The Physiology of Reproduction [1, 1 ed.] 0881672815, 9780881672817

Table of contents :
Front Cover
Title Page
Copyright
Preface
Contents
Contributors
Foreword
The Gametes, Fertilization and Early Embryogenesis
1: Sex Determination and Differentiation (George & Wilson)
2: The Spermatozoon (Eddy)
3: The Mammalian Ovum (Wassarman)
4: Gamete and Zygote Transport (Harper)
5: Mammalian Fertilization (Yanagimachi)
6: Early Mammalian Embryogenesis (Pedersen)
7: Biology of Implantation (Weitlauf)
The Reproductive Systems: The Female
8: Embryology of Mammalian Gonads and Ducts (Byskov & Hoyer)
9: The Primate Oviduct and Endometrium (Brenner & Maslar)
10: Follicular Steroidogenesis and Its Control (Gore-Langton & Armstrong)
11: Follicular Selection and Its Control (Greenwald & Terranova)
12: Mechanism of Mammalian Ovulation (Lipner)
13: The Corpus Luteum and Its Control (Niswender & Nett)
14: Local Nonsteroidal Regulators of Ovarian Function (Tsafriri)
15: Inhibin (Steinberger & Ward)
16: Relaxin (Sherwood)
17: Actions of Ovarian Steroid Hormones (Clark & Markaverich)
The Reproductive Systems: The Male
18: Perspectives in the Male Sexual Physiology of Eutherian Mammals (Williams-Ashman)
19: Anatomy, Vasculature, Innervation, and Fluids of the Male Reproductive Tract (Setchell & Brooks)
20: The Cytology of the Testis (de Kretser & Kerr)
21: The Sertoli Cell (Bardin et al.)
22: Testicular Steroid Synthesis: Organization and Regulation (Hall)
23: Efferent Ducts, Epididymis, and Vas Deferens: Structure, Functions, and Their Regulation (Robaire & Hermo)
24: Androgen Action and the Sex Accessory Tissues (Coffey)
25: Male Sexual Function: Erection, Emission, and Ejaculation (Benson)
The Pituitary and the Hypothalamus
26: Pituitary and Hypothalamus: Perspectives and Overview (Everett)
27: The Anatomy of the Hypothalamo-Hypophyseal Complex (Page)
28: Role of Classic and Peptide Neuromediators in the Neuroendocrine Regulation of LH and Prolactin (Weiner, Findell, & Kordon)
29: The Gonadotropin-Releasing Hormone (GnRH) Neuronal Systems: Immunocytochemistry (Silverman)
30: Lactotropes and Gonadotropes (Tougard & Tixier-Vidal)
31: Gonadotropins: Chemistry and Biosynthesis (Pierce)
32: Gonadotropin Secretion and Its Control (Fink)
33: Prolactin Secretion and Its Control (Neill)
Subject Index
Back Cover

Citation preview

Reproduced from Medicine and the Artist (Ars Medica) by permission of the Philadelphia Museum ofArt.

on /it iij Lx

Compliments of: W.B. Saunders

frail

Ex Etbrtjs: Peter Vasilenko,

1

1

Itteag^liBlfe

|n^

Ph.D.

Physician ministering to a patient (Hans Weiditz).

The Physiology of Reproduction

The

PHYSIOLOGY of

REPRODUCTION Volume 1

Editors-in-Chief

Ernst Knobil

Jimmy D. Neill

The H. Wayne Hightower Professor in the Medical Sciences and Director, Laboratory for Neuroendocrinology The University of Texas Health Science Center at Houston—Medical School Houston, Texas

Professor and Chairman Department of Physiology and Biophysics The University of Alabama at Birmingham Birmingham, Alabama

Associate Editors

Larry L. Ewing

Gilbert S. Greenwald

Professor, Division of Reproductive Biology Department of Population Dynamics The Johns Hopkins University School of Hygiene and Public Health

Distinguished Professor and Chairman Department of Physiology University of Kansas Medical Center Kansas City, Kansas

Baltimore, Maryland

Clement L. Markert

Donald W. Pfaff

University Research Professor Department of Animal Sciences North Carolina State University School of Agriculture and Life Sciences Raleigh, North Carolina

Professor Laboratory of Neurobiology and Behavior Rockefeller University New York, New York

Raven Press

Jp*

New York

Raven Press, 1185 Avenue of the Americas, New York, New York 10036

© 1988 by Raven Press, Ltd. All rights reserved. This book is protected by copyright. No part of it may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronical, mechanical, photocopying, or recording, or otherwise, without the prior written permission of the publisher.

Made in the United States of America

Library of Congress Cataloging-in-Publication Data

The Physiology of reproduction. Includes bibliographies and index. 1. Reproduction. 2. Mammals—Physiology. I. Knobil, Ernst. II. Neill, Jimmy D. [DNLM: 1. Reproduction. WQ 205 P5784] QP251.P525 1988 599.016 85-42819 ISBN 0-88167-281-5 The material contained in this volume was submitted as previously unpublished material, except in the instances in which credit has been given to the source from which some of the illustrative material was derived. Great care has been taken to maintain the accuracy of the information contained in the volume. However, neither Raven Press nor the editors can be held responsible for errors or for any consequences arising from the use of the information contained herein. Materials appearing in this book prepared by individuals as part of their official duties as U.S. Government employees are not covered by the abovementioned copyright.

9 8 7 6 5 4 3 2 1

Preface

This work was undertaken, after much deliberation, in an attempt to fill a need for a comprehen¬ sive, scholarly treatise on the physiology of mammalian reproduction. A major inspiration for this effort has been the volume, Sex and Internal Secretions, a factual and conceptual beacon which guided generations of reproductive biologists from the time of its first publication in 1932 to the appearance of its last edition over a quarter of a century ago. The book is divided into five major sections, and these, in turn, are loosely arrayed in two domains. The first covers the components of the reproductive system, and the second discusses reproductive processes and their physiological control. In the latter, we have included reproductive behavior in the conviction that this fundamental aspect of reproduction clearly belongs in the physi¬ ological realm and will remain a demanding challenge long after all the other mysteries in the field have been resolved. In our discussion of reproductive systems, we have been aware of the profound differences among mammals in the way some fundamental processes, such as the ovarian cycle, are controlled. We have addressed this issue, in part, by providing separate coverage of major mammalian groups where this seemed appropriate. It has been left to the reader to ascertain the similarities and differ¬ ences among them. In any case, we must ever be mindful in considering reproductive processes, from the control of ovulation to the initiation of parturition, not to extrapolate from one species to another without due reflection. It is hoped that this book will be useful to all serious students of reproductive physiology be they scientists, teachers, or physicians. The Editors

v

-

t

Contents

Foreword by Roy O. Greep

.,.

xvii

VOLUME 1 The Gametes, Fertilization and Early Embryogenesis 1.

Sex Determination and Differentiation . Fredrick W. George and Jean D. Wilson

2.

The Spermatozoon E. M. Eddy

3.

The Mammalian Ovum PaulM. Wassarman

4.

Gamete and Zygote Transport Michael J. K. Harper

5.

Mammalian Fertilization R. Yanagimachi

6.

Early Mammalian Embryogenesis Roger A. Pedersen

7.

Biology of Implantation H. M. Weitlauf

. .

3 27 69

.

103

.

135

.

187

.

231

The Reproductive Systems The Female 8.

Embryology of Mammalian Gonads and Ducts Anne Grete Byskov and Poul Erik Hfiyer

.

265

9.

The Primate Oviduct and Endometrium Robert M. Brenner and I la A. Maslar

.

303

10.

Follicular Steroidogenesis and Its Control . Robert E. Gore-Langton and David T. Armstrong

331

11.

Follicular Selection and Its Control . Gilbert S. Greenwald and Paul F. Terranova

387

12.

Mechanisms of Mammalian Ovulation Harry Lipner

447

.

vii

viii

/

Contents

13.

The Corpus Luteum and Its Control

.

489

Gordon D. Niswender and Terry M. Nett 14.

Local Nonsteroidal Regulators of Ovarian Function

. *•.

527

.

567

Alex Tsafriri 15.

Inhibin

Anna Steinberger and Darrell N. Ward 16.

Relaxin

.

585

O. David Sherwood 17.

Actions of Ovarian Steroid Hormones

.

675

James H. Clark and Barry M. Markaverich The Male

18.

Perspectives in the Male Sexual Physiology of Eutherian Mammals

.

727

H. G. Williams-Ashman 19.

Anatomy, Vasculature, Innervation, and Fluids of the Male Reproductive Tract

...

753

.

837

B. P. Setchell and D. E. Brooks 20.

The Cytology of the Testis

D. M. de Kretser and J. B. Kerr 21.

The Sertoli Cell . C. Wayne Bardin, C. Yan Cheng, Neil A. Mustow, and Glen L. Gunsalus

933

22.

Testicular Steroid Synthesis: Organization and Regulation

.

975

Efferent Ducts, Epididymis, and Vas Deferens: Structure, Functions, and Their Regulation .

999

Peter F. Hall 23.

Bernard Robaire and Louis Hermo 24.

Androgen Action and the Sex Accessory Tissues

.

1081

Donald S. Coffey 25.

Male Sexual Function: Erection, Emission, and Ejaculation

.

1121

George S. Benson

The Pituitary and the Hypothalamus 26.

Pituitary and Hypothalamus: Perspectives and Overview

.

1143

.

1161

Role of Classic and Peptide Neuromediators in the Neuroendocrine Regulation of LH and Prolactin .

1235

John W. Everett 27.

The Anatomy of the Hypothalamo-Hypophyseal Complex

Robert B. Page 28.

Richard I. Weiner, Paul R. Findell, and Claude Kordon 29.

The Gonadotropin-Releasing Hormone (GnRH) Neuronal Systems: Immunocytochemistry .

1283

Ann-Judith Silverman 30.

Lactotropes and Gonadotropes

.

Claude TougardandAndree Tixier-Vidal

1305

Contents

31.

Gonadotropins: Chemistry and Biosynthesis

.

1335

.

1349

.

1379

John G. Pierce 32.

Gonadotropin Secretion and Its Control

George Fink 33.

Prolactin Secretion and Its Control

Jimmy D. Neill Subject Index follows page 1390

VOLUME 2 Reproductive Behavior and Its Control 34.

The Physiology of Male Sexual Behavior

.

1393

Benjamin D. Sachs and Robert L. Meisel 35.

Cellular Mechanisms of Female Reproductive Behaviors

.

1487

.

1569

Donald W. Pfaff and Susan Schwartz-Giblin 36.

Maternal Behavior

Michael Numan 37.

Endocrine Basis of Communication in Reproduction and Aggression

.

1647

John C. Wingfield and Peter Marler 38.

Pheromones and Mammalian Reproduction

.

1679

John G. Vandenbergh

Reproductive Processes and Their Control 39.

Puberty in the Rat

.

1699

Sergio R. Ojeda and Henryk F. Urbanski 40.

Puberty in the Female Sheep

.

1739

.

1763

Douglas L. Foster 41.

Puberty in Primates

Tony M. Plant 42.

Rhythms in Reproduction

.

1789

Fred W. Turek and Eve Van Cauter 43.

Seasonal Regulation of Reproduction in Mammals

.

1831

F. H. Bronson 44.

The Ovarian Cycle of the Rabbit: Its Neuroendocrine Control

.

1873

.

1893

Victor D. Ramirez and C. Beyer 45.

The Ovarian Cycle of the Rat

Marc E. Freeman 46.

Neuroendocrine Control of the Ovine Estrous Cycle

Robert L. Goodman

.

1929

/

IX

x

/

Contents

47.

The Menstrual Cycle and Its Neuroendocrine Control

.

1971

. "•

1995

Ernst Knobil and Julane Hotchkiss 48.

Recognition and Maintenance of Pregnancy

Gary D. Hodgen and Joseph Itskovitz 49.

Immunological and Genetic Factors Influencing Pregnancy

.

2023

.

2043

Thomas J. Gill III 50.

Placental Transport

Frank H. Morriss, Jr. and Robert D. H. Boyd 51.

The Placenta as an Endocrine Organ: Steroids

.

2085

S. Solomon 52.

The Placenta as an Endocrine Organ: Polypeptides

.

2093

.

2145

Frank Talamantes and Linda Ogren 53.

Maternal Physiology During Gestation

James Metcalfe, Michael K. Stock, and Donald H. Barron 54.

Parturition

...

2177

J. R. G. Challis and D. M. Olson 55.

Mechanisms of Prolactin Action

.

2217

James A. Rillema, RansomeN. Etindi, John P. Ofenstein, and Steven B. Waters 56.

Lactation and Its Hormonal Control

.

2235

H. Allen Tucker 57.

The Biosynthesis and Secretion of Oxytocin and Vasopressin

.

2265

Harold Gainer, Miriam Altstein, MarkH. Whitnall, and Susan Wray 58.

Milk Ejection and Its Control

.

2283

J. B. Wakerley, G. Clarke, and A. J. S. Summerlee 59.

Suckling and the Control of Gonadotropin Secretion

.

2323

Reproductive Senescence: Phenomena and Mechanisms in Mammals and Selected Vertebrates .

2351

Alan S. McNeilly 60.

Frederick S. vom Saal and Caleb E. Finch Subject Index follows page 2414

Contributors

Laboratory of Neurochemistry and Neuroimmunology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, Maryland 20892

Miriam Altstein

Medical Research Council Group in Reproductive Biology, Department of Physiology and Obstetrics and Gynaecology, The University of Western Ontario, London, Ontario, Canada N6A 5A5

David T. Armstrong

C. Wayne Bardin

The Population Council, New York, New York 10021

Department of Obstetrics and Gynecology, University of Florida College of Medicine, Gainesville, Florida 32601

Donald H. Barron

Professor of Surgery (Urology), The University of Texas Health Science Center, Houston, Texas 77225

George S. Benson

Centro de Investigacion en Reproduccion Animal, CINVECTAVUAT Panothla, Tlaxacala, Mexico

C. Beyer

Action Research Placental and Perinatal Unit, Department of Child Health, University of Manchester, St. Mary’s Hospital, Manchester Ml 3 OJH, United Kingdom

Robert D. H. Boyd

Division of Reproductive Biology and Behavior, Oregon Regional Primate Research Center, Beaverton, Oregon 97006

Robert M. Brenner

Institute of Reproductive Biology, Department of Zoology, University of Texas, Austin, Texas 78712

F. H. Bronson

D. E. Brooks

(Deceased) Department of Animal Sciences, Waite Agricultural Research Institute,

University of Adelaide, Adelaide, South Australia Laboratory of Reproductive Biology II, Rigshospitalet, University of Copenhagen, DK-2100 Copenhagen, Denmark

Anne Grete Byskov

Departments of Obstetrics and Gynaecology and Physiology, The Research Institute, St. Joseph’s Health Centre, University of Western Ontario, London, Ontario, Canada N6A 4V2

J. R. G. Challis

C. Yan Cheng

The Population Council, New York, New York 10021

James H. Clark

Department of Cell Biology, Baylor College ofMedicine, Houston, Texas 77030

Department of Anatomy, Medical School, University of Bristol, Bristol, BS8 1TD, United Kingdom

G. Clarke

Department of Urology, The Johns Hopkins University, School of Medicine, Baltimore, Maryland 21205

Donald S. Coffey

D. M. de Kretser

Department of Anatomy, Monash University, Melbourne, Victoria 3168,

Australia Gamete Biology Section, Laboratory of Reproductive and Developmental Toxicology, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, North Carolina 27709

E. M. Eddy

xi

xii

/ Contributors Ransome N. Etindi Department of Physiology, Wayne State University, School of Medicine, Detroit, Michigan 48201 John W. Everett Department ofAnatomy, Duke University School of Medicine, Durham, North Carolina 27710 “. Caleb E. Finch Andrus Gerontology Center, Department of Biological Sciences, University of Southern California, Los Angeles, California 90081 Paul R. Findell Department of Obstetrics, Gynecology, and Reproductive Sciences, University of California at San Francisco, School of Medicine, San Francisco, California 94143 George Fink MRC Brain Metabolism Unit, University Department of Pharmacology, Edinburgh EH8 9JZ, United Kingdom Douglas L. Foster Associate Professor of Obstetrics and Gynecology, and Biological Sciences and Associate Research Scientist of Developmental and Reproductive Biology, The University of Michigan, Ann Arbor, Michigan 48106 Marc E. Freeman Florida 32306

Department of Biological Science, Florida State University, Tallahassee,

Harold Gainer Laboratory ofNeurochemistry and Neuroimmunology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, Maryland 20892 Fredrick W. George Department of Cell Biology and Anatomy, University of Texas Health Science Center at Dallas, Southwestern Medical School, Dallas, Texas 75235 Thomas J. Gill III Department of Pathology, University of Pittsburgh, School of Medicine, Pittsburgh, Pennsylvania 15261 Robert L. Goodman Virginia 26506

Department of Physiology, West Virginia University, Morgantown, West

Robert E. Gore-Langton Medical Research Council Group in Reproductive Biology, Department of Physiology and Obstetrics and Gynaecology, The University of Western Ontario, London, Ontario, Canada N6A 5A5 Gilbert S. Greenwald City, Kansas 66103

Department of Physiology, University of Kansas Medical Center, Kansas

Roy O. Greep Director of the Laboratory of Human Reproduction and Reproductive Biology, Professor of Anatomy, Emeritus, Harvard Medical School, Boston, Massachusetts 02115 Glen L. Gunsalus

The Population Council, New York, New York 10021

Peter F. Hall Department of Medicine, Prince of Wales Hospital, Randwick, New South Wales 2301, Australia Michael J. K. Harper Departments of Obstetrics and Gynecology and Physiology, The University of Texas Health Science Center at San Antonio, San Antonio, Texas 78284 Louis Hermo Centre for the Study of Reproduction, Department of Anatomy, McGill University, Montreal, Quebec, Canada H3G 1Y6 Gary D. Hodgen The Jones Institute for Reproductive Medicine and Department of Obstetrics and Gynecology, Eastern Virginia Medical School, Norfolk, Virginia 23507 Julane Hotchkiss Laboratory for Neuroendocrinology, The University of Texas Health Science Center at Houston, The University of Texas Medical School, Houston, Texas 77225 Poul Erik Hdyer Institute of Medical Anatomy A, The Panum Institute, University of Copenhagen, DK-2200 Copenhagen, Denmark Joseph Itskovitz Department of Obstetrics and Gynecology, Technion, Israel Institute of Technol¬ ogy, Faculty of Medicine, Rambam Medical Center, Haifa, Israel 31096

Contributors

J. B. Kerr

Department of Anatomy, Monash University, Melbourne, Victoria 3168, Australia

Laboratory for Neuroendocrinology, The University of Texas Health Science Center at Houston, The University of Texas Medical School, Houston, Texas 77225

Ernst Knobil

Unite de Neuroendocrinologie, INSERM U159, 75014 Paris, France

Claude Kordon Harry Lipner

Department of Biological Sciences, Florida State University, Tallahassee, Florida

32306 Barry M. Markaverich

Department of Cell Biology, Baylor College ofMedicine, Houston, Texas

77030 Peter Marler

The Rockefeller University, Field Research Center, Millbrook, New York 12545

Division of Reproductive Biology and Behavior, Oregon Regional Primate Research Center, Beaverton, Oregon 97006

Ila A. Maslar

MRC Reproductive Biology Unit, University of Edinburgh, Centre for Repro¬ ductive Biology, Edinburgh EH3 9EW, United Kingdom

Alan S. McNeilly

Department of Psychological Sciences, Purdue University, West Lafayette,

Robert L. Meisel

Indiana 47907 Heart Research Laboratory, Department of Medicine, Oregon Health Sciences University, Portland, Oregon 97201

James Metcalfe

Frank H. Morriss, Jr.

Department of Pediatrics, University of Iowa Hospitals and Clinics, Iowa

City, Iowa 52242 Neil A. Mustow

The Population Council, New York, New York 10021

Department of Physiology and Biophysics, University of Alabama at Birmingham, Birmingham, Alabama 35294

Jimmy D. Neill

Department of Physiology and Biophysics, Colorado State University, Fort Collins, Colorado 80532

Terry M. Nett

Department of Physiology and Biophysics, Colorado State University, Fort Collins, Colorado 80532

Gordon D. Niswender

Michael Numan

Department of Psychology, Boston College, Chestnut Hill, Massachusetts 02167

Department of Physiology, Wayne State University, School of Medicine, Detroit, Michigan 48201

John P. Ofenstein

Biology Board of Studies, Thimann Laboratories, University of California, Santa Cruz, California 95064

Linda Ogren

Department of Physiology, The University of Texas Health Science Center at Dallas, Dallas, Texas 75235-9040

Sergio R. Ojeda

Departments of Paediatrics and Physiology, The Research Institute, St. Joseph’s Health Centre, University of Western Ontario, London, Ontario, Canada N6A 4V2

D. M. Olson

Professor of Neurosurgery and Anatomy, Chairman, Neuroscience Program, Milton S. Hershey Medical Center, The Pennsylvania State University, Hershey, Pennsylvania 17033

Robert B. Page

Laboratory of Radiobiology and Environmental Health (LR-102), University of California at San Francisco, San Francisco, California 94143

Roger A. Pedersen

Laboratory of Neurobiology and Behavior, The Rockefeller University, New York, New York 10021

Donald W. Pfaff

Department of Biological Chemistry, UCLA School of Medicine, Los Angeles, California 90024

John G. Pierce

/

xiii

xiv

/

Contributors

Tony M. Plant 15261

Department of Physiology, University of Pittsburgh, Pittsburgh, Pennsylvania

Victor D. Ramirez Department of Physiology and Biophysics, University of Illinois at UrbanaChampaign, Urbana, Illinois 61801 “» James A. Rillema Department of Physiology, Wayne State University, School of Medicine, Detroit, Michigan 48201 Bernard Robaire Centre for the Study ofReproduction, Departments ofPharmacology and Thera¬ peutics, and Obstetrics and Gynecology, McGill University, Montreal, Quebec, Canada H3G 1Y6 Benjamin D. Sachs 06268

Department of Psychology, University of Connecticut, Storrs, Connecticut

Susan Schwartz-Giblin Laboratory of Neurobiology and Behavior, The Rockefeller University, New York, New York 10021 B. P. Setchell Department of Animal Sciences, Waite Agricultural Research Institute, University of Adelaide, Adelaide, South Australia O. David Sherwood University of Illinois, College ofMedicine and Department of Physiology and Biophysics, University of Illinois, Urbana, Illinois 61801 Ann-Judith Silverman Department of Anatomy and Cell Biology, Columbia University, College of Physicians and Surgeons, New York, New York 10032 S. Solomon Departments of Medicine and Obstetrics and Gynecology, McGill University and the Royal Victoria Hospital, Montreal, Quebec H3A 1A1, Canada Anna Steinberger Department of Obstetrics, Gynecology and Reproductive Sciences, The Univer¬ sity of Texas Medical School at Houston, Houston, Texas 77030 Michael K. Stock Heart Research Laboratory, Department of Medicine, Oregon Health Sciences University, Portland, Oregon 97201 A. J. S. Summerlee Department of Anatomy, Medical School, University of Bristol, Bristol, BS8 1TD, United Kingdom Frank Talamantes Biology Board of Studies, Thimann Laboratories, University of California, Santa Cruz, California 95064 Paul F. Terranova Kansas 66103

Department of Physiology, University of Kansas Medical Center, Kansas City,

Andree Tixier-Vidal Groupe de Neuroendocrinologie Cellulaire et Moleculaire, College de France, ll, Place Marcelin Berthelot, 75231 Paris Cedex 05, France Claude Tougard Groupe de Neuroendocrinologie Cellulaire et Moleculaire, College de France, ll, Place Mar celin Berthelot, 75231 Paris Cedex 05, France Alex Tsafriri Department of Hormone Research, The Weizmann Institute of Science, Rehovot 76100, Israel H. Allen Tucker Department of Animal Science, Michigan State University, East Lansing, Michigan 48824 Fred W. Turek Department of Neurobiology and Physiology, Northwestern University, Evanston, Illinois 60201 Henryk F. Urbanski Department of Physiology, The University of Texas Health Science Center at Dallas, Dallas, Texas 75235-9040 Eve Van Cauter Institute of Interdisciplinary Research, Free University of Brussels, B-1070 Brussels, Belgium and Department of Medicine, University of Chicago, Chicago, Illinois 60637

Contributors

John G. Vandenbergh Carolina 27695

Department of Zoology, North Carolina State University, Raleigh, North

Frederick S. vom Saal Division of Biological Sciences, John M. Dalton Research Center, Univer¬ sity of Missouri, Columbia, Missouri 65211 J. B. Wakerley Department of Anatomy, Medical School, University of Bristol, Bristol, BS8 1TD, United Kingdom Darrell N. Ward Department of Biochemistry and Molecular Biology, The University of Texas, M.D. Anderson Hospital and Tumor Institute, Houston, Texas 77030 Paul M. Wassarman Department of Cell Biology, Roche Institute of Molecular Biology, Roche Research Center, Nutley, New Jersey 07110 Steven B. Waters Department of Physiology, Wayne State University, School of Medicine, Detroit, Michigan 48201 Richard I. Weiner Department of Obstetrics, Gynecology, and Reproductive Sciences, University of California, San Francisco, School of Medicine, San Francisco, California 94143 H. M. Weitlauf Department of Cell Biology and Anatomy, Texas Tech University Health Sciences Center, Lubbock, Texas 79430 Mark H. Whitnall Laboratory of Neurochemistry and Neuroimmunology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, Maryland 20892 H. G. Williams-Ashman Ben May Institute, and Departments of Biochemistry and Molecular Biology and Pharmacological and Physiological Sciences, University of Chicago, Chicago, Illinois 60637 Jean D. Wilson Department of Internal Medicine, University of Texas Health Science Center at Dallas, Southwestern Medical School, Dallas, Texas 75235 John C. Wingfield 12545

The Rockefeller University, Field Research Center, Millbrook, New York

Susan Wray Laboratory of Neurochemistry and Neuroimmunology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, Maryland 20892 R. Yanagimachi Department ofAnatomy and Reproductive Biology, University of Hawaii School of Medicine, Honolulu, Hawaii 96822

XV

§|pi

■

i

■■■

'3^,.

;

.

*•

■

Foreword

I am pleased and honored to have been asked to prepare the Foreword to this volume of work depicting the progress in research on the physiology of reproduction as well as the resulting gains in understanding made over the past few years. The expertise that is represented by the numerous contributors to this work is so impressive that I am humbled even to contemplate adding anything of note. It is only by virtue of the perspective garnered from my vantage point of having personally witnessed a very large segment of twentieth century research on reproduction that I am emboldened to reflect on the byways and the trail blazings that have brought this field to its present proud state of enlightenment with regard to the long sought-after means of controlling the procreative process in humankind. Clearly, there are many important and knotty problems yet to be resolved, but the pace of progress over the past several years has quickened to the extent that one is left in expectant wonderment as to where and when the next revolutionizing development will occur. The experimental method of studying reproduction was initiated in 1849 with Berthold’s discov¬ ery of a blood-borne activity that came from the testis and stimulated growth of distant organs such as the comb and wattles. In so doing he utilized one of the most fundamental means of demonstrat¬ ing the function of an endocrine organ, namely, surgical removal to determine what deficiencies follow, coupled with implantation or transplantation to ascertain whether the deficiencies were repaired. At that time it was not possible to take the next step, namely, preparation of an active extract of the testes, because nothing was known about the chemical nature of the bioactivity. Forty years later, Brown-Sequard claimed to have prepared an active extract of dog testes; however, as is well known, his enthusiastic claims for restoration of his own sexual activity at an advanced age were not substantiated. Actually, these simple means of studying reproductive physiology persisted well into the twentieth century, including the studies of such pioneering stalwarts as Marshall, Heape, Prenant, Bouin, Ancel, Loeb, Cushing, and Aschner. Observations otherwise were limited to cyclic and seasonal changes in sexual behavior among common laboratory and small domestic animals. This type of eyeball research remained in vogue through the early 1920s and overlapped the extension of visualization to the microscopic level. The latter revealed, for the first time, the precise timing of events in the ovarian cycle through microscopically observable cellular changes in the vaginal fluid. My point in mentioning these early studies is to emphasize that although the tools and techniques were inordinately primitive by present standards, the results established a firm base of knowledge on which to build. The study of cyclic changes in the vaginal smear in rats and the finding of estrogenic activity in follicular fluid during the early 1920s led to an explosion of interest in the study of reproduction. The field was fortunate in attracting to its ranks a small band of exceedingly able biologists and biochemists who were to become authors of the classic compendium, the first edition, in 1932, of Sex and Internal Secretions, overwhelmingly devoted to reproductive endocrinology. It was this landmark of progress that finally gave propriety to the study of reproduction and put it on a par with the study of other major bodily systems. Incredible as it may seem, it was only a decade earlier that a distinguished panel of the National Research Council had declared that sex research was not a fitting topic for scientific study. Lest our pride in today’s spectacular pace of progress unduly bedazzle the mind, it should not be overlooked that the developments recorded in the 10-year span from 1926 to 1936 may never be equaled. Among those monumental achievements, all of the native sex steroid hormones were brought to light, their structures were determined, their functions were defined, and they were made available in pure form for research and therapy. Similarly, all of the pituitary, placental, and urinary tropic hormones were identified, and their functions were defined. Like today’s competition for priority rights, publicity, and potential financial gain, these earlier periods also were times of intense

xvil

xviii

/

Foreword

rivalries, but rarely with prospects for financial rewards. It would be difficult to overstate the boost that was given to basic and clinical research in reproduction as a result of having available estradiol17/3, testosterone, and progesterone in pure form and of known potency. The replacement of homemade extracts and such elastic entities as rat units, mouse units, capon units, etc., with micrograms of pure hormone was revolutionizing and allowed the study of reproduction to be on a quantitative basis. Prior to World War II the thrust of research on reproduction dealt predominantly with the steroid hormones. This was the heyday of steroid biochemistry. After World War II the emphasis shifted to the protein and peptide hormones, where it still remains strong. This prolonged and difficult effort yielded many biochemical triumphs. Most notable among these were the isolation of the pituitary, placental, and urinary gonadotropins as well as the determination of their primary struc¬ ture as glycoproteins comprised of two dissimilar and covalently bonded subunits, the isolation and synthesis of the gonadotropin-releasing hormone (GnRH) of hypothalamic origin, and the isolation and structural characterization of relaxin. The availability of pure protein and polypeptide hormones made possible the production of hor¬ mone-specific antibodies as well as the application of immunological techniques to the study of reproduction. An outcome of great consequence was the development of radioimmunoassay as a new means of measuring all of the hormones relating to reproduction. The sensitivity of this new technique was so great that it made possible, for the first time, the measurement of all these hor¬ mones in the body fluids. It had the further distinct advantage of requiring such a small amount of fluid that the monitoring of blood levels of the hormones of reproduction could be done throughout an estrous or menstrual cycle by close serial sampling. This revealed still another and most unex¬ pected finding, the pulsatile pattern of secretion. Identifying the homeostatic mechanism(s) responsible for maintaining a steady state in various physiologic systems of the body has been fraught with many challenging problems, but these pale in comparison with the difficulties encountered in trying to elucidate the mechanisms maintaining a constantly changing system, a characteristic of the reproductive system of female mammals. The earliest piece of evidence suggested the existence of a “push-pull” mechanism that later came to be known as negative feedback. It was based on the demonstration that an estrogenic extract adminis¬ tered to immature rats would maintain the ovaries in an infantile state. This was quickly followed by conclusive evidence that estrogen acted to inhibit pituitary follicle-stimulating hormone (FSH) stimulation of follicular growth and maturation; however, the effect on luteinizing hormone (LH), ovulation, and luteinization remained unsettled. Gaps continued to exist in all proposed explana¬ tions of reproductive cycles. None of these explanations took account of the influence of photoperi¬ odicity on seasonal breeders, nor did they account for the stimulus of mating in nonspontaneous ovulators. Following the discovery of the hypothalamic control of pituitary function, estrogen was shown to exert its action on both the pituitary and the hypothalamus; however, the problem of accounting for cyclicity remained. Adding to the complexity, radioimmunoassay revealed an unex¬ pectedly high level of blood estrogen just prior to ovulation—an event not in keeping with the negative feedback concept. Finally, after many years of searching for a way out of this frustrating situation, a glimmer of light appeared at the end of this long dark tunnel—light that soon turned to brilliance. In 1969, Goding and associates found that the administration of large doses of estrogen to ewes at the time of estrus did not block, but instead entrained, ovulation. Shortly thereafter, in more elaborate exam¬ ination of the relationship of blood estrogen levels and ovulation in rhesus monkeys in Knobil’s laboratory, it was revealed that elevated estrogen levels preceded and appeared to trigger ovulation. On further examination, Knobil and colleagues found that when blood estrogen reached a critical level the feedback mechanism switched from a negative to a positive, or stimulative, action. This utterly new finding greatly advanced our understanding of the endocrine mechanism governing reproductive cycles. There still remain, however, some uncertainties: Why does the switch in feed¬ back action occur; to what extent and at what stage of the cycle does estrogen act at the level of the pituitary or the hypothalamus, or both; and lastly, what role, if any, do the ovarian peptides, espe¬ cially inhibin, play in controlling reproductive cycles? The progress of research on reproduction has been chronicled in numerous review articles by individual authors. Many have appeared in Recent Progress in Hormone Research, Volumes 1 to 42. Other major sources include the multiple editions of such titles as: Marshall’s Physiology of Reproduction, now being produced in its fourth edition; Sex and Internal Secretions, whose third

Foreword

and last edition was issued in 1981; two volumes on the Female Reproductive System (1973), and one on the Male Reproductive System (1975) in Section 7 of the Handbook ofPhysiology, published by the American Physiological Society; and four serial volumes on reproductive physiology in the International Review of Physiology, the last one being issued in 1983. The present volume will provide comprehensive coverage and meet the current needs of the held of reproductive physiology, a held that is rapidly gathering momentum from the application of new and highly sophisticated tools and techniques. In viewing the vast literature dealing with research on the male and female reproductive systems and considering the rate at which it is accumulating, one might ask whether this staggering prolifera¬ tion of books and articles is essential to progress; the answer is an emphatic “yes.” The yardstick by which progress is measured in this or any other held is not in number of articles published or amount of hnancial support but in improved understanding. Such gains are generally marked by sharp peaks at indeterminate intervals, separated by avalanches of incremental gains, as recorded in an ever-growing list of journals. The point to remember is that without this persistent chipping away at a major problem there would be no solutions and no quantum leaps forward. In research very little comes from out of the blue. Part of the driving force in research is its adventuresome nature and the ever-present possibility that one’s efforts will pay off in an important manner. It may not be entirely fair, but in research (as in most human activities), the spoils go to the victor in the form of kudos, prizes, awards, public attention, and, increasingly in the present technological age, monetary gains—sometimes of great magnitude. What effect this latter may have, if any, on the long-cherished sanctity of science has not been determined, but it has become a matter of concern. This volume bears the title The Physiology of Reproduction. Physiology, by traditional consensus, is that branch of science which studies the functions of a living organism or any of its parts and includes the basic underlying processes. It will be understood that most of the studies reviewed here will be based more on holistic research than on research at the submicroscopic or molecular level. It is unfortunate that the excitement generated by recent fantastic advances in molecular biology and development has tended to downgrade the value of whole-animal research, and physiology in particular is sometimes looked upon as passe. Actually, the two categories of research are comple¬ mentary, and both are essential for maximal advancement of knowledge. Whole-animal research cannot become outdated because it is the quintessence of biological relevance and the means by which molecular findings must ultimately be evaluated. In this same vein, no one immersed in reproductive endocrinology can be unaware of the current tendency to regard research at the molecular level as representative of exceptional scientific talent. This is a common consequence of the opening of a new arena of investigation. I recall an incident that happened at a scientific meeting back in the 1930s. The first three papers in a session chaired by an eminent embryologist were on endocrine topics—mine was the third. That being ended, the chairman took pains to assure the audience that the meeting could now turn to considerations of more fundamental nature. One of the other three papers was given by Herbert M. Evans, who bristled noticeably but held his fire. There was also an earlier period when one either worked on steroid biochemistry or something of lesser appeal like biology. Anyone who remembers the 1950s will recall a flash-in-the pan ignited by cybernetics, a study of automatic control systems both neural and physical. The gurus of cybernetics captured the attention of the press and of audiences through¬ out the land, but eventually this obsession suffered the fate of other passing preoccupations. My own observation is that the closer one approaches the molecular level of research, the more one becomes dependent on highly sophisticated instrumentation to make the observations and to read out results that are often quite free of extraneous variables. Toward the obverse situation, one’s dependence on an extensive background of experience and physiological insight increases as does the unavoidable complex of in vivo variables that must be taken into account. In either case we have today the availability of far more diverse approaches to a given problem in any field of biomed¬ ical research than has ever existed before. In Berthold’s day there was only one experimental method available; today’s number is untold but is probably in the hundreds, perhaps thousands. That is an exceedingly promising situation and one to which investigators of all persuasions must adjust. Open minds will experience exhilaration over substantive achievements at any point on this observational spectrum. One of the major factors influencing research on reproduction has been the availability of funds or lack thereof. Prior to the institution of federal funding (i.e., prior to the middle of the twentieth century), reproductive research was sparsely supported by university departmental funds, industry,

xix

xx

/

Foreword

small grants from the Committee for Research in Problems of Sex within the National Research Council, and some aid from the Rockefeller Foundation. The National Institutes of Health were slow in providing significant support of research on reproduction because of restrictions on the support of work related in any way to birth control. This occurred despite the simultaneous postwar baby boom. What kept research afloat during this critical period was major support by the Ford Foundation plus lesser contributions by other major foundations. It was not until the establishment in 1968 of the Center for Population Research in the NICHD that major governmental funding in this area became available, but the boost was short-lived. As a result of the imposition of fiscal restraints in the early 1970s, federal support dwindled and has remained at a minimal level ever since. Support from all sources is woefully incommensurate with the distressing expansion of the human population and the need for safe, effective, economical, and readily available means of limiting human fertility. The physiology of reproduction is predominantly under hormonal control. The first essential step in studying reproduction was identification of the hormones involved and the functions they serve. This having been accomplished, efforts turned to a detailed analysis as to how hormones act within the body. During the past decade there has been a rising tide of interest in the binding of steroid, protein, or peptide hormones to receptors on specific target cells. Much effort is currently being directed toward the isolation and chemical characterization of these receptors. They are known to be composed of a protein or proteins, and sorffe information has already been gained as to their partial or provisional structure. This, however, is only a preliminary step in the complex process whereby hormone action results in an end response such as growth, secretion of a target cell hormone, or altered behavior. The curtain has already been raised on the climactic and final chapter of the story on how hormones act. This involves linkage of the hormone-receptor complex with the nuclear genetic apparatus leading through a now well-defined series of processes to the manifestation of a physiological response in the living organism. Genes that bring about the expres¬ sion of certain hormonal signals are being isolated, modified, transferred between species, and also inserted into bacteria where they direct the biosynthesis of specific hormones in large quantity. Thus genes are being manipulated in ways that raise the potential of altering the reproductive process. It is largely as a result of developments in endocrinology at the molecular level that bewildering possibilities loom on the horizons of reproductive research—they are within reach; they are science, not fiction; and they stagger the imagination. Roy O. Greep

The Physiology of Reproduction

The Gametes, Fertilization and Early Embryogenesis

The Physiology of Reproduction, edited by E. Knobil and J. Neill et at. Raven Press, Ltd., New York © 1988.

CHAPTER 1

Sex Determination and Differentiation Fredrick W. George and Jean D. Wilson 11 • Possible Role of Steroid Hormones in Gonadal Dif¬

Chromosomal Sex, 3

ferentiation, 11

The Y Chromosome, 4 • Pairing of the X and Y Chro¬ mosomes, 4 • Testis-Determining Genes, 5 • OvaryDetermining Genes, 6 • X-Chromosome Inactivation During Spermatogenesis, 6 • Development of a Testis in the Absence of a Y Chromosome, 6

Phenotypic Sex, 11 Urogenital Tract Development During the Indifferent Phase, 11 • Male Development, 12 • Female Development, 13 • Breast Development in Both Sexes, 7 J • Endocrine Control of Male Phenotypic Differentiation, 14

Gonadal Sex, 8

Do Hormones Play a Role in Female Phenotypic Devel¬ opment?, 18 Summary, 20 References, 21

Embryonic Development, 8 • Histological Differentia¬ tion of the Fetal Gonad, 8 • The Role of the Germ Cells in Gonadal Development, 8 • Endocrine Differentiation of the Testis, 9 • Endocrine Differentiation of the Ovary,

responsible for sexual differentiation has, however, re¬ mained largely enigmatic. The pioneering work in this field, performed in Drosophila, suggested that sex is determined by the number of X chromosomes. Flies with a single X chromosome (XY, XO) are male, whereas the presence of two or more X chromosomes (XX, XXX, or XXY) confers a female phenotype (reviewed in ref. 248). The mammalian Y chromosome was also originally believed to be a null chromosome without genetic information, with the possible exception of some factor that promoted fertility in males (227). However, with the development of techniques for the karyotyping of the mammalian chromosome in the early 1960s, it became apparent that in humans (and other mam¬ malian species) the Y chromosome specifies the develop¬ ment of the testis. That is, no matter how many X chro¬ mosomes are present, a testis will develop as long as a Y chromosome is present (as in XXY, XXYY, XXXY, XXXXY, etc.) (47). The phenotype of the 45,X human is female, although ovarian development is incomplete, im¬ plying that X chromosomes do participate in human ovarian differentiation. Nevertheless, the presence of a second X chromosome (as in XXY or XXSxr) does not appear to affect sexual differentiation at the gonadal level; e.g., either a Y chromosome or a critical fragment of a Y chromosome that contains the testis-determining genes is capable of inducing testicular development, no matter how many X chromo¬ somes are present. However, if two X chromosomes are present, spermatogenesis is impaired (except in XX true

Sexual differentiation is a sequential process beginning with the establishment of chromosomal sex at the time of fertilization, followed by the development of gonadal sex, and culminating in the formation of the sexual phenotypes (Fig. 1). Each step in this process is dependent on the preceding one, and, under normal circumstances, chro¬ mosomal sex agrees with the phenotypic sex. Occasionally, however, chromosomal sex and phenotypic sex do not agree, or the sexual phenotype may be ambiguous. Abnormalities of sexual development are usually not life threatening and occur at many levels. They encompass clinical conse¬ quences that range from defects in the terminal phases of male development (cryptorchidism and microphallus) to more fundamental abnormalities that result in conditions of in¬ tersex. In many cases, disorders of sexual differentiation are inherited as single gene mutations, and the analysis of these disorders has been especially informative in defining the molecular and genetic determinants involved in the nor¬ mal process of sexual development. Although the principal focus of this chapter is on human sexual determination, many aspects of the process have been best studied in other species.

CHROMOSOMAL SEX The chromosomal basis for sex determination was estab¬ lished between 1910 and 1916, largely by T. H. Morgan and his colleagues. The nature of the genetic information

3

4

/ Chapter 1 CHROMOSOMAL SEX

{ I

GONADAL SEX Normal Y

PHENOTYPIC SEX

FIG. 1. The Jost paradigm.

Y variants compatible with male development

hermaphroditism). The most likely explanation for the in¬ fertility in the XXY state is that inactivation of the X chro¬ mosome in the formation of the XY body (see below) during normal spermatogenesis is essential for fertility. In other words, some X-coded gene product must prevent male germ development (172). This concept is in keeping with the fact that some balanced translocations of X chromosomes to autosomes in mice (76) and in humans (163) are associated both with male infertility and with incomplete inactivation of X chromosomal genes. Furthermore, if the situation in the creeping vole (193), in which the X chromosome is eliminated from the testicular germ line, is applicable to other species, then it can be assumed that function of an X chromosome is not essential for either testicular differen¬ tiation or spermatogenesis.

The Y Chromosome The Y chromosome is the third smallest human chro¬ mosome, on average just larger than chromosomes 20 and 21 (248). The short arm is invariable in size whereas, among normal men, the long arm can vary considerably in length. The distal portion of the long arm exhibits brilliant fluo¬ rescence after quinacrine staining, with two or three separate fluorescent bands being visible in interphase nuclei (Fig. 2). The Y chromosomes of all species, like many other chro¬ mosomes, contain satellites that are only visible under cer¬ tain staining conditions. In the human Y chromosome, sat¬ ellite DNAs are present both within a distal heterochromatin segment and in the centromeric region. The satellite DNA on the human Y chromosome, whose function is unknown, does not cross-hybridize with Y-chromosome-associated satellite DNA of other species (248).

i(Yq)

Y variant compatible with female development

FIG.-2. Diagrams of normal and variant Y chromosomes. Cross-hatching indicates quinacrine-bright region of the long arm (Yq). Three abnormal types are depicted: i(Yp), iso¬ chrome of the short arm; i(Yq), isochrome of the long arm; Yq~, fragment arising as a result of loss of the distal long arm. division, only two types of sperm are produced, those con¬ taining X and those containing Y chromosomes (227). In some instances the pairing has been described as being “side by side”; in other instances, “end to end.” In the hamster, where this has been studied extensively, there is considerable variability among strains (225). In any event, the net effect is the formation, during spermatogenesis, of the XY body (i.e., the condensed chromatin of the fused X and Y chromosomes), which is identifiable between zygo¬ tene and mid-pachytene (207,225).

X

A

Y/ , i / i_/

Pairing of the X and Y Chromosomes The two X chromosomes in females pair at the centromere and segregate during the first meiotic division of oogenesis by a mechanism analogous to the pairing of the autosomes. The X and Y chromosomes, like homologous autosomes, must also pair during meiosis to ensure that they segregate properly. However, the pairing of X and Y does not occur at the centromere but apparently occurs at a small region of homology between the short arm of the X and the short arm of the Y (30) (Fig. 3). In this way, they duplicate and partition evenly on the spindle. Thus at the second meiotic

Td(Y)

Pairing I Region I

Td(Y)

XY Male

YYsxr Male

FIG. 3. Pairing of the X and Y chromosomes, and pseudoautosomal inheritance of the testis-determining region of the Y chromosome [Td(Y)j.

Sex Determination and Differentiation / Testis-Determining Genes For the reasons mentioned above, the primary gene (or genes) that controls testicular differentiation is thought to be located on the Y chromosome. Analyses of structural abnormalities of the Y chromosome in humans have sug¬ gested that the short arm of the Y carries the responsible genes. Loss of the fluorescent segment of the long arm of the Y (Yq~) (Fig. 2) or the formation of a Y “ring” chro¬ mosome (which requires loss of the distal-most segments of both the long and the short arms) does not interfere with the formation of a normal testis (47). An abnormal Y chro¬ mosome consisting of a duplicated short arm [i(Yp)] also has no effect on male development. Flowever, several ex¬ amples of isochromosomes for the long arm of the Y [i(Yq)] (Fig. 2) have been observed to result in failure of testicular development, with subsequent formation of a female phe¬ notype (18,102,129). It is therefore likely that the testis¬ determining gene(s) is located on the short arm of the Y near the centromere (243). Additional gene(s) on the short or long arms may be essential for normal spermatogenesis (64,100). Homologous single-copy regions of the DNA of the hu¬ man X and Y chromosomes have been identified with the use of restriction endonuclease techniques. These fragments appear to be unique to the human (e.g., different than the homologous regions in other species), and hence these re¬ gions, like the major repeat segments, are not thought to play a role in sexual differentiation (195). More importantly, some of the homologous regions appear to be on the tips of the short arms of the X and Y chromosomes (100) and are presumed to be the segments responsible for pairing of the two chromosomes during meiosis rather than to contain genes that specify gonadal development (Fig. 3). These pairing sequences are highly polymorphic; indeed almost every family appears to have unique segments (41). About 70% of the DNA of the human Y chromosome consists of repeated sequences of DNA located on the long arm (41). These sequences are related, in an evolutionary sense, to sequences on the human X. Hence no detectable phenotypic effect results when the long arm of the Y be¬ comes translocated onto autosomes. These long repetitive fragments of DNA of the Y chromosome in Drosophila, the mouse, and the human are composed of two repeating base quadruplets (GATA and GACA). In the human these repetitive fragments are believed to be responsible for flu¬ orescence of the distal long arm of the Y and for much of the variability in length among normal Y chromosomes. Sex-specific DNA was originally described in the W chro¬ mosomes of snakes and is now known to be conserved in many vertebrate species (69,132). In the mouse these se¬ quences are concentrated on the short arm of the Y chro¬ mosome (e.g., near the site thought to contain the deter¬ minants of testicular differentiation), and these regions of DNA are transcribed into a male-specific RNA of 1,250 to 1,400 bases (70,223). However, there is considerable vari¬

5

ability in the location and organization of these sequences among species, and it is thus unlikely that they are involved in sex determination (64). The molecular basis by which a gene (or genes) on the Y chromosome promotes testicular differentiation is not known. A leading theory, based on a male-specific trans¬ plantation antigen, proposes that the Y chromosome spec¬ ifies, or regulates, the production of a differentiative, cellsurface antigen that mediates transformation of the sexually indifferent gonad into a testis. This male-specific trans¬ plantation antigen was identified in certain inbred strains of mice in 1955 by Eichwald and Silmser (66). Female mice rejected skin grafts from male mice of the same strain. However, in the same strains, males do not reject skin grafts from females. The graft rejection in females was attributed to a transplantation antigen present only in the males. Pre¬ sumably the only source of genetic incompatibility between males and females of this inbred strain of mouse was the Y chromosome. Therefore it was deduced that male-tofemale skin graft rejection was caused by a Y-situated his¬ tocompatibility gene. This gene is termed the histocompa¬ tibility-Y (H-Y) gene (13). In the early 1970s, male-specific antibodies were detected in the serum of female mice with male skin grafts, and serologic assays were developed for measuring the male antigen (94). Using these assays it was found that the pres¬ ence of a testis is usually associated with the presence of serologically detected male antigen. Furthermore, the wide¬ spread phylogenetic conservation of H-Y antigen suggested that the antigen has an important biological function. Based on this evidence, it was proposed that the H-Y gene is, in fact, the testis-determining gene (250). However, the H-Y antigen was originally defined on the basis of rejection of skin transplantation, e.g., a transplan¬ tation antigen, whereas the serological tests examine for the presence of an antigen on the surface of cells. It is difficult in most instances to be certain whether the two assays mea¬ sure the same property; therefore the gene that determines the H-Y plasma membrane antigen may, in fact, not be identical to the gene that specifies H-Y transplantation an¬ tigen (219). Subsequently, questions have also arisen as to whether the relationship between H-Y antigen and testicular devel¬ opment is actually one of cause and effect (42,103, 131,262,265). One of the complicating features of the thesis is that in birds, in which the female rather than the male is the heterogametic sex (e.g., ZW/ZZ species), the female is H-Y antigen positive (249,264). As pointed out by Ohno (190), it would be more straightforward to suppose that any testis inducer conserved throughout evolution would be pres¬ ent in all testis-bearing males, whether homogametic or heterogametic. The presence of H-Y antigen in female chickens is even more puzzling in view of the fact that administration of estradiol to ZZ male chicks causes a change in phenotype from H-Y to H-Y+ (249). A second problem is that some 45,X women with gonadal dysgenesis are H-

6 / Chapter 1 Y antigen positive (249), a finding that also does not fit with the original thesis. Furthermore, studies in the Sxr mouse have been interpreted as indicating that the testis¬ determining (Tdy) gene and the H-Y antigen gene are sep¬ arate (174), although it is possible that the H-Y antigen gene codes for a factor essential for spermatogenesis (31). Even if the structural gene that specifies H-Y antigen (or a reg¬ ulatory gene that controls expression of the antigen) is not the testis determinant, its location on the short arm of the Y chromosome may be so near the Tdy gene that accurate measurement of the H-Y antigen would be clinically useful, since its presence would correspond, in most instances, to the presence of the testis determinant.

Ovary-Determining Genes Eicher and Washburn (63) have reported that genotypic males that carry a Y chromosome derived from a Mus domesticus strain on the genetic background of an inbred Mus musculus strain (C57BL/6J) develop ovaries or ovotestes. Sex reversal in these animals is apparently due to interaction between an autosomal recessive gene carried by C57BL/6J and the Y chromosome of Mus domesticus. Other autosomal testis-determining loci have subsequently been identified (64). Based on these results, Eicher and Washburn have proposed that, in addition to the testis-determining gene on the Y chromosome, autosomal genes are also necessary for normal gonadal differentiation and that there are also ovary¬ determining genes. According to their hypothesis, in XY individuals the testis-determining genes are activated prior to, and inactivate, ovary-determining gene(s). XX individ¬ uals lack the initial testis-determining gene, and hence the ovary-determining genes dominate. Mutations of the testis¬ determining loci that interfere with the timely and coordi¬ nated expression of these genes may result in failure of suppression of ovarian determinants, with subsequent de¬ velopment of ovarian tissue in XY individuals. X-Chromosome Inactivation During Spermatogenesis Soon after implantation of the eutherian embryo, X chro¬ mosomes in excess of one are randomly inactivated in so¬ matic cells (81,158,160). The X-chromosome inactivation in somatic cells is associated with chemical modification of the DNA (263) and the formation of a heterochromatin body in cells of the female (9,192). The formation, during spermatogenesis, of the XY body in pachytene is associated with features typical of X-chro¬ mosome inactivation, namely, late replication of the X chromosome (and presumably late replication of the Y chromosome as well) (189) and decreased incorporation of radioactive uridine into messenger-like RNA (181). The concept that a functional X chromosome is not essential for testicular differentiation is supported by the finding that, in the creeping vole, the X chromosome is eliminated (by nondisjunction) during spermatogenesis so that the germ cell

lines in the male are OY in composition (193). This also suggests that an X chromosome is not necessary for the formation of fertile sperm. Additional evidence in support of this view has come from studies of translocations of X chromosomes to autosomes in the mouse. Some of these translocations impair spermatogenesis, presumably by im¬ pairing X inactivation in primary spermatocytes (76,77, 153,154,212). The mechanism of X-chromosome inacti¬ vation during spermatogenesis appears to be fundamentally different than the random X-chromosome inactivation that occurs in the autosomes of XX females. The latter type of X-inactivation is associated with chemical modification of the DNA (263), whereas the X-chromosome DNA from sperm is functional in in vitro transcriptional assays and is presumably not modified (241). Development of a Testis in the Absence of a -Y Chromosome Study of several disorders that result from the alteration of the number or structure of the X and Y chromosomes has provided insight into the control of sexual differentia¬ tion. Disorders in which a testis develops in the apparent absence of a Y chromosome have been particularly infor¬ mative in this regard. The XX Male The sex-reversal mutation in the mouse (Sxr), which causes XX genotypic females to develop as phenotypic (albeit in¬ fertile) males (38), was originally assumed to be inherited as an autosomally linked gene; subsequent linkage studies in several laboratories failed to substantiate this view. In 1982, Singh and Jones (222) detected a Y-specific DNA fragment on the distal end of an X chromosome in XXSxr metaphase chromosomes. This finding, coupled with the formulation of Burgoyne (29) that crossing over can occur in the pairing region of the X and Y chromosome during meiosis, provides an explanation for the apparent (pseudo) autosomal inheritance of the Sxr mutation. In sup¬ port of this interpretation, an additional fragment has been identified at the distal end of the Y chromosome in XYSxr carrier males and at the distal end of one X chromosome in XXSxr males (71). Thus, it is now possible to explain testicular development in the Sxr mouse in the absence of a Y chromosome (171). The testis-determining portion of the Y chromosome is pres¬ ent as a duplicated element on the distal pairing region of the Y chromosome in XYSxr male mice. Translocation of this duplicated testis-determining region to an X chromo¬ some during meiosis can cause testicular differentiation in XX males (Fig. 3). Nevertheless, fertility does not occur on the background of an XX chromosome composition. In this sense, this disorder is a phenocopy of the 47,XXY Klinefelter’s syndrome, involving a Y fragment rather than an intact Y chromosome (see above).

Sex Determination and Differentiation

A disorder similar to that of the Sxr mouse occurs in humans. The incidence of the 46,XX karyotype in pheno¬ typic men is approximately 1 in 20,000 to 24,000 male births (49,50). Clinical features include small, firm testes (generally less than 2 ml in volume), normal male wolffian duct derivatives, and male external genitalia. Azoospermia and hence infertility are invariably present (197,215). 46,XX men resemble Klinefelter’s syndrome subjects (209). Four theories were proposed to explain the pathogenesis of the human disorder: (i) mosaicism for a Y-containing cell line or early loss of a Y chromosome in an individual who was originally 47,XXY; (ii) an autosomal gene mutation that acts independently of a Y chromosome; (iii) deletion or inactivation of an X-chromosome gene that normally has a negative regulatory effect on testis development; or (iv) interchange of a Y-chromosome fragment with the X chro¬ mosome, analogous to the situation in the Sxr mouse (50). Most cases in the human appear to be a result of the latter mechanism because most XX males, like the XXSxr mouse, contain Y-specific DNA on the tip of the X chromosome (4,51,111,243). Additional genes encoded on the human X and Y chromosomes are also inherited in a pseudoautosomal fashion (40,211,220), and at least one of these pseudoau¬ tosomal genes, MIC2, is located sufficiently close to the testis-determining gene(s) on the short arm of the Y to be a potential marker for the testis determinant(s) (101). The human testis determinant(s) also maps to the short arm of the Y chromosome (243). True Hermaphroditism True hermaphroditism is a condition in which both an ovary and a testis, or a gonad with histologic features of both ovary and testis (ovotestis), are present (238-240). To justify the diagnosis in the human, both types of gonadal epithelium must be present. The presence of ovarian stroma without oocytes is not sufficient. True hermaphroditism is actually several different disorders (221). About two-thirds of human subjects have a 46,XX karyotype, one-tenth have a 46,XY karyotype, and the remainder are chromosomal mosaics, i.e., either 46,XX/46,XY or 45,X/46,XY chi¬ meras. Instances of true hermaphroditism associated with mosaicism are generally assumed to be clonal in origin, with the X- or XX-containing cells giving rise to ovarian cell lines and the Y-containing cell lines giving rise to tes¬ ticular elements. XY true hermaphroditism is assumed to be caused by a mutation in the testis-determining gene(s) that impairs suppression of the ovarian determinants in some cells (63-65). Hermaphroditism in the presence of an XX karyotype, like the XX male syndrome, is an apparent contradiction to the axiom that a Y chromosome is necessary for testicular differentiation. Possible explanations include (a) undetected loss of a Y chromosome after initiation of testicular devel¬ opment or undetected chromosomal mosaicism or chimerism, (b) translocation of testicular determinants from the Y

/

7

to the X chromosome or to an autosome, or (c) a single gene mutation. In all instances reported to date, 46,XX true hermaphrodites are H-Y antigen positive, including the XX form of true hermaphroditism in the dog (19,213,217). Al¬ though several family aggregates of 46,XX true herma¬ phroditism have been reported (6,39,80,156,185,210), most are sporadic in nature. The available data in the familial cases are compatible with either autosomal recessive in¬ heritance, new autosomal dominant mutations, or translo¬ cation of a fragment of a Y chromosome in a paternal line. In addition, two families have been reported in which one affected member was a 46,XX male and the other was a 46,XX true hermaphrodite (11,140). Insight into the pathogenesis of the XX true hermaph¬ rodite has come from studying the Sxr mouse. According to the Lyon hypothesis, extragonadal cells in the female have only one active X chromosome; additional X chro¬ mosomes are inactivated and form the sex chromatin bodies characteristic of the nuclei of female cells (159). In normal females, X inactivation is random and occurs early in em¬ bryonic life so that, in individual cells, either the matemalor paternal-derived X chromosome remains active (159). This random inactivation is believed to be operative in all somatic cells of the female; however, in the ovary, X in¬ activation differs from that in other tissues. By studying women heterozygous for electrophoretic variants of the Xlinked enzyme glucose-6-phosphate dehydrogenase, it was shown that oocytes both from adult and from fetal ovaries (82,83) express both X-linked alleles. Furthermore, both X chromosomes appear to be active in germ cells prior to entering meiosis (178). However, Kratzer and Chapman (149) have shown that, up to day 10 of embryogenesis, only one X chromosome is active in the embryonic mouse ovary and that the inactive X is reactivated as germ cells enter meiosis. Thus, random X inactivation occurs in ovarian development, as in other tissues of the female, and reacti¬ vation of the inactivated X subsequently occurs in the oogonia. To summarize: In testicular development the single X chromosome appears to be inactivated as a part of the XY body; in oogenesis, however, one of two X chromosomes is active during organogenesis of the ovary, and both X chromosomes are active during oogenesis itself. Random X-chromosome inactivation of the Lyon type involves chemical modification of the DNA and hence is thought to occur by a different mechanism than that responsible for Xchromosome inactivation in the XY body (241,263). X-autosome translocations have been described in mice in which the normal X chromosome (i.e., those not undergo¬ ing autosomal translocation) is preferentially inactivated (191). When XYsxr mice are crossed with female mice carrying such an X-autosome translocation trait (216) (the translo¬ cated X is designated XT), the XSxrXT offspring develop as sterile males, fertile females, or hermaphrodites (37,173). This finding can most simply be explained by assuming that the region of the X chromosome that carries the Sxr and the Tdy genes is inactivated in some cells but not others.

8 / Chapter 1 Inactivation is conceived as spreading to a variable extent from the inactive X chromatin to the Sxr fragment, and it is assumed that the extent of inactivation is transmittable to progeny cells after the time of X-chromosome inactivation during embryogenesis. In effect, such 46,XX individuals are mosaics, with some cell lines expressing Sxr and some not. The sex of the gonads (and hence the aggregate sex of the individual) depends on the proportion of cells expressing Sxr in the gonad primordia. If 30% of the cells in a gonadal primordia are XY, a normal testis will develop. When the X chromosome containing the Sxr gene is paired with an XT, XxXSxr females and hermaphrodites will occur at a low, but measurable, frequency. However, when the X chro¬ mosome to which Sxr is attached is paired with a normal X chromosome, half of the cells will express Sxr; hence, the majority of individuals will be sterile males, and females and hermaphrodites are rare. At a theoretical level, the same type of disorder, namely duplication of the male-determining gene(s) on the Y chro¬ mosome and translocation of these genes onto the X chro¬ mosome, could explain the development of either the XX male or the XX true hermaphrodite, depending on the com¬ pleteness and frequency of the inactivation of such testis¬ determining genes during the embryogenesis of the XX in¬ dividual carrying such a translocation trait. (It has yet to be proven that this model can explain the potential fertility of the XX true hermaphrodite.) This model has two implica¬ tions for the human disorder: (i) It implies that testis-de¬ termining genes, and hence Y-chromosome fragments, will be detectable in cells from 46,XX true hermaphrodites as well as from 46,XX men, and (ii) if the translocation is to an X chromosome (as in the case of the mouse Sxr) and not to an autosome, it explains why the XX male is more common than the XX true hermaphrodite.

GONADAL SEX Embryonic Development The gonad develops as a stratification of the coelomic epithelium on the medial aspect of the mesonephric kidney (the urogenital ridge) around the fourth week of gestational development in the human. Most of the cell types of the gonads are derived from the mesoderm cells of the urogenital ridges. The primordial germ cells originate, however, out¬ side the area of the presumptive gonad and are initially identifiable in the entoderm of the yolk sac. The germ cells, which are derived from primitive ectodermal cells of the inner cell mass (81), are distinguished from other cells of the developing embryo because of their large size as well as their large round nuclei and clear cytoplasm. Histochemically, they are characterized by high alkaline phosphatase activity and glycogen (169). The mechanism by which the germ cells differentiate from other cell types is not under¬ stood, but the process must commence early in embryo¬ genesis because primordial germ cells have been recognized

in the 4^-day-old human blastocyst (114). At the beginning of the fourth week of development the germ cells commence to migrate by ameboid movement (15,78,261) through the gut entoderm and .into the mesoderm of the mesentery, fi¬ nally ending up in the coelomic epithelium of the gonadal ridges. It is not known what entices the primordial germ cells to migrate to this area. However, once there, the prim¬ itive gonocytes move from the epithelium into the gonadal parenchyma, and lose their motile characteristics (60). Closely attached epithelial cells move with them into the underlying mesenchyme. The formation of the gonadal blastema is completed by weeks 5 to 6 of gestation in human embryos. At this time the primitive (“indifferent”) gonad is composed of three distinct cell types: (i) germ cells, (ii) supporting cells of the coelomic epithelium of the gonadal ridge that give rise to the Sertoli cells of the testis and the granulosa cells of the ovary, and (iii) stromal (interstitial) cells derived from the original mesenchyme of the gonadal ridge.

Histological Differentiation of the Fetal Gonad The first morphological sign of sexual dimorphism in the gonads is the development of the primordial Sertoli cells and their aggregation into spermatogenic cords in the fetal testis (139). In the human this occurs between weeks 6 and 7 of gestational development (130). In contrast to the early development of the fetal testis, the fetal ovary shows, no characteristic development until months later in embryo¬ genesis and initially is identified histologically early only by exclusion. At weeks 6 to 7 of gestation in the human, the ovarian epithelial components form irregular groups of cells around the primordial germ cells. At about the sixth month, the primitive granulosa cells organize around the dividing oocytes to form a single layer of follicular cells, thus establishing the primordial follicle (93). In general, gonadogenesis in other mammalian species is similar to that in humans in that histologic differentiation of the fetal testis precedes that of the fetal ovary by days to weeks.

The Role of the Germ Cells in Gonadal Development It seems unlikely that gonadal development or differen¬ tiation is dependent on the presence or type of germ cell that migrates into the coelomic epithelium of the gonadal ridge. Neither selective destruction of germ cells with drugs (175) nor surgical excision of primordial germ cells in the anterior germinal crest before they reach the gonadal primordium (168) inhibits gonadal development. Furthermore, mutant mice homozygous for the atrichosis gene are ge¬ netically deficient in germ cells, yet Sertoli cells differentiate and aggregate into tubules that are devoid of germ cells (112). Thus, it appears that the somatic cells can organize into an ovary or a testis irrespective of the presence or absence of the germ cells.

Sex Determination and Differentiation / Endocrine Differentiation of the Testis By demonstrating that castration of sexually indifferent rabbit embryos invariably results in female development of embryos of both sexes (Fig. 4), Jost (136,138) established that the induced phenotype in mammals is male and that secretions from the fetal testes are necessary for male de¬ velopment. Development of the female urogenital tract oc¬ curs in the absence of gonads and apparently does not require secretions from the fetal ovaries. Furthermore, Jost deduced that two substances from the fetal testes are essential for male development: (i) a nonsteroid hormone that acts ipsilaterally to cause regression of the mullerian duct (mullerian-inhibiting substance) and (ii) an androgenic steroid responsible for virilization of the wolffian duct, urogenital sinus, and urogenital tubercle.

Mullerian-Inhibiting Substance Mullerian-inhibiting substance is a large (—140,000 daltons) dimeric glycoprotein formed by the Sertoli cells of the fetal and newborn testis (56,58,202,203,234,247). It is thought to act locally to suppress mullerian duct develop¬ ment rather than as a circulating hormone (238; however, see ref. 119). Monoclonal antibodies to mullerian-inhibiting substance (186,246) block mullerian duct regression in a species-specific manner in in vitro bioassays (245), as well as in vivo (235). Although mullerian duct regression begins in the male embryo shortly after formation of the spermatic

9

cords in the fetal testis (14) the secretion of mullerianinhibiting substance is independent of spermatogenic tubule formation in the testis (166). Even though they are not primarily responsible for mul¬ lerian duct regression, androgens and estrogens appear to influence the process. For example, regression of rat mul¬ lerian ducts in organ culture is enhanced by testosterone, although testosterone alone is inactive (74,124). Interest¬ ingly, neither dihydrotestosterone nor estradiol affects mul¬ lerian duct regression in this system. In other systems, es¬ trogens interfere with mullerian duct regression (120, 122,144,170,228,232). Thus, although mullerian-inhibiting substance is required for mullerian duct regression, steroid hormones appear to influence its action. The concept that mullerian duct regression in male de¬ velopment is an active process is supported by studies of the persistent mullerian duct syndrome in the human (5,20,21,224). In this disorder, genetic and phenotypic males have fallopian tubes and a uterus (in addition to normal wolffian-duct-derived structures) as the result of either an autosomal or an X-linked gene defect. The pathogenesis of the disorder is uncertain, but is probably due to either a failure of production of mullerian-inhibiting substance by the fetal testis or a failure of the tissue to respond to the hormone. Persistence of the mullerian ducts is usually accompanied by failure of the testes to descend (133). Furthermore, mul¬ lerian-inhibiting activity is lower in biopsied testicular cells from newborn boys with cryptorchidism than from normal newborns (57). Therefore, it is conceivable that mullerian-

FIG. 4. Fetal castration experiment of Jost. The indifferent urogenital tract (top) under¬ goes male differentiation (bottom right) if a testis develops, or female differentiation (bottom left) if an ovary develops. Embryos castrated prior to sexual differentiation de¬ velop as phenotypic females (bottom mid¬ dle). (Adapted from ref. 138.)

10

/ Chapter 1

inhibiting substance plays a role in testicular descent, pos¬ sibly by influencing the cranial anchoring of the testis to the peritoneal fold.

Androgen The second developmental hormone of the fetal testis was identified by Jost as an androgenic steroid (136). Testos¬ terone, the principal testicular androgen formed in postnatal life, is also the androgenic steroid formed by the testes of rabbit and human embryos during male phenotypic devel¬ opment (155,218,259). Testosterone formation in the testis begins shortly after the onset of differentiation of the spermatogenic cords and is concomitant with the histological differentiation of the Leydig cells (38,98). The critical role of testosterone in the development of the male urogenital tract was deduced from three types of embryologic and endocrinologic evidence. First, as shown in Fig. 5, the fact that testosterone synthesis immediately pre¬ cedes the initiation of virilization of the urogenital tract in a variety of species suggested a cause-and-effect relationship between the two events (7,155,208,218,259). Second, the administration of androgens to female embryos at the ap¬ propriate time in fetal development causes male develop¬ ment of the internal and external genitalia (214). And third, administration of pharmacologic agents that inhibit the syn¬ thesis or action of androgens in embryogenesis impairs male development (95,187). This concept, furthermore, has been substantiated on ge¬ netic grounds. In the human, five single-gene defects in androgen synthesis are known to cause inadequate testos¬ terone synthesis and incomplete virilization of the male em¬ bryo (107,257). Severely affected males may develop as phenotypic women, with complete failure of virilization of the wolffian ducts, urogenital sinus, and external genitalia. At the other extreme, mildly affected men appear normal, except for developmental abnormalities such as hypo¬ spadias. The fact that the fallopian tubes and uterus are absent in such patients indicates that regression of the mullerian ducts takes place normally during embryogenesis and that mullerian regression is not primarily dependent on tes¬ tosterone biosynthesis and action. Regulation of testosterone synthesis in the fetal tes¬ tis. Many questions concerning the regulation of testoster¬ one synthesis and secretion by the fetal testis are not re¬ solved. The enzymatic differentiation of the fetal gonads that underlies the onset of endocrine function has been char¬ acterized in detail in the rabbit embryo. Enzymatic differ¬ entiation of fetal ovaries and testes in this species is apparent by day 18 of gestation and is manifested by an increase in the rate of 3(3-hydroxysteroid dehydrogenase activity in the fetal testis and by an increase in aromatase activity in the fetal ovary (179). At this time of development, activities of all other enzymes in the pathway of steroid hormone syn¬ thesis are similar in the ovaries and testes (Fig. 6) (90).

Gestational

Age, Weeks

FIG. 5. Enzymatic differentiation of the human fetal gonad. (Adapted from refs. 86 and 218.)

CHOLESTEROL

PREGNENOLONE Three

FETAL

Enzymes

V

TESTIS

7

ANDROSTENEDIOL (

I

FETAL OVARY

3/3-Hydroxysteroid Dehydrogenase

TESTOSTERONE Aromatase

ESTRADIOL

FIG. 6. Enzymatic differentiation of fetal rabbit ovaries and testes on day 18 of gestation.

Sex Determination and Differentiation / Thus, in the rabbit, changes in the rates of only a few enzymatic reactions in the gonads at a critical time in em¬ bryonic development have profound consequences for sex¬ ual differentiation. Furthermore, this enzymatic differentia¬ tion appears to be an autonomous function of the steroidogenic cells, because it occurs at the appropriate time in fetal testes cultured in defined medium without hormones (88,89) as well as in testes that fail to develop spermatogenic cords (196). Whether the actual rate of testosterone production in the rabbit fetal testis is regulated by fetal and/or placental go¬ nadotropins at the onset of testosterone synthesis is not clear. On the one hand, receptors for luteinizing hormone (LH) are present in fetal rabbit testes at the time of initial tes¬ tosterone synthesis (36); furthermore, these LH receptors are functional, as evidenced by the enhancement of testicular cyclic AMP formation (89) and cholesterol side-chain cleav¬ age activity (90) by human chorionic gonadotropin. On the other hand, basal, unstimulated cholesterol side-chain cleav¬ age activity in fetal rabbit testes in the absence of gonad¬ otropin stimulation appears to be sufficient to provide enough steroid substrate to support testosterone synthesis at a max¬ imum rate during the initial period of male phenotypic de¬ velopment. Later in embryogenesis, when sexual differ¬ entiation is far advanced, testosterone synthesis is enhanced by gonadotropin treatment (90). As a consequence, we be¬ lieve that in rabbits the onset of testosterone synthesis and the resulting differentiation of the male urogenital tract are independent of gonadotropin control. It is uncertain whether a similar situation exists in em¬ bryonic sexual differentiation in humans. LH receptors are present in human fetal testes as early as the twelfth week of gestation (117,180), and human fetal testes respond to human chorionic gonadotropin stimulation by exhibiting in¬ creased testosterone synthesis at this time (117). It is not known what happens between weeks 8 and 11, when the major portion of male phenotypic differentiation takes place. Because testosterone synthesis is gonadotropin dependent during the latter two-thirds of gestation, analogous to the situation in the rabbit embryo, it follows that those aspects of male sexual development that take place during this time— growth of the penis and descent of the testes—are probably gonadotropin-dependent in all species (206).

Endocrine Differentiation of the Ovary In many if not most species, endocrine differentiation of the fetal ovary, as evidenced by the appearance of the ca¬ pacity to synthesize estrogen, occurs simultaneously with the development of the ability of the fetal testis to synthesize testosterone (Fig. 5) (86,179). Although estrogen formation is not essential for normal female phenotypic development (136), estrogen may play a role in the development of the ovary itself. The change in the developing ovary that cor¬

11

relates with the onset of estrogen synthesis is the accumu¬ lation of lipid in the primitive granulosa cells (99).

Possible Role of Steroid Hormones in Gonadal Differentiation In some species it is possible to influence gonadal dif¬ ferentiation with sex hormones (reviewed in ref. 33). For example, in the embryonic male (ZZ) bird, treatment with estrogens during embryonic development leads to the de¬ velopment of an ovotestis. In amphibians and fish, treatment with steroid hormones of the opposite sex can lead to sex reversal of the gonads. In contrast with the striking effects of sex hormones on the differentiation of the gonads of birds and amphibians is the apparent failure to obtain comparable effects in mammalian embryos (25,104,135,237,252,253). Nevertheless, the bipotentiality of the indifferent mammal¬ ian gonad is suggested by transplantation experiments. Al¬ though it is felt that the testicular primordium represents remarkable stability in its development, fetal ovarian de¬ velopment appears to be more malleable. For example, sev¬ eral investigators have reported that fetal rodent ovaries develop “testis-like” structures when transplanted into male hosts (34,116,167,182,230,233), and these grafted fetal ovaries secrete testosterone in vitro (231). Thus in some situations, mammalian gonadal differentiation also appears to be influenced by the endocrine environment.

PHENOTYPIC SEX Urogenital Tract Development During the Indifferent Phase Prior to the eighth week of human development, the uro¬ genital tract is identical in the two sexes. The internal ac¬ cessory organs of reproduction develop from a dual duct system (wolffian and mullerian) that forms within the me¬ sonephric kidney early in embryogenesis (Fig. 7). Within the substance of the mesonephros, tubules connect primitive capillary networks with a longitudinal mesonephric (wolf¬ fian) duct. The wolffian duct extends caudally to the prim¬ itive urogenital sinus. At about 6 weeks the development of the paramesonephric (mullerian) ducts begins in embryos of both sexes as an evagination in the coelomic epithelium, just lateral to the wolffian ducts. This evagination develops into a tubular structure, the caudal end of which becomes intimately associated with the wolffian duct (i.e., no base¬ ment membrane separates their epithelia) (110). Whether the mullerian duct “splits off” from the wolffian duct in its later caudal development to become an independent duct system emptying into the urogenital sinus or whether the wolffian duct simply acts as a guide for the subsequent evolution of the mullerian duct from the coelomic epithelium is uncertain. However, mullerian duct development cannot take place in the absence of the wolffian duct (110). At the

12 / Chapter 1 INDIFFERENT STAGE gonad mesonephros Mullerian duct Wolffian duct

epididymis testis

ovary Fallopian tube

as deferens

seminal vesicle

urethralgroove

prostate

: •F.’jVjV. 11

scrotum-^ I MALE

FIG. 7. Differentiation of the internal genitalia. (From ref. 256.) end of the indifferent phase of phenotypic sexual differ¬ entiation (prior to week 8 of gestation), a dual duct system (wolffian and mullerian) constitutes the anlagen of the in¬ ternal accessory organs of reproduction (Fig. 7). The ter¬ mination of the mesonephric ducts in the urogenital sinus divides the sinus into an upper and a lower portion. The upper portion, the vesicourethral canal, is involved in the development of the bladder and the upper urethra. The lower portion contributes to the development of the external gen¬ italia. Prior to week 6 of gestation, the anlagen of the external genitalia are also indistinguishable in the two sexes. The genital eminence is a rounded mass between the umbilicus and the tail (226) and is composed of a genital tubercle flanked by prominent genital swellings. The opening of the urogenital sinus between the genital swellings (the urethral groove) is surrounded by genital folds (Fig. 8). At week 7 of gestation in the human, the genital tubercle begins to elongate; a shallow, circular depression defines the glans of the tubercle. At this stage of development, there are no remarkable differences between the external genitalia of male and female embryos.

Male Development The first sign of male differentiation of the urogenital tract is degeneration of the mullerian ducts adjacent to the testes, a process that begins just after formation of the spermatogenic cords in the testes. Eventually, the mullerian ducts of the male undergo almost complete regression. The transformation of the wolffian ducts into the male

prepuce body of penis scrotal raphe

H?- •

fr;■'-I "~f • ■* L ,i tt‘

FIG. 8. Differentiation of the external genitalia. (From ref. 256.)

reproductive tract begins subsequent to the onset of mul¬ lerian duct regression. The portion of the wolffian duct adjacent to the testis becomes convoluted to form the epi¬ didymis; the central portion of the duct becomes the mus¬ cular vas deferens. The seminal vesicles develop as buds off the lower portions of the wolffian ducts just before they enter into the urogenital sinus. The prostatic and membra¬ nous portions of the male urethra develop from the pelvic portion of the urogenital sinus. The prostate originates as a series of endothermal buds off this portion of the urogenital sinus (10,143,157). The external genitalia of the male begin to develop shortly after the onset of virilization of the wolffian ducts and uro¬ genital sinus (Fig. 8). The genital tubercle elongates, and the urethral folds begin to fuse over the urethral groove from posterior to anterior. The two urogenital swellings move posterior to the genital tubercle and eventually fuse to form the scrotum. The elongated urogenital cleft becomes closed to form the penile urethra (Fig. 8). These events in male development are completed relatively early (during the first trimester) in human fetal development and are dependent on hormonal secretions from the testis.

Sex Determination and Differentiation

/

13

Two aspects of male phenotypic development take place during the late phases of virilization. The first involves growth of the male phallus. Just after closure of the male urethra is complete, there is little difference in the size of the genital tubercle in the two sexes. However, under the influence of androgens from the fetal testis, the male phallus grows during the latter phases of fetal development and by the time of birth is much larger than the urogenital tubercle of the female. Testicular descent also takes place during later stages of fetal development (92). Testicular descent is both complex and incompletely understood. For didactic purposes, the process can be divided into three phases. The first phase (transabdominal movement) involves, at a minimum, de¬ generation of the portion of the peritoneal fold that anchors the cranial part of the gonad to the abdominal wall, short¬ ening of the caudal gonadal ligament (gubemaculum), and rapid growth of the abdominal-pelvic region of the fetus. As a result, the testis comes to rest against the anterior abdominal wall in the inguinal region. The second phase involves formation of the processus vaginalis and devel¬ opment of the inguinal canal and scrotum. Increasing in¬ traabdominal pressure is believed to cause a herniation in the abdominal wall (the processus vaginalis) along the course of the inguinal portion of the gubemaculum. Continued pressure causes enlargement of the processus vaginalis around the gubemaculum and leads to formation of the inguinal canal. The gubemaculum increases in size until the diameter of the inguinal canal approaches that of the testis. In the final stage of testicular descent the abdominal testis traverses the inguinal canal and comes to rest in the scrotum. Descent of the testis into the scrotum probably involves a progressive degeneration of the proximal portion of the gubemaculum. Although the process is at least, in part, androgen dependent (206), the early stages (e.g., transabdominal movement) of testicular descent may be the result of other factors (118).

Female Development The internal reproductive tract of the female is formed from the mullerian ducts; the wolffian ducts persist only in remnant form. The cephalic ends of the mullerian ducts (the portions derived from coelomic epithelium) are the anlagen of the fallopian tubes, whereas the caudal portions fuse to form the uterus (Fig. 7). Contact of the mullerian ducts with the urogenital sinus induces formation of the uterovaginal plate (an intense proliferation of endodermal cells) between the mullerian ducts and the urogenital sinus (194). Although it is felt that both the mullerian ducts and wolffian ducts contribute to the formation of the uterovaginal plate, the relative degree to which they contribute is unknown (16,28,110). The cells of the uterovaginal plate proliferate, thus increasing the distance between the uterus and the uro¬ genital sinus (Fig. 9). Later, the central cells of this plate break down to form the lumen of the vagina.

FIG. 9. Development of the uterus and vagina. (From ref. 256.)

In contrast to the male, in which the phallic and pelvic portions of the urogenital sinus are enclosed by fusion of the genital swellings, most of the urogenital sinus of the female remains exposed on the surface as a cleft into which the vagina and urethra open (Fig. 8). The urogenital tubercle in the female undergoes limited growth and development to form the clitoris.

Breast Development in Both Sexes Breast development occurs along “mammary lines,” which are bilateral epidermal thickenings that extend from the fore¬ limbs to the hindlimbs on the ventral surface of the embryo. In human development, these “mammary lines” largely dis¬ appear except for a small portion on each side of the thoracic region that condenses and penetrates the underlying mes¬ enchyme. This single pair of mammary buds undergoes little change until the fifth month of human development, when secondary epithelial buds appear and nipples develop. Pro¬ liferation of the ductules occurs throughout the remainder of gestation so that, by the time of birth, 15 to 25 separate

14

/ Chapter 1

glands are present, each of which is connected to the exterior through the nipple. In some species, breast development in males is inhibited by androgens during embryogenesis (96,126,147), but this does not occur in humans; the de¬ velopment of the breast in boys and girls is identical prior to puberty (200).

Endocrine Control of Male Phenotypic Differentiation As a consequence of experiments demonstrating that cas¬ tration of sexually indifferent rabbit embryos invariably re¬ sults in female development (Fig. 4), Jost recognized that the induced phenotype in mammalian embryos was male and that secretions from the fetal testis were necessary for male phenotypic development (136,137). Development of the female urogenital tract apparently does not require se¬ cretions from the fetal ovaries, since female phenotypic development occurs in the absence of gonads. Furthermore, Jost deduced that two substances from the fetal testes are essential for male development: (i) a polypeptide that acts ipsilaterally to cause regression of the mullerian ducts and (ii) an androgenic steroid, testosterone, responsible for vir¬ ilization of the wolffian duct, urogenital sinus, and genital tubercle.

Mechanism of Action of Mullerian-Inhibiting Substance The initial endocrine function of the fetal testis is probably secretion of the protein hormone that causes regression of the mullerian ducts, namely, mullerian-inhibiting substance. Although mullerian-inhibiting substance has been purified and much is known about the structure of the protein itself (26,54,134) and about the genes that code for the protein in humans and cows (35,201), the mechanism of action of the hormone is poorly understood. Dissolution of the base¬ ment membrane and condensation of the mesenchymal cells around the ducts is an early event in mullerian duct regres¬ sion (62,113,204,236). Mullerian-inhibiting substance may act by modulating cell-surface protein phosphorylation in a manner similar to the mechanism of action of epidermal growth factor (57,121). The finding that partially purified mullerian-inhibiting substance is cytotoxic to a human ovarian-cancer-derived cell line, but not to cell lines derived from an adenocarci¬

noma of the colon, raises the possibility that mullerianinhibiting substance may also inhibit growth of malignant tumors of the female genital tract (55,59,79). %

Mechanism of Androgen Action Basic model. The current concept of how androgens act within target cells is schematically depicted in Fig. 10. Testosterone, the major androgen secreted by the testis and circulating in plasma, is thought to enter target tissues by a passive diffusion process. In some cells, testosterone is converted to dihydrotestosterone by a 5a-reductase enzyme present within the cells. In androgen target tissues, testos¬ terone or dihydrotestosterone binds to a specific high-affinity, intracellular receptor protein. Subsequently the hor¬ mone-receptor complex undergoes a poorly understood transformation process in which the hormone-receptor com¬ plex acquires the capacity to bind to DNA and other anionic substances. It is not clear whether the primary hormonereceptor interaction takes place in the nucleus or the cyto¬ plasm. However, the transformed nuclear hormone-receptor complexes are presumed to interact with specific acceptor sites on the chromosomes. The nature of the acceptor sites within the nucleus (i.e., whether protein or DNA) and their number are not clear, but the consequence of the interaction of the hormone-receptor complexes with chromatin is to increase the transcription of tissue-specific structural genes, with the subsequent appearance of new messenger RNAs and new proteins in the cytoplasm of the cell. If the andro¬ gen-receptor system is analogous to other steroid hormone systems, it is likely that the androgen-receptor complexes bind to DNA at specific regulatory sites near the structural genes under regulatory control by the hormones (48,199). On the basis of a variety of evidence (see below), it is now felt that testosterone-receptor complexes mediate fetal wolf¬ fian duct virilization and that dihydrotestosterone-receptor complexes mediate differentiation of the male external gen¬ italia. The androgen receptors from fetal urogenital sinus and urogenital tubercles appear to be identical to those of adult prostate (84), so that the hormone is presumed to act by the same mechanism in embryonic as in postnatal life. 5aDihydrotestosterone binds to the androgen receptor of most species with greater affinity than does testosterone (84,146, 165,254), and the dihydrotestosterone-receptor complex is

FIG. 10. Androgen action. (T) Testosterone; (D) dihy¬ drotestosterone; (R) androgen receptor; (R*) trans¬ formed androgen receptor.

Sex Determination and Differentiation

more readily transformed to the DNA-binding state than is the testosterone-receptor complex (145). The net conse¬ quence is that dihydrotestosterone formation generally am¬ plifies the androgenic signal. Since only one receptor ap¬ pears to mediate the action of both androgens, a central, as yet unresolved, issue of androgen physiology is concerned with the mechanism by which testosterone and dihydrotes¬ tosterone exert different actions during embryogenesis. One possibility is that the two hormones act in exactly the same manner to promote virilization. By this schema, dihydro¬ testosterone, although amplifying the hormonal signal within most androgen target cells, is not absolutely required for androgen action, provided that the intracellular concentra¬ tion of testosterone is sufficiently high and that sufficient time is available to allow registration of the weaker hor¬ monal signal produced by testosterone. Alternatively, tes¬ tosterone may promote virilization of the wolffian ducts by an indirect means, analogous to the involvement of eryth¬ ropoietin in androgen-mediated control of erythropoiesis (72). In some tissues (e.g., hypothalamus), testosterone can be aromatized to estradiol, and androgen action is, paradoxi¬ cally, mediated by an estrogen. Role of dihydrotestosterone in virilization. Separate roles for testosterone and dihydrotestosterone in male differen¬ tiation were postulated on the basis of studies of androgen metabolism in embryos (218,255,258). In rat, rabbit, guinea pig, and human embryos, 5a-reductase activity is maximal in the urogenital sinus and urogenital tubercle prior to vir¬ ilization; however, the enzyme is virtually undetectable in the wolffian duct derivatives until after virilization is ad¬ vanced (Fig. 11). Therefore, it was deduced that testosterone mediates virilization of the wolffian ducts, whereas dihy¬ drotestosterone is responsible for differentiation of the male urethra, prostate, and external genitalia (258). This hypothesis was substantiated by studies of patients with 5a-reductase deficiency, a rare autosomal recessive mutation causing abnormal sexual development in affected men (75,198,251). Affected persons are 46,XY males who have predominantly female external genitalia in association with bilateral testes. The internal genitalia (epididymis, vas deferens, seminal vesicle, and ejaculatory duct) that are derived from the wolffian ducts are virilized normally, how¬ ever, and terminate in a pseudovagina. At the time of ex¬ pected puberty, testosterone production increases into the male range, and the external genitalia may virilize to a limited extent. Gynecomastia does not develop; axillary and pubic hair develop normally. There is considerable heterogeneity in the mutant 5areductase among different families with the disorder (75,127,260). The most common defect appears to be a marked deficiency in the amount of a catalytically active 5a-reductase enzyme (183,184). Some patients, however, have structural abnormalities of the enzyme that affect co¬ factor (NADPH) and/or steroid binding (128,151). Because a mutation that causes decreased dihydrotestos¬ terone formation has not yet been described in an animal

6-8

/

15

8-10 10-12 12-13 13-14 14-17 17-21 >21 Gestational Age, Weeks

FIG. 11. Developmental study of 5a-reductase activity in uro¬ genital tracts of human fetuses. (Adapted from ref. 218.)

species, direct characterization of the separate roles for tes¬ tosterone and dihydrotestosterone in formation of the male phenotype has been difficult (22,23,152). However, the administration of 5a-reductase inhibitors to pregnant rats during the period of embryonic sexual differentiation re¬ produces many of the characteristics of the 5a-reductasedeficiency phenotype in the male offspring. Virilization of the external genitalia is impaired, whereas no effect on the male differentiation of wolffian-duct-derived structures is apparent (23,125). Since the androgen receptor system of the wolffian duct derivatives, like other androgen receptors, appears to bind dihydrotestosterone preferentially (85), it has been difficult to uaderstand why testosterone can me-

16

/ Chapter 1

diate wolffian differentiation but cannot virilize the urogen¬ ital sinus and external genitalia. It is conceivable that the local concentration of testosterone in the wolffian ducts is exceptionally high because it is secreted directly from the testis into the lumen of the wolffian duct; it is also possible that this high concentration of testosterone compensates for its relative ineffectiveness as an androgen. Two types of evidence are in keeping with this theory. First, active im¬ munization of pregnant rabbits against testosterone reduces circulating androgen and causes pseudohermaphroditism in male offspring similar to the phenotype described for human males with 5a-reductase deficiency (12,244), suggesting that androgens that are not exposed to antibody (androgens within the lumen of the wolffian ducts) may remain effec¬ tive. Second, a heritable trait in the rat causes unilateral hypoplasia of the testis in 50% of males (123). Hypoplasia of the- testis is accompanied by ipsilateral aplasia of the epididymis and vas deferens despite the fact that prostate development and virilization of the external genitalia are normal, presumably mediated by plasma androgens derived from the normal testis. This finding is consistent with the concept that a noncirculating factor from the testis (testos¬ terone?) causes virilization of the adjacent wolffian duct. A perplexing aspect of 5a-reductase deficiency is the partial virilization that occurs in some patients at the time of expected puberty (198,205). Late virilization in these patients may be caused by the combination of (a) higher levels of plasma testosterone at puberty than during embryogenesis and (b) the presence of some residual 5a-reductase activity in all patients with this defect who have been char¬ acterized to date. Role of androgen receptor in male development. A spe¬ cific, high-affinity receptor protein mediates the action of both testosterone and dihydrotestosterone in all androgendependent tissues. Androgen receptors in fetal urogenital tissues have characteristics similar to those in adult andro¬ gen-dependent tissues, and they probably mediate viriliza¬ tion of the fetus by the same mechanisms as in postnatal life (85). Studies of single-gene mutations that impair an¬ drogen receptor function are in keeping with this concept. The first disorder of the androgen receptor to be char¬ acterized in molecular terms was the testicular feminization (Tfm) mutation in the mouse, an X-linked disorder in which affected males have testes and normal testosterone produc¬ tion but differentiate as phenotypic females (8,97,161). The mullerian-inhibiting function of the fetal testis is presumed to be normal because no mullerian duct derivatives (uterus or fallopian tubes) are present in affected males. Dihydro¬ testosterone formation is normal. However, profound re¬ sistance to androgen action results in failure of all androgenmediated aspects of male development in the wolffian duct, urogenital sinus, and external genitalia. Functional cyto¬ plasmic androgen receptors are not detectable (27,84,242) (Fig. 12). Consequently, the hormone cannot reach the nu¬ cleus of the cell and interact with the chromosomes. Studies of subjects with the human counterpart of the Tfm

1000 r

14,

C- Albumin

C

\

o

o o 500

W \ 9 w

CL O

•t

/Vx/Y

Tfm/Y■V L

J

•

°:r

'

V'

J-'oo-6-00-0 o6__

10

20

Fraction Number

FIG. 12. Sucrose density gradient analysis of [3H]dihydrotestosterone binding in kidney cytosol from normal (X/Y) and androgen-resistant (Tfm/Y) male mice. (Adapted from ref. 242.)

mouse have provided additional insight into the role of the androgen receptor in embryonic virilization (260). Defects in the human androgen receptor cause a spectrum of ab¬ normalities that vary in severity, ranging from men with mild defects in androgen action to phenotypic women with testicular feminization. Women with testicular feminization usually come to the attention of the physician when they are evaluated for primary amenorrhea. The karyotype is 46,XY, but the general habitus is female. Axillary, facial, and pubic hair is absent or scanty. The external genitalia are unambiguously female, and the vagina is short and blind¬ ending. All internal genitalia are absent except testes, which may be located in the abdomen, along the course of the inguinal canal, or in the labia majora. Female breast de¬ velopment occurs at the time of expected puberty and is probably due to increased estrogen synthesis by the testis (162). A small percentage of patients with the phenotype of testicular feminization have axillary and pubic hair as well as a modest degree of virilization (164). These patients are designated as having an “incomplete” form of testicular feminization. Patients with Reifenstein syndrome (most commonly with hypospadias, azoospermia, and gyneco¬ mastia; see ref. 260) and phenotypically normal men with infertility (1,2) complete the spectrum of individuals with disorders of the androgen receptor. The molecular defect in some patients is similar to that in the Tfm mouse (i.e., no high-affinity androgen receptor can be detected; see refs. 109 and 142). It is not known whether the absence of binding in these instances is due to absence of the receptor protein or to a defect in the protein that precludes its ability to bind androgen. Other patients

Sex Determination and Differentiation

with testicular feminization have either a diminished amount or a qualitative abnormality of the receptor. Qualitative ab¬ normalities of the androgen receptor were initially identified in studies of thermolability of androgen receptors in skin fibroblasts cultured from patients with phenotypic androgen resistance (105). Subsequently, other mutations have been identified in which the cytosolic androgen receptor is un¬ stable in molybdate-containing buffers (106). Studies of the process by which androgen-receptor complexes are “trans¬ formed” to the DNA-binding state have identified yet an¬ other subset of patients with qualitatively abnormal andro¬ gen receptors in whom the abnormality of the receptor becomes manifest during transforming conditions (145,146). Quali¬ tative abnormalities of the androgen receptor that have been described in other laboratories include altered affinity of binding (24), impaired nuclear retention (67), and impaired augmentation of receptor binding following prolonged in¬ cubation with ligand (141). Since some patients with qual¬ itatively abnormal receptors have androgen resistance as profound as that observed in patients in whom androgen binding cannot be detected, it appears that such structural abnormalities can totally prevent function of the receptor. The normal gene that codes for the androgen receptor and the mutant gene that causes absence of the receptor are Xlinked (176,177); mutations that cause qualitative abnor¬ malities of the receptor are probably allelic mutations of the same gene (68). In some patients, androgen resistance occurs despite ap¬ parently normal androgen receptors. It was originally pos¬ tulated that androgen resistance in these patients was caused by defects in the later phases of androgen action, the socalled “postreceptor androgen resistance” (3). Individuals from 10 such families that have been analyzed in our lab¬ oratory span a phenotypic spectrum ranging from testicular feminization to the infertile male syndrome (108). As more sensitive techniques for characterizing qualitative abnor¬

/

17

malities in the androgen receptor have been developed, many of these subjects, including those in the original family described by Amrhein et al. (3), are known to have subtle functional alterations of the androgen receptor. Role of the mesenchyme in androgen action. In many tissues the embryonic mesenchyme (stroma) appears to con¬ trol the differentiation of the associated epithelium (re¬ viewed in ref. 46). A compelling case for stromal-epithelial interactions in androgen action comes from studies of tissue recombinants in the development of the urogenital sinus. When mesenchyme from the urogenital sinus of embryonic mice is recombined with homotypic urogenital sinus epi¬ thelium and grown as intraocular grafts in male animals, the epithelium acquires the characteristics of the glandular epithelium of the prostate (43,44). In contrast, heterotypic recombinants of epithelium from urogenital sinus and mes¬ enchyme of integumental origin are incapable of glandular (prostatic) development, but form keratinized epithelium characteristic of skin under similar circumstances (Fig. 13). Furthermore, mesenchyme derived from the urogenital sinus of androgen-resistant (Tfm/Y) mice is incapable of mediat¬ ing prostatic growth when recombined with normal uro¬ genital sinus epithelium and exposed to androgen. However, when reciprocal recombinants (Tfm/Y epithelium with nor¬ mal mesenchyme) are made, prostate development occurs (45,150). Autoradiographic studies of androgen binding in the uro¬ genital sinus of developing rats (229) provide additional insight into the role of the mesenchyme in morphogenesis of the prostate. At the time of prostatic bud formation, androgen-binding sites are located predominantly over the nuclei of mesenchymal cells that surround the developing buds. The urogenital sinus of female embryos also has an¬ drogen-binding sites in the nuclei of mesenchymal cells. In contrast, no labeling was detected in the epithelia of fetal urogenital sinuses. After postnatal day 10, androgen binding

+ UROGENITAL SINUS MESENCHYME

UROGENITAL SINUS EPITHELIUM

EPITHELIUM (PROSTATE)

FIG. 13. Diagram of tissue recombinant experi¬ ments of Cunha. (Adapted from ref. 46.)

INTEGUMENT MESENCHYME

UROGENITAL SINUS EPITHELIUM

KERATINIZED EPITHELIUM (SKIN)

18 / Chapter 1 was present in the epithelial cells of the prostate, and la¬ beling in mesenchymal cells became less prominent. These results suggest that, during morphogenesis of the male uro¬ genital tract, androgen action is initiated through mesen¬ chymal cells, whereas androgen responses of the prostate after differentiation are mediated by interaction with epi¬ thelium and mesenchyme. A similar system is responsible for the androgen-mediated regression of the embryonic mammary bud in the mouse (61,148). However, in this tissue the response to androgen requires specific interaction of the mesenchyme with mam¬ mary epithelium (115), suggesting that in some cases the epithelium may also play a role in the differentiation pro¬ cess. There are also indications that epithelium of meso¬ dermal origin, when recombined with androgen-responsive mesenchyme, responds differently than does epithelium of endoderm (46). Elucidating the nature of these mesenchy¬ mal-epithelial interactions is of critical importance in un¬ derstanding androgen-mediated differentiative processes. Despite the fact that a great deal of information has been accumulated about the hormones involved in differentiation of the urogenital tract and about the cellular sites of their initial actions, little is known about the specific gene prod¬ ucts that are synthesized in response to the hormones or how such products direct cellular organization during embryogenesis. Effects of androgens in female embryos. Female embryos have the same androgen receptor system in the urogenital tract as do male embryos (85). For example, androgen re¬ ceptors are as readily detectable in the same regions of the female and male urogenital tracts (Fig. 14). Therefore, it is not surprising that exposure to androgens during the time of sexual differentiation causes profound virilization of fe-

3200

d Prostatic

2400 -

C

Urethra

o o

? Urethra + - Vagina

ui 1600 Q_ O

800

A, 4 ■

-I?

•#' /

•■

Urinary

,A\

o o «\

St*.0 0'0'

\

_i_°'°i8s 10

20

Fraction Number

FIG. 14. Sucrose gradient analysis of [3H]dihydrotestosterone binding in cytosol of urogenital sinuses and bladder of fetal rabbits on the 28th day of gestation.

male offspring (214). The anatomical consequences of such an experiment in rats are demonstrated in Fig. 15. Figure 15A and B show urogenital tracts dissected from female and male newborn rats whose mothers had been treated with oil (control) from day 14 through day 21 of gestation. Figure 15C shows a urogenital tract from a newborn female exposed in utero to an inactive androgen analog (5(3-dihydrotestosterone). Figure 15D shows a urogenital tract from a female exposed to 5a-dihydrotestosterone. Exposure to this active androgen caused differentiation of the wolffian ducts in this female embryo into prominent epididymides, vasa deferentia, and seminal vesicles. Furthermore, the urogenital si¬ nuses from female rats exposed in utero to 5a-dihydrotestosterone contain prostatic buds and a male-type urethra and exhibited no vaginal development (F. W. George, unpub¬ lished observations). The fact that females virilize when exposed to androgens indicates that differences in anatom¬ ical, development between males and females depend on differences in the hormonal signals themselves and not on differences in the hormone receptors in target tissues. It also follows that the sexual fate of the normal embryo is deter¬ mined largely by whether testosterone production com¬ mences in the fetal testis at the precise time in embryonic development. The most common cause of virilization of human female embryos is congenital adrenal hyperplasia (188). Inherited mutations that result in decreased synthesis of cortisol in the adrenal gland lead to a compensatory increase in ACTH secretion by the pituitary. This, in turn, leads to an increase in adrenal androgen secretion. The adrenal androgens then virilize the external genitalia of the female. The internal genitalia are, however, not virilized, and wolffian duct rem¬ nants are no more prominent in women with congenital adrenal hyperplasia than in normal women. Therefore, it is likely that the degeneration of the wolffian ducts precedes the onset of adrenal androgen synthesis or that the wolffian ducts are insensitive to the predominant androgens formed in affected females.

DO HORMONES PLAY A ROLE IN FEMALE PHENOTYPIC DEVELOPMENT? In the eutherian mammal, embryogenesis takes place in a “sea” of hormones (steroidal and nonsteroidal) derived from the placenta, the maternal circulation, the fetal adrenal gland, the fetal testis, and possibly from the fetal ovary. It is not known whether any of these substances influence female phenotypic differentiation or development. It is con¬ ceivable that estrogens or progestogens, or both, are in¬ volved in the growth and maturation of the female urogenital tract during the latter part of embryonic development, even if they are not required for their differentiation. Presumably, fetal castration experiments [such as those performed by Jost (136) to elucidate the role of the fetal testis in male development in the rabbit] would be uninformative, since

Sex Determination and Differentiation /

19

FIG. 15. Virilization of the female rat urogenital tract caused by 5a-dihydrotestosterone administration to the mother. A and B, respectively: Female and male uro¬ genital tracts of newborn rats from a mother given oil from days 14-21 of gestation. C: Female urogenital tract following administration of 16 mg/day of 5(3-dihydrotestosterone. D: Female urogenital tract following administration of 16 mg/day of 5a-dihydrotestosterone. (o) Ovary; (u) uterus; (c) coils of oviduct; (v) vagina; (t) testis; (e) epididymis; (vd) vas deferens; (sv) seminal vesicle; (p) prostate. (From ref. 214.)

removal of fetal ovaries would not remove the major source of female hormones. Experimental agents that block estro¬ gen synthesis or action interfere with placental function and precipitate abortion. Furthermore, no mutations have been identified that result in either deficient estrogen synthesis or resistance to estrogen action. This is in contrast to the sit¬ uation in regard to testosterone synthesis and action where single-gene mutations that interfere with both processes have been characterized in many species (260). In the rabbit, estrogen synthesis is temporarily activated in both male and female embryos at the time the blastocyst implants in the uterine wall between days 6 and 7 of gestation (87) (Fig. 16). Eater in the embryogenesis of the rabbit, estrogen syn¬ thesis (aromatase activity) is activated in placenta (tran¬ siently), brain, and ovary, but specific activity of aromatase in these tissues is never as high as in the implanting blas¬ tocyst (Fig. 16). Estrogens may be necessary for implan¬

tation and survival of the blastocyst (52,53), suggesting that estrogen action is essential for life in eutherian mammals. If this is true, mutations that prevent either the synthesis of, or the response to, estrogens may be lethal at an early stage of development by preventing implantation of the blas¬ tocyst. In many species, estradiol formation is initiated in the ovary before definitive histological differentiation has oc¬ curred; it is possible that cellular organization of the ovary may be mediated, in part, by estrogens formed locally (99), analogous to the postulated role of testosterone in maturation of the spermatogenic cords of the testis. In contrast to eutherian mammals in which sexual dif¬ ferentiation occurs in utero, marsupial young are bom sex¬ ually indifferent. Development of the sexual phenotypes takes place in the pouch, independent of the maternal milieu, and the pouch young are accessible for experimentation

20

/

Chapter 1

FIG. 16. Distribution of aromatase ac¬ tivity in the developing rabbit embryo. (Composite of data in refs. 87, 91, and 179.)

4

6

8

10

12

14

16

18

20

22

24

£6

28

30

DAY OF GESTATION

throughout sexual development. Marsupials therefore rep¬ resent an important model system for studying the hormonal factors controlling sexual differentiation. Bums (32) studied the effects of the administration of androgen or estrogen on the development of the sexual phenotypes in newborn opos¬ sums (Didelphis virginiana). Testosterone administration caused a marked hypertrophy and male differentiation of the phallus in both male and female pouch young. In con¬ trast, estrogen caused female-type development of the phal¬ lus and cloacal region in both sexes. Similar effects have been reported in the gray opossum (Monodelphis domestica) (73). Thus, estrogens appear to play a role in the devel¬ opment of the female external genitalia in these marsupial species. The development of the pouch in female, and of the scrotum in male, opossums occurs 10 days after birth, and the two structures are thought to derive from common anlage (17). Pouch development is the earliest evidence of sexual dimorphism that can be identified grossly. Interestingly, in the opossum pouch young that Bums treated with androgen or estrogen, some of which were treated soon after entering the pouch, the development of the pouch or scrotum was not influenced. Thus, phenotypic differentiation in this marsupial appears to be an exception to Jost’s formulation for sexual differ¬ entiation in eutherian mammals in that (a) estrogens may play a role in development of the female urogenital tract and (b) some aspects of the sexual phenotype (scrotum and pouch development) appear to be independent of hormonal control. SUMMARY Jost’s formulation has proved to be a powerful paradigm for understanding normal and abnormal sexual differentia¬

tion. Chromosomal sex determines gonadal sex, and go¬ nadal sex determines phenotypic sex. A minimum of 19 genes have been implicated in sexual differentiation in hu¬ mans (257). Some of these genes are located on the sex chromosomes, some on the autosomes. Thus, the relatively simple mechanism that imposes male development on the indifferent embryo requires the participation of many genes common to both the male and female embryo. Much of our understanding of the process of sexual differentiation is due to the fact that, unlike many other congenital defects, ab¬ normalities of sexual differentiation are not lethal, and such individuals, even those with mild abnormalities of sexual development, come to the attention of physicians and sci¬ entists and are systematically studied. Many of these ab¬ normalities are due to single-gene defects, and detailed anal¬ yses of these disorders in humans and animals have provided a great deal of insight into the endocrine, molecular, and genetic determinants that regulate sexual differentiation. Determinants on the Y chromosome cause the indifferent gonad to develop into a testis. Two hormonal secretions from the fetal testis, namely mullerian-inhibiting substance and testosterone, then transform the indifferent urogenital tract into one that is characteristic of the male. Mullerianinhibiting substance, secreted by the Sertoli cells, causes regression of the female (mullerian) duct system. Testos¬ terone, secreted by the Leydig cells, is responsible for the remainder of male development, including stabilization and differentiation of the wolffian ducts into the male accessory organs of reproduction as well as differentiation of the male external genitalia and prostate. The role of hormones in female development is less clear. In the opossum, ovarian hormones may play a role in the differentiation of the female urogenital tract and develop¬ ment of the pouch. However, in the eutherian mammal it has not been possible to design experiments to determine

Sex Determination and Differentiation

whether hormones from the placenta, the maternal circu¬ lation, or the fetal ovary play an essential role in female development. A major portion of androgen action in the fetus and in postembryonic life is mediated by 5a-reduced metabolites of testosterone rather than by testosterone itself. Thus, a genetic deficiency in the 5a-reductase enzyme that catalyzes the formation of dihydrotestosterone from testosterone im¬ pairs androgen action and can cause male pseudohermaph¬ roditism. Although wolffian duct development is apparently normal in these individuals, other aspects of male devel¬ opment are defective. Testosterone and dihydrotestosterone act via a common receptor to virilize male fetuses. Consequently, normal phe¬ notypic sexual development is determined by the presence (in males) or the absence (in females) of specific hormonal signals at the critical time in embryonic development. At the time of sexual differentiation, differences in the activity of only a few enzymes involved in steroid biosynthesis in the gonads have profound consequences on the character of the hormones secreted and thus on the sexual development of the fetus. Although we now understand, in considerable detail, the hormonal and genetic factors that are responsible for mam¬ malian sexual differentiation, many fundamental issues in the embryonic development of the urogenital tract remain poorly understood. What, for instance, is the mechanism by which the same hormonal signal is translated into dif¬ ferent physiologic effects in different tissues? What are the molecular and cellular changes that cause these diverse differentiative events? Ultimately, these fundamental issues of embryogenesis will have to be clarified before it will be possible to understand the entire program by which the myriad of genetic determinants and hormones interact to cause the development of phenotypic sex.

REFERENCES 1. Aiman, J., and Griffin, J. E. (1982): The frequency of androgen receptor deficiency in infertile men. J. Clin. Endocrinol. Metab., 54:725-732. 2. Aiman, J., Griffin, J. E., Gazak, J. M., Wilson, J. D., and MacDonald, P. C. (1979): Androgen insensitivity as a cause of infertility in oth¬ erwise normal men. N. Engl. J. Med., 300:223-227. 3. Amrhein, J. A., Meyer, W. J. Ill, Jones, H. W. Jr., and Migeon, C. J. (1976): Androgen insensitivity in man: Evidence for genetic heterogeneity. Proc. Natl. Acad. Sci. USA, 73:891-894. 4. Andersson, M., Page, D. C., and de la Chapelle, A. (1986): Chro¬ mosome Y-specific DNA is transferred to the short arm of X chro¬ mosome in human XX males. Science, 233:786-788. 5. Armendares, S., Buentello, L., and Frenk, S. (1973): Two male sibs with uterus and fallopian tubes. A rare, probably inherited disorder. Clin. Genet., 4:291-296. 6. Armendares, S., Salamanca, F., Cantu, J. M., del Castillo, V., Nava, S., Dominguez-de-la-Piedra, E., Cortes-Gallegos, V., Gal¬ legos, A., Cervantes, C., and Parra, A. (1975): Familial true hermaphrodism in three siblings. Clinical, cytogenetic, histological and hormonal studies. Humangenetik, 29:99-109. 7. Attal, J. (1969): Levels of testosterone, androstenedione, estrone and estradiol-173 in the testes of fetal sheep. Endocrinology, 85:280289.

/

21

8. Bardin, C. W., Bullock, L. P., Sherins, R. J., Mowszowisz, I., and Blackburn, W. R. (1973): Androgen metabolism and mechanism of action in male pseudohermaphroditism: A study of testicular fem¬ inization. Recent Prog. Horm. Res., 29:65-105. 9. Barr, M. L., and Bertram, L. F. (1949): A morphological distinction between neurones of the male and female and the behavior of the nucleolar satellite during accelerated nucleoprotein synthesis. Nature, 163:676-677. 10. Bengmark, S. (1958): The Prostatic Urethra and Prostate Glands. Berlingska Boktryckeriet, Lund, Sweden. 11. Berger, R., Abonyi, D., Nodot, A., Vialatte, J., and Lejeune, J. (1970): Hermaphrodisme vrai et “Garcon XX” dans une fratrie. Rev. Eur. Etud. Clin. Biol., 15:330-333. 12. Bidlingmaier, F., Knorr, D., and Neumann, F. (1977): Inhibition of masculine differentiation in male offspring of rabbits actively im¬ munized against testosterone before pregnancy. Nature, 266:647-648. 13. Billingham, R. E., and Silvers, W. K. (1960): Studies on tolerance of the Y chromosome antigen in mice. J. Immunol., 85:14-26. 14. Blanchard, M. G., and Josso, N. (1974): Source of the anti-mullerian hormone synthesized by the fetal testis: Mullerian-inhibiting activity of the fetal bovine Sertoli cells in tissue culture. Pediatr. Res., 8:968-971. 15. Blandau, R. J., White, B. J., and Rumery, R. E. (1963): Obser¬ vations on the movements of the living primordial germ cells in the mouse. Fertil. Steril., 14:482-489. 16. Bok, G., and Drews, U. (1983): The role of the wolffian ducts in the formation of the sinus vagina; an organ culture study. J. Embryol. Exp. Morphol., 73:275-295. 17. Bolliger, A. (1944): An experiment on the complete transformation of the scrotum into a marsupial pouch in Trichosurus vulpecula. Med. J. Aust., 2:56-58. 18. Book, J. A., Eilon, B., Halbrecht, I., Komlos, L., and Shabtay, F. (1973): Isochromosome Y [46,X,i(Yq)] and female phenotype. Clin. Genet., 4:410-414. 19. Boucekkine, C., Menasria, A., Choutier, A., Benelkadi, N., and Benmiloud, M. (1981): H-Y positive 46 XX true hermaphroditism with intrascrotal uterus. Clin. Endocrinol., 15:529-535. 20. Brook, C. G. D. (1981): Persistent mullerian duct syndrome. Pediatr. Adolesc. Endocrinol., 8:100-104. 21. Brook, C. G. D., Wagner, H., Zachmann, M., Prader, A., Armen¬ dares, S., Frenk, S., Abeman, P., Najjar, S. S., Slim, M. S.,Genton, N., andBozic, C. (1973): Familial occurrence of persistent mullerian structures in otherwise normal males. Br. Med. J., 1:771-773. 22. Brooks, J. R., Baptista, E. M., Berman, C., Ham, E. A., Hichens, M., Johnston, D. B. R., Primka, R. L., Rasmusson, G. H., Rey¬ nolds, G. F., Schmitt, S. M., and Arth, G. E. (1981): Response of rat ventral prostate to a new and novel 5a-reductase inhibitor. En¬ docrinology, 109:830-836. 23. Brooks, J. R., Berman, C., Hichens, M., Primka, R. L., Reynolds, G. F., and Rasmusson, G. H. (1982): Biological activities of a new steroidal inhibitor of A4-5a-reductase (41309). Proc. Soc. Exp. Biol. Med., 169:67-73. 24. Brown, T. R., Maes, M., Rothwell, S. W., and Migeon, C. J. (1982): Human complete androgen insensitivity with normal dihydrotestos¬ terone receptor binding capacity in cultured genital skin fibroblasts. Evidence for a qualitative abnormality of the receptor. J. Clin. En¬ docrinol. Metab., 55:61-69. 25. Bruner, J. A., and Witschi, E. (1946): Testosterone-induced mod¬ ifications of sex development in female hamsters. Am. J. Anat., 79:293-320. 26. Budzik, G. P., Powell, S. M., Kamagata, S., and Donahoe, P. K. (1983): Mullerian-inhibiting substance fractionation by dye affinity chromatography. Cell, 34:307-314. 27. Bullock, L. P., Bardin, C. W., and Ohno, S. (1971): The androgen insensitive mouse: Absence of intranuclear androgen retention in the kidney. Biochem. Biophys. Res. Commun., 44:1537-1543. 28. Bulmer, D. (1957): The development of the human vagina. J. Anat., 91:490-509. 29. Burgoyne, P. S. (1982): Genetic homology and crossing over in the X and Y chromosomes of mammals. Hum. Genet., 61:85-90. 30. Burgoyne, P. S. (1986): Mammalian X and Y crossover. Nature, 319:258-259. 31. Burgoyne, P. S., Levy, E. R., and McLaren, A. (1986): Spermatogenic failure in male mice lacking H-Y antigen. Nature, 320:170-172.

22 / Chapter 1 32. Bums, R. K. (1945): The differentiation of the phallus in the opossum and its reaction to sex hormones. Contrib. Embryol., 31:147-162. 33. Bums, R. K. (1961): Role of hormones in the differentiation of sex. In: Sex and Internal Secretions, edited by W. C. Young, pp. 76-158. Williams & Wilkins, Baltimore. 34. Buyse, A. (1935): The differentiation of transplanted mammalian gonad primordia. J. Exp. Zool., 70:1-41. 35. Cate, R. L., Mattaliano, R. J., Hession, C., Tizard, R., Faber, N. M., Cheung, A., Ninfa, E. G., Frey, A. Z., Gash, D. J., Chow, E. P., Fisher, R. A., Bertonis, J. M., Torres, G., Wallner, B. P., Ramachandran, K. L., Ragin, R. C., Manganaro, T. F., MacLaughlin, D. T., and Donahoe, P. K. (1986): Isolation of the bovine and human genes for mullerian inhibiting substance and expression of the human gene in animal cells. Cell, 45:685-698. 36. Catt, K. J., Dufau, M. L., Neaves, W. B., Walsh, P. C., and Wilson, J. D. (1975): LH-hCG receptors and testosterone content during differentiation of the testis in the rabbit embryo. Endocrinology, 97:1157-1165. 37. Cattanach, B. M., Evans, E. P., Burtenshaw, M. D., and Barlow, J. (1982): Male, female and intersex development in mice of identical chromosome constitution. Nature, 300:445-446. 38. Cattanach, B. M., Pollard, C. E., and Hawkes, S. G. (1971): Sexreversed mice XX and XO males. Cytogenetics, 10:318-337. 39. Clayton, G. W., Smith, J. D., and Rosenberg, H. S. (1958): Familial true hermaphroditism in pre- and postpubertal genetic females. Hor¬ monal and morphologic studies. J. Clin. Endocrinol. Me tab., 18:1349-1358. 40. Cooke, H. J., Brown, W. R. A., and Rappold, G. A. (1985): Hy¬ pervariable telomeric sequences from the human sex chromosomes are pseudoautosomal. Nature, 317:687-692. 41. Cooke, H. J., Fantes, J., and Green, D. (1983): Structure and ev¬ olution of human Y chromosome DNA. Differentiation, 23:S48-S55. 42. Crichton, D. N., and Steel, C. M. (1985): Serologically detectable H-Y (‘male’) antigen: Mr or myth? Immunol. Today, 6:202-203. 43. Cunha, G. R. (1972): Epithelial-mesenchymal interactions in pri¬ mordial gland structures which become responsive to androgenic stimulation. Anat. Rec., 172:179-196. 44. Cunha, G. R. (1972): Tissue interactions between epithelium and mesenchyme of urogenital and integumental origin. Anat. Rec., 172:529-542. 45. Cunha, G. R., and Chung, L. W. K. (1981): Stromal-epithelial interactions. I. Induction of prostatic phenotype in urothelium of testicular feminized (Tfm/Y) mice. J. SteroidBiochem., 14:1317-1321. 46. Cunha, G. R., Chung, L. W. K., Shannon, J. M., and Reese, B. A. (1980): Stromal-epithelial interactions in sex differentiation. Biol. Reprod., 22:19—42. 47. Davis, R. M. (1981): Localisation of male determining factors in man: A thorough review of structural anomalies of the Y chromo¬ some. J. Med. Genet., 18:161-195. 48. Dean, D. C., Gope, R., Knoll, B. J., Riser, M. E., and O’Malley, B. W. (1984): A similar 5'-flanking region is required for estrogen and progesterone induction of ovalbumin gene expression. J. Biol. Chem., 259:9967-9970. 49. De la Chapelle, A. (1972): Nature and origin of males with XX sex chromosomes. Am. J. Hum. Genet., 24:71-105. 50. De la Chapelle, A. (1981): The etiology of maleness in XX men. Hum. Genet., 58:105-116. 51. De la Chapelle, A., Tippett, P. A., Wetterstrand, G., and Page, D. (1984): Genetic evidence of X-Y interchange in a human XX male. Nature, 307:170-171. 52. Dickmann, Z., and Dey, S. K. (1976): A new concept: Control of early pregnancy by steroid hormones originating in the preimplan¬ tation embryo. Vitam. Horm., 34:215-242. 53. Dickmann, Z., Gupta, J. S., and Dey, S. K. (1977): Does “blastocyst estrogen” initiate implantation? Science, 195:687-688. 54. Donahoe, P. K., Budzik, G. P., Trelstad, R., Mudgett-Hunter, M., Fuller, A. Jr., Hutson, J. M., Ikawa, H., Hayashi, A., and MacLaughlin, D. (1982): Mullerian-inhibiting substance: An update. Recent Prog. Horm. Res., 38:279-326. 55. Donahoe, P. K., Fuller, A. F. Jr., Sailly, R. E., Guy, S. R., and Budzik, G. P. (1981): Mullerian inhibiting substance inhibits growth of a human ovarian cancer in nude mice. Ann. Surg., 194:472-480. 56. Donahoe, P. K., Ho, Y., Morikawa, Y., and Hendren, W. H. (1977): Mullerian inhibiting substance in human testes after birth. J. Pediatr. Surg., 12:323-330.

57. Donahoe, P. K., Hutson, J. M., Fallat, M. E., Kamagata, S., and Budzik, G. P. (1984): Mechanism of action of mullerian inhibiting substance. Annu. Rev. Physiol., 46:53-65. 58. Donahoe, P. K., Ito, Y., Price, J. M., and Herndon, W. H. Ill (1977): Mullerian inhibiting substance activity in bovine fetal, new¬ born and prepubertal testes. Biol. Reprod., 16:238-243. 59. Donahoe, P. K., Swann, D. A., Hayashi, A., and Sullivan, M. D. (1979): Mullerian duct regression in the embryo correlated with cy¬ totoxic activity against human ovarian cancer. Science, 205:913-915. 60. Donovan, P. J., Stott, D., Cairns, L. A., Heasman, J., and Wylie, C. C. (1986): Migratory and postmigratory mouse primordial germ cells behave differently in culture. Cell, 44:831-838. 61. Drews, U., and Drews, U. (1977): Regression of mouse mammary gland anlagen in recombinants of Tfm and wild-type tissues: Tes¬ tosterone acts via the mesenchyme. Cell, 10:401-404. 62. Dyche, W. J. (1979): A comparative study of the differentiation and involution of the mullerian duct and wolffian duct in the male and female fetal mouse. J. Morphol., 162:175-210. 63. Eicher, E. M., and Washburn, L. L. (1983): Inherited sex reversal in mice: Identification of a new primary sex-determining gene. J. Exp. Zool., 228:297-304. 64. Eicher, E. M., and Washburn, L. L. (1986): Genetic control of primary sex determination in mice. Annu. Rev. Genet., 20:327-360. 65. ‘“Eicher, E. M., Washburn, L. L., Whitney, J. B. Ill, and Morrow, K. E. (1982): Mus poschiavinus Y chromosome in the C57BL/6J murine genome causes sex reversal. Science, 217:535-537. 66. Eichwald, E. J., and Silmser, C. R. (1955): Untitled communication. Transplant. Bull., 2:148-149. 67. Eli, C. (1983): Familial incomplete male pseudohermaphroditism associated with impaired nuclear androgen retention. J. Clin. Invest., 71:850-858. 68. Elawady, M. K., Allman, D. R., Griffin, J. E., and Wilson, J. D. (1983): Expression of a mutant androgen receptor in cloned fibro¬ blasts derived from a heterozygous carrier for the syndrome of tes¬ ticular feminization. Am. J. Hum. Genet., 35:376-384. 69. Epplen, J. T., Cellini, A., Shorte, M., and Ohno, S. (1983): On evolutionarily conserved simple repetitive DNA sequences: Do “sexspecific” satellite components serve any sequence dependent func¬ tion? Differentiation, 23:S60-S63. 70. Epplen, J. T., McCarrey, J. R., Sutou, S., and Ohno, S. (1982): Base sequence of a cloned snake W-chromosome DNA fragment and identification of a male-specific putative mRNA in the mouse. Proc. Natl. Acad. Sci. USA, 79:2798-3802. 71. Evans, E. P., Burtenshaw, M. D., and Cattanach, B. H. (1982): Meiotic crossing-over between the X and Y chromosomes of male mice carrying the sex-reversing (Sxr) factor. Nature, 300:443-445. 72. Evens, R. P., and Amerson, A. B. (1974): Androgens and erythropoiesis. J. Clin. Pharmacol. 14:94-101. 73. Fadern, B. H., and Tesoriero, J. V. (1986): Inhibition of testicular development and feminization of the male genitalia by neonatal es¬ trogen treatment in a marsupial. Biol. Reprod. 34:771-776. 74. Fallat, M. E., Hutson, J. M., Budzik, D. P., and Donahoe, P. K. (1984): Androgen stimulation of nucleotide pyrophosphatase during mullerian duct regression. Endocrinology, 114:1592-1598. 75. Fisher, L. K., Kogut, M. D., Moore, R. J., Goebelsmann, U., Weitzmann, J. J., Isaacs, H. Jr., Griffin, J. E., and Wilson, J. D. (1978): Clinical, endocrinological, and enzymatic characterization of two patients with 5a-reductase deficiency. Evidence that a single enzyme is responsible for the 5a-reduction of cortisol and testoster¬ one. J. Clin. Endocrinol., 47:653-664. 76. Forejt, J. (1979): Meiotic studies of translocations causing male sterility in the mouse. II. Double heterozygotes for Robertsonian translocations. Cytogenet. Cell Genet., 21:163-170. 77. Forejt, J., and Gregorova, S. (1977): Meiotic studies of translocations causing male sterility in the mouse. I. Autosomal reciprocal trans¬ location. Cytogenet. Cell Genet., 19:159-179. 78. Fujimoto, T., Miyayama, Y., and Fuyuta, M. (1977): The origin, migration and fine morphology of human primordial germ cells Anat Rec., 188:315-330. 79. Fuller, A. F. Jr., Guy, S., Budzik, G. P., and Donahoe, P. K. (1982): Mullerian-inhibiting substance inhibits colony growth of a human ovarian carcinoma cell line. J. Clin. Endocrinol. Metab 54:1051-1055. 80. Gallegos, A. J., Guizar, E., Cortes-Gallegos, V., Cervantes, C., Bedolla, N., and Parra, A. (1976): Familial true hermaphrodism in

Sex Determination and Differentiation

81.

82.

83. 84.

85. 86.

87. 88. 89.

90.

91.

92. 93.

94.

95.

96.

97.

98.

99.

100. 101. 102. 103.

104.

105.

106.

three siblings: Plasma hormonal profile and in vitro steroid biosyn¬ thesis in gonadal structures. J. Clin. Endocrinol. Metab., 42:653-660. Gardner, R. L., Lyon, M. F., Evans, E. P., and Burtenshaw, M. D. (1985): Clonal analysis of X-chromosome inactivation and the origin of the germ line in the mouse embryo. J. Embryol. Exp. Morphol., 88:349-363. Gartier, S. M., Liskay, R. M., Campbell, B. K., Sparkes, R., and Gant, N. (1972): Evidence for two functional X chromosomes in human oocytes. Cell Differ. 1:215-218. Gartler, S. M., Liskay, R. M., and Gant, N. (1973): Two functional X chromosomes in human fetal oocytes. Exp. Cell Res., 82:464-465. Gehring, U., Tomkins, G. M., and Ohno, S. (1971): Effect of the androgen-insensitivity mutation on a cytoplasmic receptor for di¬ hydrotestosterone. Nature (New Biol.), 232:106-107. George, F. W., and Noble, J. F. (1984): Androgen receptors are similar in fetal and adult rabbits. Endocrinology, 115:1451-1458. George, F. W., and Wilson, J. D. (1978): Conversion of androgen to estrogen by the human fetal ovary. J. Clin. Endocrinol. Metab., 47:550-555. George, F. W., and Wilson, J. D. (1978): Estrogen formation in the early rabbit embryo. Science, 199:200-202. George, F. W., and Wilson, J. D. (1980): Endocrine differentiation of the fetal rabbit ovary in culture. Nature, 283:861-863. George, F. W., Catt, K. J., Neaves, W. B., and Wilson, J. D. (1978): Studies on the regulation of testosterone synthesis in the rabbit fetal testis. Endocrinology, 102:106-107. George, F. W., Simpson, E. R., Milewich, L., and Wilson, J. D. (1979): Studies on the regulation of steroid hormone biosynthesis in fetal rabbit gonads. Endocrinology, 105:1100-1106. George, F. W., Tobleman, W. T., Milewich, L., and Wilson, J. D. (1978): Aromatase activity in the developing rabbit brain. Endocri¬ nology, 102:86-91. Gier, H. T., and Marion, G. B. (1969): Development of the mam¬ malian testis and genital ducts. Biol. Reprod., 1:1-23. Gillman, J. (1948): The development of the gonads in man, with a consideration of the role of fetal endocrines and the histogenesis of ovarian tumors. Carnegie Contrib. Embryol., 32:83-131. Goldberg, E. H., Boyse, E. A., Bennett, D., Scheid, M., and Cars¬ well, E. A. (1971): Serological demonstration of H-Y (male) antigen on mouse sperm. Nature, 232:478-480. Goldman, A. S. (1971): Production of hypospadias in the rat by selective inhibition of fetal testicular 17a-hydroxyla.se and Ci7.2olyase. Endocrinology, 88:527-531. Goldman, A. S., Shapiro, B. H., and Neuman, F. (1976): Role of testosterone and its metabolites in the differentiation of the mammary gland in rats. Endocrinology, 99:1490-1495. Goldstein, J. L., and Wilson, J. D. (1972): Studies on the patho¬ genesis of the pseudohermaphroditism in the mouse with testicular feminization. J. Clin. Invest., 51:1647-1658. Gondos, B. (1980): Development and differentiation of the testis and male reproductive tract. In: Testicular Development, Structure, and Function, edited by A. Steinberger and E. Steinberger, pp. 3-20. Raven Press, New York. Gondos, B., George, F. W., and Wilson, J. D. (1983): Granulosa cell differentiation and estrogen synthesis in the fetal rabbit ovary. Biol. Reprod., 29:791-798. Goodfellow, P., Darling, S., and Wolfe, J. (1985): The human Y chromosome. J. Med. Genet., 22:329-344. Goodfellow, P. J., Darling, S. M., Thomas, N. S., and Goodfellow, P. N. (1986): A pseudoautosomal gene in man. Science, 234:740-743. Gordon, J. W., and Ruddle, F. H. (1981): Mammalian gonadal determination and gametogenesis. Science, 211:1265-1271. Gore-Langston, R. E., Tung, P. S., and Fritz, I. B. (1983): The absence of specific interaction of Sertoli-cell-secreted proteins with antibodies directed against FI-Y antigen. Cell, 32:289-301. Greene, R. R. (1942): Hormonal factors in sex inversion: The effects of sex hormones on embryonic sexual structures of the rat. Biol. Symp., 9:105-123. Griffin, J. E. (1979): Testicular feminization associated with a thermolabile androgen receptor in cultured human fibroblasts. J. Clin. Invest., 64:1624-1631. Griffin, J. E., and Durrant, J. L. (1982): Qualitative receptor defects in families with androgen resistance: Failure of stabilization of the fibroblast cytosol androgen receptor. J. Clin. Endocrinol. Metab., 55:465-474.

/

23

107. Griffin, J. E., and Wilson, J. D. (1978): Hereditary male pseudo¬ hermaphroditism. Clin. Obstet. Gynaecol., 5:457-479. 108. Griffin, J. E., Kovacs, W. J., and Wilson, J. D. (1985): Charac¬ teristics of androgen resistance. In: Regulation of Androgen Action, edited by N. Bruchovsky, A. Chapdelaine, and F. Neumann, The Proceedings of an International Symposium, pp. 127-131. Congressdruck R. Bruckner, Berlin. 109. Griffin, J. E., Punyashthiti, K., and Wilson, J. D. (1976): Dihy¬ drotestosterone binding by cultured human fibroblasts. Comparison of cells from control subjects and from patients with hereditary male pseudohermaphroditism due to androgen resistance. J. Clin. Invest., 57:1342-1351. 110. Gruenwald, P. (1941): The relation of the growing mullerian duct to the wolffian duct and its importance for the genesis of malfor¬ mation. Anat. Rec., 81:1-19. 111. Guellaen, G., Casanova, M., Bishop, C., Geldwerth, D., Andre, G., Fellous, M., and Weissenbach, J. (1984): Human XX males with Y single-copy DNA fragments. Nature, 307:172-173. 112. Handel, M. A., and Eppig, J. J. (1979): Sertoli cell differentiation in the testes of mice genetically deficient in germ cells. Biol. Reprod., 20:1031-1038. 113. Hayashi, A., Donahoe, P. K., Budzik, G. P., and Trelstad, R. L. (1982): Periductal and matrix glycosaminoglycans in rat mullerian duct regression. Dev. Biol., 92:16-26. 114. Hertig, A. T., Adams, E. C., McKay, D. G., Rock, J., Mulligan, W. J., and Menkin, M. (1956): A description of 34 human ova within the first 17 days of development. Am. J. Anat., 98:435-493. 115. Heuberger, B., Fritzica, I., Wasner, G., and Kratochwil, K. X1982): Induction of androgen receptor formation by epithelium-mesenchyme interaction in embryonic mouse mammary gland. Proc. Natl. Acad. Sci. USA, 79:2957-2961. 116. Holyoke, E. A. (1949): The differentiation of embryonic gonads transplanted to the adult omentum in the albino rat. Anat. Rec., 103:675-699. 117. Huhtaniemi, I. T., Korenbrat, C. C., and Jaffe, R. B. (1977): hCG binding and stimulation of testosterone biosynthesis in the human fetal testis. J. Clin. Endocrinol. Metab., 44:963-967. 118. Hutson, J. M. (1985): A biphasic model for the hormonal control of testicular descent. Lancet, 2:419-421. 119. Hutson, J. M., and Donahoe, P. K. (1983): Is mullerian-inhibiting substance a circulating hormone in the chick-quail chimera? Endo¬ crinology, 113:1470-1475. 120. Hutson, J. M., Donahoe, P. K., and MacLaughlin, D. T. (1985): Steroid modulation of mullerian duct regression in the chick embryo. Gen. Comp. Endocrinol., 57:88-102. 121. Hutson, J. M., Fallat, M. E., Kamagata, S., Donahoe, P. K., and Budzik, G. P. (1984): Phosphorylation events during mullerian duct regression. Science, 223:586-588. 122. Hutson, J. M., Ikawa, H., and Donahoe, P. K. (1982): Estrogen inhibition of mullerian inhibiting substance in the chick embryo. J. Pediatr. Surg., 17:953-959. 123. Ikadai, H., Sakuma, Y., Suzuki, K., and Imamichi, T. (1985): Congenital abnormalities of the male genital organs in the newly established TW rat strain. Cong. Anom., 26:65-71. 124. Ikawa, H., Hutson, J. M., Budzik, D. P., MacLaughlin, D. T., and Donahoe, P. K. (1982): Steroid enhancement of mullerian duct regression. J. Pediatr. Surg. 17:453-458. 125. Imperato-McGinley, J., Binienda, Z., Arthur, A., Mininberg, D. T., Vaughan, D. Jr., and Quimby, F. W. (1985): The develop¬ ment of a male pseudohermaphroditic rat using an inhibitor of the en¬ zyme 5a-reductase. Endocrinology, 116:807-812. 126. Imperato-McGinley, J., Binienda, Z., Gedney, J., and Vaughan, E. D. (1986): Nipple differentiation in fetal male rats treated with an inhibitor of the enzyme 5a-reductase: Definition of a selective role for dihydrotestosterone. Endocrinology, 118:132-137. 127. Imperato-McGinley, J., Guerrero, L., Gautier, T., and Peterson, R. E. (1974): Steroid 5a-reductase deficiency in man: An inherited form of male pseudohermaphroditism. Science, 186:1213-1215. 128. Imperato-McGinley, J., Peterson, R. E., Leshin, M., Griffin, J. E., Cooper, G., Draghi, S., Berenyi, M., and Wilson, J. D. (1980): Ste¬ roid 5a-reductase deficiency in a 65 year old pseudohermaphrodite: The natural history, ultrastructure of the testis and evidence for inherited enzyme heterogeneity. J. Clin. Endocrinol. Metab., 50:15-22. 129. Jacobs, P. A., and Ross, A. (1966): Structural abnormalities of the Y chromosome in man. Nature, 210:352-354.

24

/ Chapter

1

130. Jirasek, J. E. (1976): Principles of reproductive embryology. In: Disorders of Sexual Differentiation, edited by J. L. Simpson, pp. 51-110. Academic Press, New York. 131. Jones, H. W. Jr., Rary, J. M., Rock, J. A., and Cummings, D. (1979): The role of H-Y antigen in human sexual development. Johns Hopkins Med. J., 145:33-43. 132. Jones, K. W. (1983): Evolutionary conservation of sex specific DNA sequences. Differentiation, 23:S56-S59. 133. Josso, N., Fekete, C., Cachin, O., Nezelof, C., and Rappaport, R. (1983): Persistence of mullerian ducts in male pseudohermaphrodit¬ ism, and its relationship to cryptorchidism. Clin. Endocrinol., 19:247-258. 134. Josso, N., Picard, J.-Y., and Tran, D. (1977): The anti Mullerian hormone. Recent Prog. Horm. Res., 33:117-167. 135. Jost, A. (1947): Recherches sur la differenciation de l’embryon de lapin. II. Action des androgenes synthese sur sur l’histogenses genitale. Arch. Anat. Microsc. Morphol. Exp., 36:242-270. 136. Jost, A. (1953): Problems in fetal endocrinology: The gonadal and hypophyseal hormones. Recent Prog. Horm. Res., 8:379418. 137. Jost, A. (1961): The role of fetal hormones in prenatal development. Harvey Lect., 55:201-226. 138. Jost, A. (1972): A new look at the mechanisms controlling sexual differentiation in mammals. Johns Hopkins Med. J., 130:38-53. 139. Jost, A., and Magre, S. (1984): Testicular development phases and dual hormonal control of sexual organogenesis. In: Sexual Differ¬ entiation: Basic and Clinical Aspects, edited by M. Serio et al., pp. 1-15. Raven Press, New York. 140. Kasdan, R., Nankin, H. R., Troen, P., Wald, N., Pan, S., and Yanaihara, T. (1973): Paternal transmission of maleness in XX hu¬ man beings. N. Engl. J. Med., 288:539-545. 141. Kaufman, M., Pinsky, L., Hollander, R., and Bailey, J. D. (1983): Regulation of the androgen in normal and androgen resistant genital skin fibroblasts. J. Steroid Biochem., 18:383-390. 142. Keenan, B. S., Meyer, W. J. Ill, Hadjian, A. J., Jones, H. W., and Migeon, C. J. (1974): Syndrome of androgen insensitivity in man: Absence of 5a-dihydrotestosterone binding protein in skin fibro¬ blasts. J. Clin. Endocrinol. Metab., 38:1143-1146. 143. Kellokumpo-Lehtinen, P., Santti, R., and Pelliniemi, L. J. (1980): Correlation of early cytodifferentiation of the human fetal prostate and Leydig cells. Anat. Rec., 196:263-273. 144. Kobayashi, S. (1984): Induction of mullerian duct derivatives in testicular feminized (Tfm) mice by prenatal exposure to diethylstilbestrol. Anat. Embryol. 169:35-39. 145. Kovacs, W. J., Griffin, J. E., and Wilson, J. D. (1983): Transfor¬ mation of human androgen receptors to the deoxyribonucleic acid¬ binding state. Endocrinology, 113:1574—1581. 146. Kovacs, W. J., Griffin, J. E., Weaver, D. D., Carlson, B. R., and Wilson, J. D. (1984): A mutation that causes lability of the androgen receptor under conditions that normally promote DNA-binding state. J. Clin. Invest., 73:1095-1104. 147. Kratochwil, K. (1971): In vitro analysis of the hormonal basis for the sexual dimorphism in the embryonic development of the mouse mammary gland. Embryol. Exp. Morphol., 25:141-153. 148. Kratochwil, K., and Schwartz, P. (1976): Tissue interaction in an¬ drogen response of embryonic mammary rudiment of mouse: Iden¬ tification of target tissue for testosterone. Proc. Natl. Acad. Sci. USA, 73:4041-4044. 149. Kratzer, P. G., and Chapman, V. M. (1981): X chromosome reac¬ tivation in oocytes of Mus caroli. Proc. Natl. Acad. Sci. USA, 78:20932097. 150. Lasnitzki, I., and Mizuno, T. (1980): Prostatic induction: Interaction of epithelium and mesenchyme from normal wild-type and androgen insensitive mice with testicular feminization. J. Endocrinol., 85:423-428. 151. Leshin, M., Griffin, J. E., and Wilson, J. D. (1978): Hereditary male pseudohermaphroditism associated with an unstable form of 5a-reductase. J. Clin. Invest., 62:685-691. 152. Liang, T., and Heiss, C. E. (1981): Inhibition of 5a-reductase, receptor binding, and nuclear uptake of androgens in the prostate by a 4-methyl-4-aza-steroid. J. Biol. Chem., 256:7998-8005. 153. Lifschytz, E. (1971): X-chromosome inactivation: An essential fea¬ ture of normal spermiogenesis in male heterogametic organisms. In: The Genetics of the Spermatozoon, edited by R. A. Beatty and S.

Gluecksohn-Waelsch, pp. 223-232. Bogtrykkenet Forum, Copen¬ hagen. 154. Lifschytz, E., and Lindsley, D. L. (1972): The role of X-chromosome inactivation during spermatogenesis. Proc. Natl. Acad. Sci.. USA, 69:182-186. 155. Lipsett, M. B., and Tullner, W. W. (1965): Testosterone synthesis by the fetal rabbit gonad. Endocrinology, 77:273-277. 156. Lowry, R. B., Honore, L. H., Arnold, W. J. D., Johnson, H. W., Kliman, M. R., and Marhsall, R. H. (1975): Familial true herma¬ phroditism. Birth Defects, 11:105-113. 157. Lowsley, O. S. (1912): The development of the human prostate with reference to the development of other structures of the neck of the urinary bladder. Am. J. Anat., 13:299-349. 158. Lyon, M. F. (1961): Gene action in the X-chromosome of the mouse. Nature, 190:372-373. 159. Lyon, M. F. (1962): Sex chromatin and gene action in the mammalian X-chromosome. Am. J. Hum. Genet., 14:135-148. 160. Lyon, M. F. (1968): Chromosomal and sub-chromosomal inactiva¬ tion. Annu. Rev. Genet., 2:31-52. 161. Lyon, M. F., and Hawkes, S. G. (1970): X-Linked gene for testicular feminization in the mouse. Nature, 227:1217-1219. 162. MacDonald, P. C., Madden, J. D., Brenner, P. F., Wilson, J. D., and Siiteri, P. K. (1979): Origin of estrogen in normal men and in “women with testicular feminization. J. Clin. Endocrinol. Metab., 49:905-916. 163. Madan, K. (1983): Balanced structural changes involving the human X: Effect on sexual phenotype. Hum. Genet., 63:216-221. 164. Madden, J. D., Walsh, P. C., MacDonald, P. C., and Wilson, J. D. (1975): Clinical and endocrinological characterization of a pa¬ tient with the syndrome of incomplete testicular feminization. J. Clin. Endocrinol. Metab., 40:751-760. 165. Maes, M., Sultan, C., Zerhouni, N., Rothwell, S. W., and Migeon, C. J. (1979): Role of testosterone binding to the androgen receptor in male sexual differentiation of patients with 5a-reductase defi¬ ciency. J. Steroid Biochem., 11:1385-1390. 166. Magre, S., and Jost, A. (1984): Dissociation between testicular or¬ ganogenesis and endocrine cytodifferentiation of Sertoli cells. Proc. Natl. Acad. Sci. USA, 81:7831-7834. 167. Mangoushi, M. A. (1975): Scrotal allografts of fetal ovaries. J. Anat., 120:595-599. 168. McCarrey, J. R., and Abbott, U. K. (1978): Chick gonad differ¬ entiation following excision of primordial germ cells. Dev. Biol., 66:256-265. 169. McKay, D. G., Hertig, A. T., Adams, E. C., and Danziger, S. (1953): Histochemical observation on the germ cells of the human embryos. Anat. Rec., 117:201-220. 170. McLachlan, J. A. (1977): Prenatal exposure to diethylstilbestrol in mice: Toxicological studies. J. Toxicol. Environ. Health, 2:527-537. 171. McLaren, A. (1983): Sex reversal in the mouse. Differentiation 23:S93-S98. 172. McLaren, A., and Monk, M. (1981): X chromosome activity in the germ cells of sex-reversed mouse embryos. J. Reprod. Fertil 63:533-537. 173. McLaren, A., and Monk, M. (1982): Fertile females produced by inactivation of an X chromosome of ‘sex-reversed’ mice Nature 300:446-448. 174. McLaren, A., Simpson, E. Tomonari, K., Chandler, P., and Hogg, H. (1984): Male sexual differentiation in mice lacking H-Y antigen. Nature, 312:552-555. 175. Merchant, H. (1975): Rat gonadal and ovarian organogenesis with and without germ cells. An ultrastructural study. Dev. Biol., 44:1-21. 176. Meyer, W. J. Ill, Migeon, B. R., and Migeon, C. J. (1975): Locus on human X chromosome for dihydrotesterone receptor and androgen insensitivity. Proc. Natl. Acad. Sci. USA, 72:1469-1472. 177. Migeon, B. R., Brown, T. R., Axelman, J., and Migeon, C. J. (1981): Studies of the locus for androgen receptor: Localization on the human X chromosome and evidence for homology with the Tfm locus in the mouse. Proc. Natl. Acad. Sci. USA, 78:6339-6343. 178. Migeon, B. R., and Jelalian, K. (1977): Evidence for two active X chromosomes in germ cells of female before meiotic entry Nature 269:242-243. 179. Milewich, L., George, F. W., and Wilson, J. D. (1977): Estrogen formation by the ovary of the rabbit embryo. Endocrinology 100:187-196.

Sex Determination and Differentiation / 180. Molsberry, R. L., Carr, B. R., Mendelson, C. R., and Simpson, E. R. (1982): Human chorionic gonadotropin binding to human fetal testes as a function of gestation age. J. Clin. Endocrinol. Metab., 55:791-794. 181. Monesi, V. (1965): Synthetic activities during spermatogenesis in the mouse. RNA and protein. Exp. Cell Res., 39:197-224. 182. Moore, C. R., and Price, D. (1942): Differentiation of embryonic gonads transplanted into post-natal hosts. J. Exp. Zool., 90:229-265. 183. Moore, R. J., and Wilson, J. D. (1976): Steroid 5a-reductase in cultured human fibroblasts: Biochemical and genetic evidence for two enzyme activities. J. Biol. Chem., 251:5895-5900. 184. Moore, R. J., Griffin, J. E., and Wilson, J. D. (1975): Diminished 5a-reductase activity in extracts of fibroblasts cultured from patients with familial incomplete male pseudohermaphroditism, type 2. J. Biol. Chem., 250:7168-7172. 185. Mori, Y., and Mitzutani, S. (1968): Familial true hermaphroditism in genetic females. Jpn. J. Urol., 59:857-864. 186. Mudgett-Hunter, M., Budzik, G. P., Sullivan, M., and Donahoe, P. K. (1982): Monoclonal antibody to mullerian inhibiting substance. J. Immunol., 128:1327-1333. 187. Neumann, F., von Berswordt-Wallrabe, R., Eiger, W., Steinbeck, H., Hahn, J. D., and Kramer, M. (1970): Aspects of androgendependent events as studied by antiandrogens. Recent Prog. Horm. Res., 26:337-405. 188. New, M. I., Dupont, B., Grunback, K., and Levine, L. S. (1983): Congenital adrenal hyperplasia and related conditions. In: The Met¬ abolic Basis of Inherited Disease, 5th ed., edited by J. B. Stanbury, J. B. Wyngaarden, D. S. Fredrickson, J. L. Goldstein, and M. S. Brown, pp. 973-1000. McGraw-Hill, New York. 189. Odartchenko, N., and Pavillard, M. (1970): Late DNA replication in male mouse meiotic chromosomes. Science, 167:1133-1134. 190. Ohno, S. (1978): The role of H-Y antigen in primary sex determi¬ nation. J. Am. Med. Assoc., 239:217-220. 191. Ohno, S., and Lyon, M. F. (1965): Cytological study of Searle’s Xautosome translocation in Mus musculus. Chromosoma, 16:90-100. 192. Ohno, S., and Makino, S. (1961): The single-X nature of sex chro¬ matin in man. Lancet, 1:78-79. 193. Ohno, S., Jainchill, J., and Stenius, C. (1963): The creeping vole (Microtus oregoni) as a gonosomic mosaic. The OY/XY constitution of the male. Cytogenetics, 2:232-239. 194. O’Rahilly, R. (1977): The development of the vagina in the human. In: Morphogenesis and Malformation of the Genital System, edited by R. J. Blandua and D. Bergsma. Birth Defects, 13:123-136. 195. Page, D., de Martinville, B., Barker, D., Wyman, A., White, R., Francke, U., and Botstein, D. (1982): Single-copy sequence hy¬ bridizes to polymorphic and homologous loci on human X and Y chromosomes. Proc. Natl. Acad. Sci. USA, 79:5352-5356. 196. Patsavoudi, E., Magre, S., Castanier, M., Scholler, R., and Jost, A. (1985): Dissociation between testicular morphogenesis and func¬ tional differentiation of Leydig cells. J. Endocrinol., 105:235-238. 197. Perez-Palacios, G., Medina, M., Ullao-Aguirre, A., Chavez, B. A., Villareal, G., Dutrem, M. T., Cahill, L. T., and Wachtel, S. (1981): Gonadotropin dynamics in XX males. J. Clin. Endocrinol. Metab., 53:254-257. 198. Peterson, R. E., Imperato-McGinley, J., Gautier, T., and Sturla, E. (1977): Male pseudohermaphroditism due to steroid 5a-reductase deficiency. Am. J. Med., 62:170-191. 199. Pfahl, M. (1982): Specific binding of the glucocorticoid-receptor complex to the mouse mammary tumor proviral promotor region. Cell, 31:475-482. 200. Pfaitz, C. R. (1949): Das embryonale und postnatale Verholten der mannlichen Brust driise beim menschen. II. Das mammarorgan im Kindes-, Jiinglings-, Mannes-und Greisenalter. Acta Anat., 8:293328. 201. Picard, J. Y., Benarous, R., Guerrier, D., Josso, N., and Kahn, A. (1986): Cloning and expression of cDNA for anti-Mullerian hormone. Proc. Natl. Acad. Sci. USA, 83:5464-5468. 202. Picard, J. Y., Tran, D., and Josso, N. (1978): Biosynthesis of la¬ belled anti-mullerian hormone by fetal testes: Evidence for the gly¬ coprotein nature of the hormone and for its disulfide-bonded structure. Mol. Cell. Endocrinol., 12:17-30. 203. Price, J. M. (1979): The secretion of mullerian inhibiting substance by cultured isolated Sertoli cells of the neonatal calf. Am. J. Anat., 156:147-157.

25

204. Price, J. M., Donahoe, P. K., Ito, Y., and Hendren, W. H. Ill (1977): Programmed cell death in the mullerian duct induced by mullerian inhibiting substance. Am. J. Anat., 149:353-376. 205. Price, P., Wass, J. A. H., Griffin, J. E., Leshin, M., Savage, M. O., Large, D. M., Bu’Lock, D. E., Anderson, D. C., Wilson, J. D., and Besser, G. M. (1984): High dose androgen therapy in male pseudohermaphroditism due to 5a-reductase deficiency and dis¬ orders of the androgen receptor. J. Clin. Invest., 74:1496-1508. 206. Rajfer, J., and Walsh, P. C. (1977): Hormonal regulation of testicular descent: Experimental and clinical observations. J. Urol., 118:985-990. 207. Rasmussen, S. W., and Holm, P. B. (1980): Mechanics of meiosis. Hereditas, 93:187-216. 208. Rigaudiere, N. (1979): The androgens in the guinea-pig foetus throughout the embryonic development. Acta Endocrinol., 92:174-186. 209. Roe, T. F., and Alfi, O. S. (1977): Ambiguous genitalia in XX male children: Report of two infants. Pediatrics, 60:55-59. 210. Rosenberg, H. S., Clayton, G. W., and Hsu, T. C. (1963): Familial true hermaphrodism. J. Clin. Endocrinol. Metab., 23:203-206. 211. Rouyer, F., Simmler, M. C., Johnsson, C., Vergnaud, G., Cooke, H. J., and Weissenbach, J. (1986): A gradient of sex linkage in the pseudoautosomal region of the human sex chromosomes. Nature, 319:291-295. 212. Russell, L. B., and Montgomery, C. S. (1969): Comparative studies on X-autosome translocations in the mouse. I. Origin, viability, fertility, and weight of five T(X,I)’S. Genetics, 63:103-120. 213. Saenger, P., Levine, L. S., Wachtel, S. S., Korth-Schutz, S., Doberne, Y., Koo, G. C., Lavengood, R. W. Jr., German, J. L. Ill, and New, M. I. (1976): Presence of H-Y antigen and testis in 46,XX true hermaphroditism, evidence of Y-chromosomal function. J. Clin. Endocrinol. Metab., 43:1234-1239. 214. Schultz, F. M., and Wilson, J. D. (1974): Virilization of the wolffian duct in the rat fetus by various androgens. Endocrinology, 94:979-986. 215. Schweikert, H. U., Weissbach, L., Leyendecker, G., Schwinger, E., Wartenberg, H., and Kruck, F. (1982): Clinical, endocrinolog¬ ical, and cytological characterization of two 46,XX males. J. Clin. Endocrinol. Metab., 54:745-752. 216. Searle, A. G. (1962): Is sex-linked Tabby really recessive in the mouse? Heredity, 17:297. 217. Selden, J. R., Wachtel, S. S., Koo, G. C., Haskins, M. E., and Patterson, D. F. (1978): Genetic basis of XX male syndrome and XX true hermaphroditism: Evidence in the dog. Science, 201:644—646. 218. Siiteri, P. K., and Wilson, J. D. (1974): Testosterone formation and metabolism during male sexual differentiation in the human embryo. J. Clin. Endocrinol. Metab., 38:113-125. 219. Silvers, W. K., Gasser, D. L., and Eicher, E. M. (1982): H-Y antigen, serologically detectable male antigen and sex determination. Cell, 28:439-440. 220. Simmler, M. C., Rouyer, F., Vergnaud, G., Nystrom-Lahti, M., Ngo, K. Y., de la Chapelle, A., and Weissenbach, J. (1985): Pseu¬ doautosomal DNA sequences in the pairing region of the human sex chromosomes. Nature, 317:692-697. 221. Simpson, J. L. (1978): Tme hermaphroditism: Etiology and phe¬ notypic considerations. Birth Defects, 14:9-35. 222. Singh, L., and Jones, K. W. (1982): Sex reversal in the mouse (Mus musculus) is caused by a recurrent non-reciprocal crossover involving the X and the aberrant Y chromosome. Cell, 28:205-216. 223. Singh, L., Purdom, I. F., and Jones, K. W. (1980): Sex chromosome associated satellite DNA: Evolution and conservation. Chromosoma 79:137-157. 224. Sloan, W. R., and Walsh, P. C. (1976): Familial persistent mullerian duct syndrome. J. Urol., 115:459-461. 225. Solari, A. J. (1974): The behavior of the XY pair in mammals. Int. Rev. Cytol., 38:273-317. 226. Spaulding, M. H. (1921): The development of the external genitalia in the human embryo. Carnegie Contrib. Embryol., 13:67. 227. Stem, C. (1957): The problem of complete Y-linkage in man. Am. J. Hum. Genet., 9:147-165. 228. Suzuki, Y., Ishii, H., Fumya, H., and Arai, Y. (1982): Develop¬ mental changes of the hypogastric ganglion associated with the dif¬ ferentiation of the reproductive tracts in the mouse. Neurosci. Lett., 32:271-276. 229. Takeda, H., Mizuno, T., and Lasnitzki, I. (1985): Autoradiographic studies of androgen-binding sites in the rat urogenital sinus and post¬ natal prostate. J. Endocrinol., 104:87-92.

26

/ Chapter 1

230. Taketo, T., Merchant-Larios, H., and Koide, S. S. (1984): Induc¬ tion of testicular differentiation in the fetal mouse ovary by transplan¬ tation into adult male mice. Proc. Soc. Exp. Biol. Med., 176: 148-153. 231. Taketo-Hosotani, T., Merchant-Larios, H., Thau, R. B., and Koide, S. S. (1985): Testicular differentiation in fetal mouse ovaries fol¬ lowing transplantation into adult male mice. J. Exp. Zool., 236:229237. 232. Teng, C. S., andTeng, C. T. (1979): Prenatal effect of the estrogenic hormone on embryonic genital organ differentiation. In: Ontogeny of Receptors and Reproductive Hormone Action, edited by T. H. Hamilton, J. H. Clark, and N. A. Sadler, pp. 421-440. Raven Press, New York. 233. Torrey, T. W. (1950): Intraocular grafts of embryonic gonads of the rat. J. Exp. Zool., 115:37-38. 234. Tran, D., and Josso, N. (1982): Localization of antimullerian hor¬ mone in the rough endoplasmic reticulum of the developing bovine Sertoli cell using immunocytochemistry with a monoclonal antibody. Endocrinology, 111:1562-1567. 235. Tran, D., Picard, J. Y., Vigier, B., Berger, R., and Josso, N. (1986): Persistence of mullerian ducts in male rabbits passively immunized against bovine anti-mullerian hormone during fetal life. Dev. Biol., 116:160-167. 236. Trelstad, R. L., Hayashi, A., Hayashi, K., and Donahoe, P. K. (1982): The epithelial-mesenchymal interface of the male mullerian duct: Basement membrane integrity and ductal regression. Dev. Biol., 92:27-40. 237. Turner, C. D. (1940): The influence of testosterone proprionate upon sexual differentiation in genetic female mice (etc.). J. Exp. Zool., 83:1-31. 238. van Niekerk, W. A. (1974): True Hermaphroditism. Clinical Mor¬ phologic and Cytogenetic Aspects. Harper and Row, New York. 239. van Niekerk, W. A. (1981): True hermaphroditism. Pediatr. Adolesc. Endocrinol., 8:80-99. 240. van Niekerk, W. A., and Retief, A. E. (1981): The gonads of human true hermaphrodites. Hum. Genet., 58:117-122. 241. Venolia, L., Cooper, D. W., O’Brien, D. A., Millette, C. F., and Gartler, S. M. (1984): Transformation of the Hprt gene with DNA from spermatogenic cells. Chromosoma, 90:185-189. 242. Verhoeven, G., and Wilson, J. D. (1976): Cytosol androgen binding in submandibular gland and kidney of the normal mouse and the mouse with testicular feminization. Endocrinology, 99:79-92. 243. Vergnaud, G., Page, D. C., Simmler, M. C., Brown, L., Rouyer, F., Noel, B., Botstein, D., de la Chapelle, A., and Weissenbach, J. (1986): A deletion map of the human Y chromosome based on DNA hybridization. Am. J. Hum. Genet., 38:109-124. 244. Veyssiere, G., Corre, M., Berger, M., Jean-Faucher, Ch., de Turikheim, M., and Jean, Cl. (1980): Androgenes circulants etorganogenese sexuelle male chez le foetus de lapin. Etude apres immu¬ nisation active delamere contre la testosterone. Arch. Anat. Microsc. Morphol. Exp., 69:17-28. 245. Vigier, B., Picard, J-Y., and Josso, N. (1982): A monoclonal an¬ tibody against bovine anti-mullerian hormone. Endocrinology, 110:131137. 246. Vigier, B., Legali, L., Picard, J.-Y., and Josso, N. (1982): A sen¬ sitive radioimmunoassay for bovine anti-mullerian hormone, allow¬ ing its detection in male and freemartin fetal serum. Endocrinology, 111:1409-1411. 247. Vigier, B., Picard, J.-Y., Champargue, J., Forest, M. G., Heyman,

Y., and Josso, N. (1985): Secretion of anti-mullerian hormone by immature bovine Sertoli cells in primary culture studied by a com¬ petition-type radioimmunoassay: Lack of modulation by either FSH or testosterone. Mol. Cell. Endocrinol., 43:141-150. 248. Vogel, F., and Motulsky, A. G. (1979): Human Genetics. SpringerVerlag, Berlin. 249. Wachtel, S. S. (1983): H-Y Antigen and the Biology of Sex Deter¬ mination. Grune and Stratton, New York. 250. Wachtel, S. S., Ohno, S., Koo, G. C., and Boyse, E. A. (1975): Possible role of H-Y antigen in the primary determination of sex. Nature, 257:235-236. 251. Walsh, P. C., Madden, J. D., Harrod, M. J., Goldstein, J. L. MacDonald, P. C., and Wilson, J. D. (1974): Familial incomplete male pseudohermaphroditism, type 2. Decreased dihydrotestosterone formation in pseudovaginal perineoscrotal hypospadias. N. Engl. J. Med., 291:944-949. 252. Wells, L. J., and van Wagenen, G. (1954): Androgen-induced female pseudohermaphroditism in the monkey (Macaca mulatto); anatomy of the reproductive organs. Contrib. Embryol. Carnegie Inst. (Wash.), 35:93-106. 253. White, M. R. (1949): Effects of hormones on embryonic sex dif¬ ferentiation in the golden hamster. J. Exp. Zool., 110:153-181. 254. Wilbert, D. M., Griffin, J. E., and Wilson, J. D. (1983): Charac¬ terization of the cytosol androgen receptor of the human prostate. J. Clin. Endocrinol. Metab., 56:113—120. 255. Wilson, J. D. (1971): Testosterone metabolism in skin. Symp. Dtsch. Ges. Endokrinol., 17:11-18. 256. Wilson, J. D. (1979): Embryology of the genital tract. In: Urology, Vol. 2, 4th ed., edited by J. H. Harrison, R. F. Gittes, A. D. Perlmutter, T. A. Stamey, and P. C. Walsh, Chapter 41, pp. 1469— 1483. W. B. Saunders, Philadelphia. 257. Wilson, J. D., and Goldstein, J. L. (1975): Classification of hered¬ itary disorders of sexual development. Birth Defects, 11:1-16. 258. Wilson, J. D., and Lasnitzki, I. (1971): Dihydrotestosterone for¬ mation in fetal tissues of the rabbit and rat. Endocrinology, 89:659-

668. 259. Wilson, J. D., and Siiteri, P. K. (1973): Developmental pattern of testosterone synthesis in the fetal gonad of the rabbit. Endocrinology, 92:1182-1191. 260. Wilson, J. D., Griffin, J. E., Leshin, M., and MacDonald, P. C. (1983): The androgen resistance syndromes: 5a-reductase deficiency, testicular feminization, and related disorders. In: The Metabolic Basis of Inherited Disease, edited by J. B. Stanbury, J. B. Wyngaarden, D. S. Fredrickson, J. L. Goldstein, and M. S. Brown, pp. 10011026. McGraw-Hill, New York. 261. Witschi, E. (1948): Migration of the germ cells of human embryos from the yolk sac to the primitive gonadal folds. Contrib. Embryol. Carnegie Inst. (Wash:), 32:67-80. 262. Wolfe, J., and Goodfellow, P. N. (1985): The elusive testis deter¬ mining factor. Trends Genet., 1:3-4. 263. Yen, P. H., Patel, P., Chinault, A. C., Mohandas, T., and Shapiro, L. J. (1984): Differential methylation of hypoxanthine phosphoribosyltransferase genes on active and inactive human X chromosomes. Proc. Natl. Acad. Sci. USA, 83:1759-1763. 264. Zaborski, P. (1985): H-Y antigen in nonmammalian vertebrates. Arch. Aust. Microsc. Morphol. Exp., 74:33-37. 265. Zenzes, M. T., and Reed, T. E. (1984): Variability in serologically detected male antigen titer and some resulting problems: A critical review. Hum. Genet., 66:103-109.

The Physiology of Reproduction, edited by E. Knobil and J. Neill et al. Raven Press, Ltd., New York © 1988.

CHAPTER

2

The Spermatozoon E. M. Eddy

The Plasma Membrane, 28 Surface Domains, 28 • Formation of Domains, 33 • Maintenance of Domains, 34 • Plasma Membrane Composition, 35 • Modification of the Sperm Plasma Membrane During Epididymal Maturation, 35 •

The Sperm Nucleus, 41 • Cytoskeleton of Sperm Head, 43 • The Acrosome, 45

The Flagellum, 47 Connecting Piece, 47 • Axoneme, 48 • Mitochondrial Sheath, 49 • Outer Dense Fibers, 50 • Fibrous Sheath, 52 • Abnormal Flagella, 53 • Flagellar Motion, 54

Modification of the Sperm Plasma Membrane During Ejaculation and Capacitation, 40

Summary, 55 References, 57

The Sperm Head, 41

The spermatozoon is the end product of the process of gametogenesis in the male, occurring within the seminif¬ erous tubules of the testis. This involves a series of mitotic division of spermatogonial stem cells, two meiotic divisions by spermatocytes, extensive morphological remodeling of the spermatid during spermiogenesis, and the release of the free cell into the lumen of the seminiferous tubule by spermiation. It is an interesting paradox that the process of spermatogenesis produces a cell that is highly differentiated in structure and function, while at the same time is developmentally totipotent, being able to combine with the egg and thereby begin the process that gives rise to a new in¬ dividual. The mammalian spermatozoon has two main components, the head and the tail or flagellum (Fig. 1). The head consists of the acrosome, the nucleus, and lesser amounts of cytoskeletal structures and cytoplasm. The acrosome is a large secretory granule that closely surrounds and overlies the anterior end of the nucleus. The sperm nucleus is haploid, containing only one member of each chromosome pair, and the chromatin becomes highly condensed during the latter part of spermatogenesis. The tail contains a centrally placed axoneme, which is a highly ordered complex of microtubles surrounded by dense fibers extending from the head to near the posterior end of the axoneme. In addition, the anterior part of the flagellum contains mitochondria wrapped in a tight helix around the dense fibers, and the most posterior part of the tail contains the fibrous sheath surrounding the dense fibers. The dense fibers and the fibrous sheath form the cytoskeleton of the flagellum. These cytoskeletal fea¬ tures appear to have evolved with the development of in¬

ternal fertilization (1). The tail, like the head, is closely wrapped by the plasma membrane and contains little cy¬ toplasm. Although all mammalian spermatozoa have these general characteristics, there are species-specific differences in the size and shape of the head as well as the length and relative size of the components of the flagellum. Nonmam¬ malian species show greater variation in sperm structure; although sperm in most invertebrates and nonmammalian vertebrates have an acrosome, they often contain few mi¬ tochondria, and their flagellum consists only of an axoneme. In some species, spermatozoa are amoeboid cells lacking an acrosome and a flagellum (2,3). The specialized structural features of the spermatozoon are a reflection of its unique functional activities. The ac¬ rosome contains enzymes essential for fertilization, while the flagellum contains the energy sources and machinery necessary to produce spermatozoon motility. The roles of these components are to ensure delivery of the genetic ma¬ terial contained in the nucleus to the egg, where combination of the haploid male and female pronuclei occurs, ending the process of reproduction and initiating the process of development. In most vertebrates, the sex chromosome car¬ ried in the haploid sperm nucleus determines the gender of the resulting individual (4). Both a maternal and a paternal genome are required for development to proceed to term, probably because differential imprinting of gamete genomes occurs during gametogenesis in males and females (5). This chapter examines the structure and function of the mammalian spermatozoon, with an emphasis on the mol¬ ecules currently known to be involved. The major topics considered are: the organization of the sperm plasma mem-

27

28

/ Chapter 2

HEAD

FLAGELLUM

FIG. 1. General features of the mammalian sperm. The head of the spermatozoon is attached to the connecting piece of the fla¬ gellum. The other regions of the flagellum are the middle piece, the principal piece, and the end piece. The middle piece con¬ tains the mitochondrial sheath, while the principal piece contains the fibrous sheath. Longitudinal and cross-sectional views of the principal piece and a segment of fi¬ brous sheath are indicated by arrows; the internal components of the flagellum are identified in Fig. 9.

brane into domains, changes in the composition and function of the domains during the life of the cell, the structural components of the head of the sperm, and the features of the flagellum. Other chapters are concerned with the for¬ mation of the male gamete (6), participation of the sper¬ matozoon in fertilization (7), and the subsequent develop¬ ment of a new individual (8). They contain additional information that is important for understanding the structure and function of the spermatozoon and its role in the repro¬ ductive process.

THE PLASMA MEMBRANE Surface Domains A unique feature of the spermatozoon is that the plasma membrane is subdivided into sharply delineated regional domains that differ in composition and function. The het¬ erogeneous nature of the sperm surface first became apparent from studies of surface charge, lectin binding to specific sugar moieties, freeze-fracture patterns, and antibody la¬ beling. The evidence that the organization and composition

of the plasma membrane varied between different regions of the sperm surface led to the concept that the sperm plasma membrane is a mosaic of restricted domains that reflect the specialized functions of surface and cytoplasmic compo¬ nents of the spermatozoon (9). Subsequent studies have supported this and have further demonstrated that the do¬ mains are dynamic features that undergo changes in organ¬ ization and composition during the life of the cell. The major domains of the head region of the sperm sur¬ face in most mammals (Fig. 2) are (a) the anterior acrosome (acrosomal cap) and equatorial segment (posterior acro¬ some) overlying the acrosome and (b) the postacrosomal region (postacrosomal sheath, postnuclear sheath) covering the portion of the head posterior to the acrosome. In ad¬ dition, the less well-defined anterior band is situated be¬ tween the anterior acrosome and the equatorial segment (10), and a serrated band girdles the sperm head at the posterior margin of the equatorial segment. The posterior ring (nuclear ring, striated ring) lies at the junction between head and tail and apparently forms a tight seal between the cytoplasmic compartments of the two main portions of the spermatozoon. The plasma membrane of the flagellum is

The Spermatozoon

/

29

FIG. 2. Plasma membrane domains on the sur¬ face of the head of mouse and rabbit sperma¬ tozoa. The anterior acrosome domain is more extensive on sperm containing a spatulate head, such as that of the rabbit, whereas the equatorial segment domain overlying the posterior part of the acrosome is larger on sperm containing a falciform head. The posterior ring forms a bound¬ ary between the postacrosomal domain of the head and the middle piece domain of the flagel¬ lum. The distribution of the domains on the heads of sperm of these and other species are com¬ pared in Fig. 3.

separated into domains overlying the middle piece and pos¬ terior tail (distal tail) by the annulus, a fibrous ring that surrounds the components of the axoneme and is firmly attached to the membrane.

The Mosaic Sperm Surface The first studies to suggest the heterogeneity of the sperm plasma membrane were those that examined regional dif¬ ferences in surface charge (Table 1). When spermatozoa from the ram, rabbit, or bull were suspended between op¬ positely charged electrodes, they were found to be drawn tail first by electrophoresis toward the anode. This suggested that sperm have a net negative charge and that more of this charge is on the tail than at the head (11-13). Other studies indicated that at least some of the moieties responsible for sperm net negative charge were on the surface and that there were regional differences in this surface charge. Electron microscopy was used to show that binding of positively charged colloidal iron hydroxide to the surface of rabbit sperm was greater on the flagellum than on the head (14,15). Lectins have relatively specific affinities for particular saccharide molecules (16,17). They are often multimeric and can bind simultaneously to more than one saccharide ligand on a given cell or can cause agglutination by linking adjacent cells. Lectins can be tagged with fluorescent mark¬ ers, enzymes, radioactive labels, or materials visible in the electron microscope. Agglutination assays carried out with spermatozoa from different species have demonstrated re¬ gional differences in the location and amount of specific saccharides on the sperm surface (Table 1). An early study with soybean agglutinin (SBA; recognizing a-D-GalNAc, D-Gal) indicated that it caused predominantly tail-to-tail ag¬ glutination of ejaculated bull sperm (18). Other studies in¬ dicated that hamster sperm agglutinated more readily by the tail than the head, following treatment with concanavalin A (Con A; recognizing a-D-Glc, a-D-Man) or wheat-germ agglutinin (WGA; recognizing [(3(l-4)D-GalNAc]2) (19).

However, it has been noted that there are discrepancies between the number of lectin-binding sites and the agglu¬ tination of treated cells (20,21). Quantitative studies using iodinated lectins indicated that there are 1 to 3 x 107 ConA- and WGA-binding sites per rabbit or hamster epididymal spermatozoon, but fewer RCA-binding sites (22). When expressed on the basis of surface area, the average density of binding sites was similar between sperm from those spe¬ cies. However, when the data on Con-A binding were re¬ calculated on the basis of square-micrometer surface area, the sperm head possessed approximately 10 times more sites than the flagellum (21). The greater surface area and motion of the tail, compared to the head, apparently lead to tailto-tail agglutination, even though the tail has a lower number of lectin-binding sites. The use of lectins conjugated with fluorescent labels or ultrastructural markers has confirmed the regional hetero¬ geneity of distribution of saccharides on the sperm surface (Table 1). However, they have given somewhat different results than were seen in agglutination studies. With flu¬ orescent lectins, Con A was found to bind predominantly to the acrosome region of mouse sperm (23,24), WGA was found to bind to the head of mouse sperm and over the acrosome of guinea-pig sperm (25,26), SB A was found to bind over the anterior acrosome of guinea-pig sperm (21), and peanut agglutinin (PNA; recognizing D-Gal, (3[l-3]GalNAc) was found to bind over the anterior acrosome of mouse sperm (27). A peroxidase histochemical method was used to show that Con-A binding is more intense on the head than on the tail of rabbit sperm (28). Other ultrastruc¬ tural studies used ferritin-conjugated castor-bean agglutinin (RCA; recognizing (3-D-Gal, D-GalNAc) on rabbit sperm (29) and used ferritin- and hemocyanin-labeled Con A on hamster sperm (30) and also noted greater binding on the head than on the tail. These studies confirmed that lectin¬ binding sites are generally present in higher density on the head than on the tail of the sperm. However, the surface domains recognized by lectins are often not well defined, and different binding patterns have been observed for the

30

/ Chapter 2 TABLE 1. Regional heterogeneity of the sperm plasma membrane References

Characteristic

same lectin on sperm from the same species (see, e.g., refs. 21 and 24). Freeze-fracture, freeze-etch, and surface replica studies have shown differences in the number and patterns of dis¬ tribution of intramembranous particles and in membraneassociated structures in different regions of the sperm plasma membrane (Table 1). Although there are significant species differences, these studies have demonstrated characteristic features of the plasma membrane associated with the anterior region of the acrosome, the equatorial segment, the posterior margin of the equatorial segment, and the postacrosomal area. Other regions of the sperm plasma membrane showing structural specialization are (a) the posterior ring at the junc¬ tion between head and tail and (b) the annulus situated between the midpiece region and the more distal part of the tail. In most species, the plasma membrane overlying the an¬ terior acrosome and equatorial segment contains randomly distributed intramembranous particles, but numerous hex¬ agonal arrays of particles are present in guinea-pig sperm stacked closely together in the epididymis, and small patches

11-13 14, 15

SI--33 26, 31, 34,

00

18, 19 21, 23--30

CO

Surface charge distribution Sperm drawn tail-first toward the anode in electrophoretic field Positively charged colloidal iron hydroxide particle binding greater on the tail than on the head Surface saccharide distribution Sperm agglutinated tail-to-tail following lectin treatment Different lectins bind to specific regions of the sperm surface Intramembranous particle distribution Hexagonal arrays of particles in the plasma membrane overlying the acrosome Dense populations of intramembranous particles in the plasma membrane overlying the postacrosomal region Oblique strands of particles in plasma membrane of middle piece overlying mitochondria Localized surface features Serrated band at posterior margin of equatorial segment of acrosome Palisades of prominent particles at posterior margin of postacrosomal region Belt of fine periodicities in plasma membrane over the posterior ring Close array of fine particles in plasma membrane over annulus Staggered row of large particles running longitudinally on principal piece Membrane intercalating agents Filipin-induced complexes with sterols frequent in plasma membrane over acrosome Polymyxin B binds anionic phospholipids in plasma membrane over anterior acrosome Fluorescent lipid analog integrates preferentially into plasma membrane over acrosome Sperm plasma membrane antigens recognized by antisera Antigens shared with other cells in restricted domains on sperm surface Equatorial segment Potacrosomal region Anterior acrosome Antigens in single domains recognized by antisera to male germ cells Anterior acrosome Postacrosomal region Principal piece Antigens in multiple domains recognized by antisera to male germ cells Equatorial segment and postacrosomal region Acrosome and midpiece Head and midpiece Head and tail Entire sperm surface

41

31, 38, 40, 49, 51 31, 31, 31, 31, 31,

33--36, 38, 39 41--43 34, 37, 38, 49 49 49, 51, 55, 56, 58

40, 43, 56, 59-61 60, 62--64 65

67 68 69 70 71 72 73 72 74 75 76-78

of similarly organized particles are seen occasionally in rat sperm (31). These particle arrays have been noted by other investigators studying rat (32) and boar sperm (33). The posterior margin of the equatorial segment usually has a saw-toothed pattern, formed of rows of closely packed par¬ ticles (Table 1). The location and general topography of this serrated band can be seen as a surface feature on whole spermatozoa by light microscopy (34), scanning electron microscopy (34-36), and transmission electron microscopy of surface replicas (37-39). The plasma membrane of the postacrosomal area of the sperm head usually contains a more dense population of intramembranous particles (26,31,34,39-41) than does the acrosomal area, and the particles are sometimes in clusters (Table 1). The basal part of the postacrosomal area contains prominent cords or bands of particles (31,41-43), which often lie in palisades (Table 1). They lie adjacent to the posterior ring, which is a belt of fine periodicities or cords of small particles in the plasma membrane at the junction between head and tail of the spermatozoon (31,34,37,44). This structure was first reported by light microscopists (45,46),

The Spermatozoon and its presence was confirmed by electron microscopists studying sections (47) and replicas (48) of spermatozoa. The posterior ring is formed by plasma membrane being closely applied to a belt of dense fibrous material lying upon the nuclear envelope (31,34,37,38,44,48) and appears to produce a seal between head and tail compartments of the spermatozoon. The posterior ring of mammalian sperma¬ tozoa may allow substantially different ionic and metabolic conditions to be maintained in the cytoplasmic compart¬ ments of the head and flagellum (49). The plasma membrane of the midpiece region of the fla¬ gellum in some species contains strands of particles, which run in diagonal arrays that coincide in pitch with the un¬ derlying helically wound mitochondria (Table 1). The par¬ ticle strands appear to be present only when the plasma membrane is closely applied to mitochondria; these strands are usually not present in cytoplasmic droplets (31.38.40.41.50.51) . It has been suggested (52) that the particle strands might be homologous to the necklace of particles at the base of cilia (see, e.g., ref. 53). The strands are particularly prominent in guinea-pig sperm; in other species, however, patches of hexagonally packed particles are present in the midpiece plasma membrane and tend to follow the contour of the mitochondrial helix (49). A dif¬ ferent pattern has been reported in opossum spermatozoa, where intramembrane particles are present in parallel lon¬ gitudinally arranged aggregates (54). The annulus is a dense fibrous ring surrounding the axonemal complex at the junction between the middle piece and the principal piece of the flagellum. Upon use of freezefracture techniques, the plasma membrane in this area ap¬ pears to have a rough texture or to contain a close array of small particles, sometimes in circumferential strands (31). The plasma membrane is closely applied to the annulus, and these features may represent anchoring of the annulus into the membrane (49). The intramembranous particles in the plasma membrane of the principal piece posterior to the annulus appear larger than those in the middle piece. In addition, sperm of some species have a staggered double row of yet larger particles (31.51) , resembling a zipper coursing longitudinally over the ribs of the fibrous sheath, opposite outer dense fiber number 1 (31). The zipper-like structure terminates before reaching the posterior end of the principal piece of the fla¬ gellum. The large particles that form this structure are seen on surface replicas and are probably part of a transmembrane complex (49,55). The particles appear to be slightly oval and to have a depression in their center, possibly indicative of a pore. This feature has led to the suggestion that zipper particles may be sites of ion transport (56). It has also been postulated that the zipper is a membrane-anchoring device for some of the axonemal components (55). Treatment of guinea-pig sperm with digitonin, a detergent that disrupts cholesterol-rich plasma membrane (57), does not remove the zipper. However, the zipper particles can be removed by subsequent treatment with Triton X-100 (55). The zipper

/

31

binds the lectins Con A, RCA, and WGA, and the TritonX-100-soluble fraction includes four polypeptides ranging from 24,000 to 110,000 daltons (58). Freeze-fracture techniques used in conjunction with mem¬ brane intercalating agents has provided evidence of com¬ positional differences between sperm surface membrane do¬ mains (Table 1). Treatment of guinea-pig sperm with filipin, a polyene that complexes (3-hydroxysterols with sterol, in¬ dicated that the regions between the plaques of intramem¬ branous particles in the plasma membrane over the anterior acrosome and equatorial segment are sterol-rich (43,59). The sterols (cholesterol and dermosterol) appear to be pres¬ ent in higher concentration in the inner half than in the outer half of the membrane leaflet (60). However, filipin fluo¬ rescence indicates that sterols may exist throughout the en¬ tire plasma membrane of the anterior acrosome, not just between the plaques (10). The plasma membrane of the postacrosomal region has less than one-fourth as many filipin-induced complexes as that of the anterior acrosome and equatorial segment, with a pattern suggesting that they are present mainly in the outer half of the bilayer (10,60). These studies also indicated that the anterior acrosome is separated from the equatorial segment by an anterior band, a subdomain that, in guinea-pig sperm, often contains circles of membrane cleared of sterols and intramembranous par¬ ticles (56,61). The antibiotic polymyxin B binds anionic phospholipids, producing crenulations of anionic phospholipid-rich mem¬ branes (62,63). Treatment of guinea-pig sperm with poly¬ myxin B indicated that the plasma membrane over the an¬ terior acrosome has a high anionic lipid concentration (64). In sperm from the epididymis, the concentration of anionic lipids appears to be greatest at the anterior tip of the ac¬ rosome. The high anionic lipid concentration is not seen in the anterior band or posterior to the equatorial segment (60,62,64). The fluorescent lipid analog, l,l'-dihexadecyl-3,3'-tetramethyl-indocarbocyanine perchlorate (C16dil), intercalates into the outer leaflet of the sperm plasma membrane, and it was found that the anterior part of the head of ram sperm was labeled more intensely than the posterior head with this agent (65). The differences in affinity of the probe for these regions of the sperm apparently result from interactions of the probe with lipids and proteins heterogeneously distrib¬ uted within the plane of the membrane.

Sperm-Surface Antigens Antibodies to spermatozoa, germ cells, other cell types, or isolated molecules have identified more distinct surface regions on living sperm than those seen with other methods (Table 1). Antibodies can be conjugated directly with var¬ ious labels or detected indirectly with second antibodies (66) that carry labels visible by light or electron microscopy. Antibodies can also be used to isolate and identify specific

32

/ Chapter 2

molecules and to test their roles in bioassays. In some cases, it has been found that antigens shared with other cell types are restricted to specific regions of the sperm surface. With mouse sperm, (a) an antiserum to the H-Y antigen has been found to react with the plasma membrane over the equatorial segment (67), (b) an antiserum to F9 teratocarcinoma cells to bind to the postacrosomal region (68), and (c) an anti¬ serum to galactosyltransferase to bind to the anterior acrosome (69). In addition, some antisera to sperm or spermatogenic cells react with specific regions, such as anterior acrosome (70), postacrosomal region (71), or principal piece (72). Other antisera to germ cells react with multiple regions of the sperm surface, such as equatorial segment and post¬ acrosomal region (73), acrosome and midpiece (72), head and midpiece (74) or whole tail (75), or even with the entire sperm surface (76-78). The molecules recognized by these antisera have not been identified in most cases. It is tempting to speculate that antisera reacting with specific regions may recognize one or a few antigens, whereas those reacting with the whole surface recognize multiple antigens. How¬ ever, binding to the whole sperm surface has been seen with both an antiserum to a single antigen (77) and one to multiple antigens (78). The multiple and variable specificities of antisera to whole cells or mixtures of antigens limit their usefulness for dis¬ secting the distribution of sperm-surface components and defining the biochemical characteristics and functional roles of such components. Monoclonal antibodies have been pro¬ duced against spermatozoa to overcome some of these lim¬ itations (Table 2). Although monoclonal antibodies are often specific to particular molecules, they may also recognize epitopes shared by many molecules (see, e.g., ref. 79). Furthermore, monoclonal antibodies, which label only one domain on the sperm surface, may immunoprecipitate pro¬ teins of more than one molecular weight from that domain (see, e.g., ref. 80). It is possible that these proteins are either subunits of a membrane complex that coprecipitate under such conditions or are not associated but share a common epitope recognized by the antibody. Studies with monoclonal antibodies have been carried out most fre¬ quently in mouse, rat, guinea pig, and human, but less often in boar, hamster, and rabbit. In some cases, the monoclonal antibody recognizes the whole sperm surface (81-83), the whole head (80,83,84,86,100,101), or the whole tail (80,84,87-93). Other monoclonal antibodies bind specifi¬ cally to anterior acrosome, equatorial segment, postacro¬ somal region, middle piece, or posterior tail of living sperm (79,80,84,85,94-108) (Table 2). Other patterns have been reported on air-dried or fixed sperm, but these treatments can expose internal antigens (see, e.g., ref. 82). In addition, some monoclonal antibodies recognize variable patterns or apparent subdomains on spermatozoa, possibly as a result of shedding of sperm-surface components during process¬ ing, partial masking of components by extrinsic molecules, or relocalization of components due to membrane fluidity (see, e.g., ref. 109). However, a variety of studies have

TABLE 2. Sperm-surface domains recognized with monoclonal antibodies

Antibodies

Species

Mass (kilodaltons)

Antigens restricted to single domains Anterior acrosome HS 1 A.1 Human NDa MA 1 Human 84 MA 2 Human ND MA 3 Human 240 D81,G112, Human ND G176, G225b SMA1 Mouse ND MS-1 Mouse 69 1B3 Mouse 28 AMSIV-33fa Mouse 200 — Mouse ND J1, C6 Mouse ND M5 Mouse 54 M4F Mouse 21, 35, 60 M42 Mouse 220-240 AH-1 Guinea pig 52 AH-2 Guinea pig 18, 25, 46, 62, 70 AH-3 Guinea pig 38, 52, 62 AH-4 Guinea pig 16, 38 AH-5 Guinea pig ND 8C10.5 Rabbit 63 Equatorial segment D3 Human ND M2C Mouse 36, 44 M29c Mouse 40 Postacrosomal region HS 1 E.1 Human 53, 56, 73 PH-1 Guinea pig 60 PH-2 Guinea pig 41, 48, 66 PH-3 Guinea pig 48, 58 PH-4 Guinea pig ND 1B6 Rat ND WS 35.22 Hamster ND Middle piece MA 4 Human 30 HSA-1 Human ND AMSIV-256 Mouse ND Posterior tail PT-1 Guinea pig ND

References

79 94 94 94 95 84 96 97 92 98 27 99 99 85, 99 80, 100, 101 100, 101 100, 101 100, 101 100, 101 102 95 103 103 79 100, 100, 100, 100, 93 104

101 101 101 101

94 88 92 80, 100, 101

Antigens present in multiple domains Whole head SMA3 OBF13 WH-1 WH-2 WH-3 HMS3.1

Mouse Mouse Guinea pig Guinea pig Guinea pig Hamster

ND ND 42 45, 89 ND ND

84 86 80 80, 100, 101 80, 100, 101 83 (continued)

used monoclonal antibodies against sperm from different species to confirm that the sperm surface consists of welldefined domains and that different proteins and glycopro¬ teins may be either confined to individual domains or shared by multiple domains (Table 2).

The Spermatozoon TABLE 2. (Continued) Antibodies

Species

Mass (kilodaltons)

Acrosome and midpiece G177 Human WH 97.25 Hamster

ND >500 Equatorial segment and middle piece bF4 Human ND HS 2M.1 Human 32, 83 HS 2N.1 Human 26-105 Equatorial segment and principal piece M2 Mouse 36, 44 Equatorial segment and whole tail MA 5 Human 71 MA 6 Human ND Postacrosomal region and middle piece HS-4 Human 130 Postacrosomal region and whole tail MA-24b Human 23 XT-1 Mouse ND Neck and middle piece MS 76.11 Hamster ND

References 95 104 95 79 79 103 94 94 105 106 107, 108 104

Middle piece and end piece WS 64.23 Hamster Whole tail YWK-1 Human HSA-1 Human SP1D1, AP7A7 Human SMA4 Mouse T21 Mouse AMSIV-54i), Mouse AMSIV-75b WT-1 Guinea pig 2B1 Rat

84 ND ND 54 31 ND

87 88 89 84, 90 91 92

ND 40

80 93

Whole sperm 2D6 HMS1.0, HMS2.0

23 ND

81, 82 83

Rat Hamster

23

104

aND, not determined. ^Determined by indirect immunofluorescence on fixed or air-dried sperm. determined by indirect immunofluorescence on acrosome-reacted sperm.

Formation of Domains It is likely that most sperm-surface domains are estab¬ lished during spermiogenesis, while the round spermatid is being remodeled into the spermatozoon. However, sper¬ matozoa of some species undergo shape changes in the epididymis, and their surface domains reach final distri¬ bution and form after spermiogenesis. Probably most af¬ fected are the middle piece and the acrosome. The cyto¬ plasmic droplet migrates from the anterior to the posterior end of the middle piece and is usually shed as the sperm transits the epididymis. The acrosome can undergo a re¬

/

33

duction in size (110,111) and, in some species, most notably the guinea pig, a substantial change in shape (112). New surface antigens also appear in specific domains during mat¬ uration, but they probably arise through modification or unmasking of preexisting molecules or by attachment of new molecules to acceptor sites already segregated into do¬ mains (see, e.g., ref. 113). It has also been reported that a monoclonal antibody reacts with the whole head of testicular sperm in the guinea pig, but only with the postacrosomal region of sperm from the tail of the epididymis (101). Such a change could occur by lateral migration of that antigen in the plasma membrane or by loss of an antigen from one domain and appearance of another antigen with the same epitope in a different domain (101). The mechanisms responsible for establishing sperm-sur¬ face domains during spermiogenesis have not been defined. However, most of the surface domains overlie distinct cy¬ toplasmic organelles or features. Some of the mechanisms that establish the shape and organization of the spermato¬ zoon may also be involved in determining the location of surface domains. Morphogenetic processes that shape the sperm are probably carried out by cytoskeletal components of the germ cell. Transmembrane proteins that are stabilized by attachment through linkage proteins to cytoskeletal struc¬ tures (114) may define the boundary and contents of different domains. An acrfn-specific monoclonal antibody was used in con¬ junction with biochemical procedures to show that testicular spermatozoa of the rabbit contain filamentous actin in a sheath around the nucleus and throughout the equatorial segment and postacrosomal region (115). This study also confirmed an earlier report that actin is present in the post¬ acrosomal region of rabbit epididymal spermatozoa (116). In addition, actin has been reported to be present in the subacrosomal space of testicular spermatozoa of the rat (116); in the posterior region of the head and in the connecting piece, midpiece, and principal piece of the tail of human sperm (116-118); and in the postacrosomal region of sper¬ matozoa from boar (119,120). One study has reported that myosin is present in the acrosomal region of mammalian spermatozoa (116). Filamentous structures have been ob¬ served associated with the plasma membrane on the concave margin of the acrosome in vole spermatozoa (121). These may correspond to a cytoskeletal structure recently identified lying between the plasma membrane and the acrosome in hamster sperm (122). Spectrin is a major actin-binding protein and usually is associated closely with the cytoplasmic surface of the plasma membrane (123,124). It is also a main calmodulin-binding protein and a structural component of the cytoskeleton (124). It was present in the cytoplasm overlying the acrosome of round spermatids and elongating spermatids as well as in that of some testicular spermatozoa in the mouse, but was not detected in epididymal spermatozoa (125). Guinea-pig spermatozoa are reported to lack spectrin (126), but human spermatozoa apparently contain spectrin in the anterior ac-

34

/ Chapter 2

rosome and principal piece regions (127). This suggests that spectrin and possibly other actin-binding proteins might be involved in establishing sperm-surface domains. Microtubules have been shown to play an important role in determining the cell shape, and it has been suggested that microtubules in the manchette may be responsible for the elongation and shaping of the spermatid nucleus (128). The manchette is a sheath of microtubules that assemble as sper¬ matid elongation begins. It attaches to a deposition of dense fibrillar material on the plasma membrane which migrates from the anterior to the caudal end of the nucleus during spermiogenesis (129). This attachment site later becomes the posterior ring lying at the junction between head and tail, which serves to separate the domains of the postacrosomal region and the middle piece. This suggests that the manchette contributes to the definition of the boundary be¬ tween these two surface domains. However, a comparative study of morphogenetic factors influencing the shape of the sperm head concluded that its form is probably a conse¬ quence of the aggregation of DNA and protein during con¬ densation of chromatin rather than due to forces external to the nucleus (129). In addition, microtubule-like components have been observed in close association with the plasma membrane overlying the postacrosomal region of the sper¬ matozoon of the bull (130) and vole (121). However, they have not been shown to contain tubulin. Such features may represent the periodic densities seen in the postnuclear dense lamina (130,132). The annulus also appears during spermiogenesis, encir¬ cling the axonemal complex at the distal end of the basal body, where the plasma membrane is reflected onto the forming flagellum of the elongating spermatid (133). The annulus remains firmly adherent to the plasma membrane as it moves down the flagellum to take up its final position, late in spermiogenesis, at the posterior end of the mito¬ chondrial sheath and the anterior end of the fibrous sheath. At this location it also serves to separate the plasma mem¬ brane domains of middle piece and posterior tail. The Sertoli cells may influence the establishment of do¬ mains, particularly those on the sperm head. Junctional structures, referred to as Sertoli-cell “ectoplasmic special¬ izations,” maintain a tight association between Sertoli cells and spermatids (134). The ectoplasmic specializations ap¬ pear to grasp the head of spermatids and are present in large amounts facing cells undergoing the elongation and matu¬ ration phases of spermiogenesis (135). This association is maintained until near the time of sperm release (136), ap¬ parently holding the spermatid in a recess extending deeply into the Sertoli cell. The ectoplasmic specialization consists of bundles of actin or actin-like filaments and more deeply placed saccules of endoplasmic reticulum within the Sertoli cell (134); this might aid in shaping the sperm head or in maintaining cell polarity necessary for domain formation. Another specialized relationship between Sertoli cells and spermatids appears during an even later part of spermiogen¬ esis. The “tubulobulbar complexes” form as spermatids be¬

gin to move from deep recesses in the Sertoli-cell plasma membrane toward the lumen of the seminiferous tubule (137). The tubulobulbar complexes arise around the spermatid head as the ectoplasmic specialization begins to dissociate (134). Successive generations of complexes extend from sperma¬ tids into Sertoli cells, where they are actively phagocytosed (138). This process continues until the time of spermiation and morphometric data indicate that as much as 70% of the cytoplasm of spermatids may be eliminated by this mech¬ anism (138). This process also has the potential to substan¬ tially alter the composition and organization of the plasma membrane and could serve a role in defining the nature and organization of proximate surface domains.

Maintenance of Domains Tt has been suggested that sperm-surface domains might be maintained by restriction of mobility of surface molecules in their final domain, by the existence of a membrane barrier to surface component movement at the domain boundary, or through thermodynamic partitioning of molecules into a specific region (101). Restriction of mobility might occur through interactions of intramembranous components with molecules outside or inside the membrane. Although there is no direct evidence that external constraints can be imposed on domains by molecules applied to the sperm surface, capacitation involves loss of extrinsic components and a concomitant increase in membrane mobility in some do¬ mains (discussed below). Also, the glycocalyx between closely stacked sperm heads in the guinea pig has a septate structure (139) that may be congruent with the quilted pattern of intramembranous particles in that area of the plasma mem¬ brane (31). This quilted pattern is diminished by treatments that remove the cell coat (60,61). Furthermore, there is a correlation between the loss of the quilted pattern and changes in the rat sperm glycocalyx in the epididymis (32). Internal constraints seem likely to play a major role in maintaining domains in the plasma membrane. The regions of plasma membrane over the posterior ring and the annulus hardly deserve to be called domains, but cytoplasmic com¬ ponents are involved in the formation and stabilization of these specialized regions. Furthermore, the transmembrane particles that comprise the zipper-like structure of the fla¬ gellum appear to be held in place by attachments to the ribs of the fibrous sheath (31). Also, the oblique strands of intramembranous particles in the middle piece are associated with the underlying mitochondria (31). In addition, the plasma membrane of the postacrosomal region is underlaid by long rod-like structures that form the postnuclear dense lamina (34,140) and that may stabilize the plasma membrane of this domain. Finally, a cytoplasmic tuft underlies the par¬ ticle-bare filipin-complex sparse region of the anterior band, between the anterior acrosome and equatorial segment (10,64). Barriers within the plane of the plasma membrane may restrict the movement of surface components between do-

The Spermatozoon mains. Such potential barriers are present between the do¬ mains of the equatorial segment and the postacrosomal re¬ gion (serrated band), the postacrosomal region and the middle piece (posterior ring), and the midpiece and the distal tail (annulus). For example, a surface antigen (PT-1) has been found to exhibit free diffusion within the plasma membrane of the distal tail domain, but apparently is prevented from migrating into the midpiece by the annulus (141). Also, two antigens (2B1, 2D6) present over the whole surface of the flagellum of rat sperm show antibody-induced patching, but the patches do not migrate onto the head (81). In addition, the lectin RCA caused clustering of binding sites within the postacrosomal region of rabbit sperm, but additional lectin¬ binding sites did not appear to migrate into this area from the acrosome (29). Finally, an antigen (RSA-1) over the entire surface of rabbit sperm undergoes antibody-induced clustering over the acrosome, but not in other regions of the plasma membrane (77). It appears that even though sperm-surface antigens may be mobile within their own domain, they are unable to migrate into other domains. However, this does not appear to be the case for all mem¬ brane components. Using fluorescence recovery after photobleaching, it was observed that there is free exchange of a lipid analog by lateral diffusion between plasma membrane of head and midpiece and between plasma membrane of midpiece and distal tail (65). Plasma Membrane Composition The membranes of human spermatozoa are said to be similar to the plasma membrane of the erythrocyte with regard to lipid composition (142). However, many earlier studies were carried out on whole spermatozoa from the cauda epididymis or ejaculates. More accurate determina¬ tions can be made with purified plasma membranes. Using this approach, it was found that phospholipids make up about 70% of the total plasma membrane lipid in boar sperm, with the major phospholipids being phosphatidylcholine, phosphatidylethanolamine, sphingomyelin, phosphatidylserine, phosphatidylinositol, and lysophosphatidylcholine (143) . Sterols are the next most abundant lipid, with a cho¬ lesterol/phospholipid molar ratio of about 0.12. Glycolipids are less abundant, and this lipid class consists mainly of sulfatoxygalactosylacylalkylglycerol (SGG) (144). Free fatty acid makes up a relatively small amount of the lipid in boar spermatozoa, whereas diacylglycerols are present in about the same amounts as glycolipids (143). SGG is a major component of the germ-cell plasma mem¬ brane in mammals (144). This glycolipid is synthesized in primary spermatocytes (145,146) and remains a stable com¬ ponent without turnover throughout the life of the cell. SGG is present in both the head and tail fractions of spermatozoa (144) . It was detected with an antiserum on spermatogenic cells but not on spermatozoa (147). However, a monoclonal antibody to SGG (148) reacts with the equatorial segment and midpiece of living mouse spermatozoa. SGG binds three

/

35

proteins of 68,000, 34,000, and 24,000 daltons isolated from rat spermatogenic cells (146) and is believed to be tightly associated with those proteins in the sperm plasma membrane. The phospholipid/protein ratio is approximately 0.68 on a weight basis in plasma membranes isolated from boar spermatozoa (143), suggesting that the amounts of total lipid and protein in the sperm plasma membrane are about the same. However, this is for the whole plasma membrane, and the amount and type of lipids and the lipid/protein ratios are probably different in various domains. Freeze-fracture studies with filipin suggest that the amount of sterol in the anterior acrosome is about four times that present in the postacrosomal region in guinea-pig and bull spermatozoa (43,60,61). The postacrosomal region apparently contains few sterols or anionic lipids in guinea-pig spermatozoa (64), but the cytoplasmic droplet of the middle piece is probably rich in both (10,60). Also, cholesterol sulfate makes up only a small fraction of the total sterol but is a major component of the plasma membrane over the acrosome of human sperm (149). However, changes in the lipid content of the spermatozoal plasma membrane occur during maturation and capacitation, and these may have substantial effects on the composition and function of the membrane in different do¬ mains.

Modification of the Sperm Plasma Membrane During Epididymal Maturation Spermatozoa undergo changes during transit through the epididymis that give them the ability to fertilize. Functional modifications that have been identified include alterations in metabolism (150), changes in the pattern and effective¬ ness of flagellar activity (see, e.g., refs. 151 and 152), and acquisition of ability to bind to the zona pellucida (153,154). The sperm plasma membrane is modified in composition during epididymal maturation (Table 3), and some of these changes appear to be required for the functional modifica¬ tions to occur. Changes have been reported in surface charge, lectin binding, intramembranous particle distribution, mem¬ brane fluidity, lipid composition, protein composition, and antibody binding as sperm travel through the epididymis.

Changes in Surface Charge and Intramembranous Particle Distribution The net negative surface charge has been found to be greater on spermatozoa from the cauda than on those from the caput epididymidis in several species (13,155,156). Fur¬ thermore, the density of colloidal iron particles bound to the tail, and to a lesser extent those bound to the head, was greater on rabbit sperm from the cauda epididymidis than on sperm that has not undergone epididymal maturation (14,15). It has been suggested that an increase in sialic acid moieties may be responsible for the change in surface charge

36

/ Chapter 2 TABLE 3. Sperm-surface modifications during epididymal maturation Modification

References

Increase in net negative 13, 14, 155, 156 surface charge Increase in binding of 14, 15, 158, 159, 284 cationic colloidal iron Changes in lectin28, 157, 160-164, binding properties 187, 284 Changes in 32, 33 intramembranous particle distribution Changes in membrane 65 fluidity Changes in lipid 143 composition Decreases in cholesterol, phosphatidylethanolamine, phosphatidylserine, phosphotidylinositol Increases in diacylglycerol, cholesterol sulfate, dermosterol, phosphatidylcholine, sphingomyelin, polyphosphoinositides Changes in protein and glycoprotein composition New surface 186-193 components detected with vectorial labels New surface 194-201 components detected by gel electrophoresis New surface 193, 202-219, 222-224 components detected with antisera New surface 81, 82, 90, 97, 207, 208, components 225-227 detected with monoclonal antibodies Loss of surface 27, 97, 189, 190, 221 components

during maturation (157). One cytochemical study using neuraminidase treatment and colloidal iron staining of ram sperm during epididymal maturation supported this hypoth¬ esis (158), but other similar studies did not (15,159). Such changes in saccharides also have been detected with lectins. One study found that WGA and RCA binding to rabbit sperm decreased as the sperm proceeded from caput to cauda epididymidis, although Con-A binding did not change (157). Other investigators reported a decrease in WGA binding to

hyrax sperm (160); they also reported decreased binding of RCA (161) and Con A (162) to ram sperm. However, an¬ other study found that washed rabbit sperm from the caput bound Con A poorly, while washed sperm from the cauda bound substantially more Con A than did unwashed sperm, particularly over the head (28). It was concluded that sperm¬ coating substances from the seminal fluid were masking lectin-binding sites on unwashed sperm. Additional studies indicated that Con-A binding to rat spermatozoa increased over the acrosome during maturation (163) and that the total amount of rat sperm surface material that will bind to a Con-A-affinity column approximately doubled during this process (164). Boar sperm studied using freeze-fracture techniques tran¬ siently showed regular geometric arrays of intramembranous particles in the plasma membrane over the anterior acrosome as they passed through the distal region of the caput epi¬ didymis. A different hexagonal array developed as sper¬ matozoa approached the cauda epididymis, appearing ini¬ tially at the margin of the acrosome and then extending to the postacrosomal region, but disappearing almost com¬ pletely upon ejaculation (33). In rat spermatozoa, plaques of parallel rows of particles appeared in the plasma mem¬ brane of the head at the distal end of the proximal caput epididymis, but largely disappeared by the proximal cauda epididymis (32). It has been suggested that these changes in patterns of intramembranous particles reflect changes in the nature of the sperm glycocalyx during epididymal transit (32).

Changes in Lipid Content The lipid content of whole sperm has been reported to decrease during epididymal maturation in boar, bull, ram, and rat (165-171); the cholesterol content of whole sperm has been reported to decrease in ram, rat, and hamster (172-174). The cholesterol/phospholipid ratio and the con¬ centration of phosphatidylserine, phosphatidylethanolamine, cardiolipin, and ethanolamine plasmalogen has also been reported to decrease in the ram (166,172). However, increases have been reported in the amount of sulfoconjugated sterols in hamster and human spermatozoa (173,175) as well as in unsaturated fatty acids in ram spermatozoa (172). When similar studies were carried out using plasma membranes isolated from boar spermatozoa, they confirmed that the amount of lipid in the plasma membrane decreased during epididymal maturation (143). Although there was a decrease in cholesterol, no significant change was seen in the cholesterol/phospholipid ratio. These studies also found decreases in phosphatidylethanolamine and phosphatidylinositol, as well as increases in dermosterol, cholesterol sulfate, phosphatidylcholine, and polyphosphoinositides. There was a decrease in the level of fatty acids and an increase in diacylglycerol, but no change in the degree of unsaturation of fatty acids. Preparations enriched for plasma

The Spermatozoon membrane from the anterior head region of ram sperm were particularly rich in ethanolamine and choline phosphoglycerides (176). The amount of dermosterol and ethanolamine in this region of the plasma membrane was reported to decrease, while the cholesterol to phospholipid ratio in¬ creased, during epididymal maturation. Changes in the amount and composition of lipids in the plasma membrane of spermatozoa during maturation are thought to explain why ejaculated sperm are more sensitive to cold shock than are testicular sperm (177-179). These changes may also account for the maturation-dependent de¬ crease in charge density at the phospholipid-water interface of ram spermatozoa, detected by electron spin resonance (177), and the decrease in membrane fluidity of bull sper¬ matozoa, seen by fluorescence polarization spectroscopy (180). Analysis of testicular and ejaculated ram spermatozoa by fluorescence recovery after photobleaching indicated that there are regional differences in the decrease of plasma membrane fluidity (65). During maturation, the diffusion rate of a fluorescent lipid analog increased in all regions of the sperm except the midpiece.

Changes in Protein Composition The sperm plasma membrane also undergoes major changes in protein composition during epididymal maturation (181-185). These changes occur by addition of new com¬ ponents to the sperm surface, by unmasking or modification of preexisting sperm surface moieties, or by loss of spermsurface components. They have been identified using bio¬ chemical approaches and antibodies to sperm-surface com¬ ponents or epididymal fluid components (Table 3). These changes are probably responsible for most of the previously described modifications in sperm-surface charge and lectin binding that occur during epididymal maturation, although changes in glycolipid composition could also be involved. Changes involving a particular component often occur within specific regions of the epididymis, suggesting that special¬ izations of epididymal function in these regions play im¬ portant roles in modifications of the sperm surface during epdidymai maturation. New components that appear on the sperm surface have been detected in biochemical studies using vectorial labeling of sperm from different regions of the epididymis. For ex¬ ample, the galactose-oxidase-tritium-borohydride proce¬ dure was used to label accessible D-galactose or A-acetylD-galactosamine and to detect a 37,000-dalton glycoprotein present on sperm from the cauda, but not on sperm from the corpus epididymidis of the rat (186). Similar results were seen when lactoperoxidase-catalyzed iodination was used to label surface tyrosine residues or when fluoresceinconjugated Con-A lectin was used to identify glycoproteincontaining bands on polyacrylamide gels (187). Other stud¬ ies using similar approaches have reported that the major change during rat sperm maturation is the increase in a

/

37

31,000- (188), a 32,000- (189), a 34,000- (190-192), or a 37,500-dalton (193) surface glycoprotein. In addition, bio¬ chemical approaches have been used to identify changes during maturation in spermatozoa from ram (194-197), bull (198), rabbit (199), boar (200), and chimpanzee (201).

Changes in Sperm-Surface Antigens Immunological approaches have also proven to be effec¬ tive for identifying differences in the surface composition of spermatozoa which arise during maturation (Table 3). Studies using these approaches have complemented bio¬ chemical studies, and the two procedures have frequently been used together. Antisera raised against either sperma¬ tozoa or epididymal fluid have often reacted with both, suggesting that an epididymal fluid component binds to the sperm surface. This has given rise to the term “sperm¬ coating antigens” for components found both in the fluid and on sperm (202-206). It is also possible that spermatozoa might shed components into the epididymal fluid during maturation which would be recognized by antisera to sperm or components in the fluid. However, several studies have identified components that are secreted by the epididymis and become bound to the sperm surface (see below). An antiserum to a 33,000-dalton acidic epididymal gly¬ coprotein (AEG) purified from the rat epididymis was used in immunohistochemical studies to demonstrate that AEG was secreted by principal cells in the epithelium of the caput and corpus epididymis and bound to sperm as they left the initial segment of the caput epididymis (207). AEG was found to have a slight stimulatory effect on sperm motility, but bovine serum albumin and rabbit serum were equally effective (208). Another study used an antiserum to partially purified specific epididymal proteins (SEP) in the rat (209) to confirm the organ specificity of SEP (210). The SEP were found by gel electrophoresis to be a complex mixture of proteins, including the PAS-positive glycoproteins “C” (22,400 daltons and pi 5.35-5.79) and “D” and “E” (37,000 daltons and pi 5.13 and 4.95, respectively) (210). The main region of synthesis was determined to be the caput epidi¬ dymis, and sperm were coated with SEP as they left the initial segment of the caput epididymis (211). Proteins D and E apparently bound weakly and could be removed from sperm by a modest increase in ionic strength of the medium (212) . The antiserum bound to the head of mature sperm and inhibited fertilization in artificially inseminated animals (213) . In another series of studies, different proteins isolated and purified from the rat caput epididymis were designated pro¬ teins “B” and “C” (16,000 daltons “D” (30,000 daltons), and “E” (32,000 daltons) (214-216). By using antisera and surface labeling methods, it was found that proteins D and E were absent from testis sperm but present on cauda sperm, whereas proteins B and C showed little, if any, binding to sperm (217,218). Antisera to D and E (redesignated as 27,000-

38

/ Chapter 2

and 28,000-dalton glycoproteins, respectively) bound to the head of sperm from the cauda, but not to sperm from the testis (218). Proteins B and C were biosynthetically labeled, along with a new protein “G” (37,000 daltons), and were found to bind to sperm from testis and cauda epididymis and to erythrocytes to a similar degree, suggesting that the binding was not specific to epididymal sperm (217). A sialoprotein, designated “SP” (37,500 daltons and pi 4.7), was also purified from rat epididymis and shown by immunohistochemistry to be present on the apical surface of cells in the caput and corpus epididymides, in the cy¬ toplasm of certain epididymal epithelial cells, and on sperm in the lumen of the epididymis (193). The antiserum did not cross-react with supernatants of epididymal homoge¬ nates from dog, rabbit, guinea pig, or human. Two extrinsic glycoproteins of 50,000 and 100,000 daltons were extracted from cauda sperm in the rat by washing with a high-ionicstrength buffer (219). Antisera to these glycoproteins reacted with the “periacrosomal” region of the head of cauda sperm, and immunohistochemical studies indicated that the antigen first appeared in the cytoplasm of prin¬ cipal cells in the proximal region of the cauda epididymis. Peptide maps suggested that the two glycoproteins had very similar composition and that the larger molecule may be a dimer of the smaller molecule. Both glyco¬ proteins bound lectins on Western blots, which indi¬ cated that their glycoconjugates contained mannose and A-acetylglucosamine (219). It is possible that the same sperm-surface maturation gly¬ coprotein has been identified in some of the different bio¬ chemical and immunological studies in the rat (217,220). It is the major glycoprotein present on sperm from the cauda epididymis, has sialic acid or galactose and tyrosine acces¬ sible to surface labels, and has an estimated mass between 24,000 and 37,000 daltons (186,187,189,190,192,220,221). It has been suggested that this molecule is present on caput sperm and that glycosylation during movement through the epididymis changes its pi from a range of 5.5 to 5.6 to a range of 5 to 5.5 and makes it more readily labeled by tritium borohydride (220). This molecule may be the a-lactalbumin that regulates galactosyl-transferase activity on the sperm surface (220). Although epididymal secretory products that become as¬ sociated with the sperm surface have been studied most extensively in the rat, studies in other species have shown that this is a general phenomenon. In the rabbit, epididymal glycoproteins first become associated with spermatozoa in the distal caput and proximal corpus epididymis (222); in the hamster, however, epididymal glycoproteins first be¬ come associated with the sperm in the proximal caput region (222). Other studies in the hamster also identified epidid¬ ymal glycoproteins that became deposited on the sperma¬ tozoa during maturation (223). In the human, an antiserum to ejaculated spermatozoa reacted specifically with epidi¬ dymal fluid and with epididymal sperm, but not testicular

sperm, and apparently identified an epididymal secretory product that binds to sperm (224). A few studies have used monoclonal antibodies to ex¬ amine sperm-surface changes during maturation. The ad¬ vantages of these probes are that they are highly specific and often identify antigens restricted to specific domains on the sperm surface. Two rat sperm-surface antigens that first appear in the caput epididymis were identified; one was present on the postacrosomal portion of the head (identified with monoclonal antibody 1B6), and the other was uni¬ formly distributed over the entire sperm surface (identified with antibody 2D6) (81). The latter antigen was susceptible to antibody-induced patching and was observed to be in¬ serted into the egg surface upon fertilization (207). Two antigens were also identified on hamster spermatozoa that were modified during maturation (208). One was over the head at an apparently higher concentration and on a greater percentage of cauda than on testicular sperm (recognized by antibody HM 3.1). The other was over the entire tail and appeared to be present in higher concentration on corpus sperm than on caput or cauda sperm (recognized by antibody HM 5.8). Antibody HM 3.1 blocked fertilization in vitro, whereas HM 5.8 reduced fertilization in vitro due to sperm agglutination. A 28,000-dalton antigen (identified with monoclonal antibody 1B3) was present over the entire surface of spermatozoa from the testis or caput epidi¬ dymis in the mouse, but was present only on the tip of the head of sperm from the cauda epididymidis (97). Other studies used monoclonal antibodies to identify four sperm-maturation antigens (SMA) in the mouse which were restricted to the anterior acrosome (SMA 1), posterior acrosome and midpiece (SMA 2), whole head (SMA 3), and whole tail (SMA 4) (84). SMA 4 was se¬ creted by the epithelium in the distal-caput-proximalcorpus region of the epididymis as an 85,000-dalton com¬ ponent and was trimmed to a 54,000-dalton component upon attachment to the surface of the flagellum (90,225,226). Another monoclonal antibody was used to show, in the mouse, that an antigen carrying the fucosylated lactosaminoglycan SSEA-1 was secreted by the caput epididymis and absorbed by sperm (227). The appearance of new antigens on the sperm surface during epididymal maturation might also occur because of unmasking or alteration of preexisting sperm-surface pro¬ teins. Although there do not appear to be well-documented examples of appearance due to unmasking, a possible case of this has been seen in the mouse. A monoclonal antibody was used to identify a 31,000-dalton antigen on the surface of the flagellum of spermatozoa from the cauda epididymis (91). The antigen gradually appeared on sperm as they passed through the mid-corpus region of the epididymis. It appar¬ ently was present first in higher-molecular-weight forms that were converted to the final protein, suggesting that the un¬ masking involved removal of moieties by limited proteo¬ lysis. Proteases (150,228,229) and glycosidases (230-232)

The Spermatozoon are present in the epididymis, and [3-galactosidase and (3glucoronidase are secreted in substantial amounts by rat epididymal cells in culture (233). In addition, the secretion of glycosidases in the epididymis is reported to be androgen dependent (234,235). Such enzymes might be responsible for unmasking of sperm-surface maturation components by partial degradation of surface glycoproteins. A proteinase inhibitor has been isolated from the epididymis in the mouse (236), and such molecules might be involved in regulating or limiting enzyme-mediated sperm-surface modifications in the epididymis.

Changes in Sperm-Surface Carbohydrates Another way that sperm-surface components are altered in the epididymis is by glycosylation. Epididymal homog¬ enates are rich in glycosyltransferases; dolichol, the polyisoprenoid carrier of oligosaccharide side chains used in as¬ sembly of A-glycosylated glycoproteins, is present in high concentration in the caput and corpus epididymis in the rat (237). The rat epididymis contains androgen-dependent glucosyl and mannosyl transferases (238), and the corpus has the highest (3-A-glucosaminidase, (3-A-acetyl-galactosaminidase, and (3-A-galactosidase activities in the epididymis (232). Also, A-acetylneuraminyl transferase activity is higher in caput than cauda epididymidis (239), although some sialoglycoproteins in the epididymal fluids are produced only in the cauda (188). There is evidence that the increase in negative surface charge by ram and bull spermatozoa in the epididymis is due to the addition of sialic acid groups to the sperm surface (9). Other studies have suggested that lactosaminoglycans present on the surface of testicular sper¬ matozoa are fucosylated by an epididymal fucosyltransferase (240). UDP-galactose:A-acetylglucosamine galactosyltransferase activity has been detected in fluids from the vas deferens of mice and rats (241-243) and in the rete testis fluid of rats (244). The galactosyltransferase appeared to be pro¬ duced in the testis and concentrated in the caput epididymi (244). Rat spermatozoa were capable of incorporating gal¬ actose from UDP-[14C]galactose into surface glycoproteins, and the addition of rete testis fluid increased the amount incorporated (245). The major incorporation was into 37,000and 23,000-dalton proteins, suggesting that sperm-surface components are altered during maturation by galactosylation of exposed A-acetylglucosamine residues (245). The 37,000dalton glycoprotein may have been the same maturationdependent sperm-surface glycoprotein identified in earlier surface-labeling studies (186). There is also galactosyltransferase activity on the sperm surface (69,247,248) which may be involved in sperm-sur¬ face modifications during epididymal maturation. This en¬ zyme appears to be different from the one in epididymal fluid (249). The galactosyltransferases may be regulated by

/

39

a-lactalbumin, which is apparently present in epididymal fluid (246) and on the surface of the sperm flagellum (250,251) and is produced by the epididymis (252). Galactosyltrans¬ ferase interacts with a-lactalbumin to form lactose synthe¬ tase, which catalyzes the transfer of galactose from UDPgalactose to glucose, forming lactose (253). It has also been hypothesized that the sperm-surface ga¬ lactosyltransferase binds to /V-acetylglucosamine on the zona pellucida as part of the fertilization process (249,254). The evidence for this comes from studies on a strain of mice that have a genetic predisposition for increased fertilizing ability and have elevated sperm-surface galactosyltransfer¬ ase activity (248). However, sperm from recombinant strains of these mice did not show elevated galactosyltransferase activity or increased fertilizing ability (249); competitive substrates for the enzyme, a-lactalbumin, and enzyme sub¬ strate analogs inhibited sperm binding to the zona pellucida (249,254). Also, enzymatic removal or unmasking of ter¬ minal A-acetylglucosamine residues produced coincident in¬ hibition or stimulation, respectively, of binding (249,254). Other studies have shown (a) that purified galactosyltrans¬ ferase produced a dose-dependent inhibition of sperm bind¬ ing to the zona pellucida and caused sperm bound to the zona pellucida to be released and (b) that monospecific antiserum to the enzyme produced a dose-dependent inhi¬ bition of sperm binding to the zona pellucida and concom¬ itantly blocked sperm galactosyltransferase activity (69). The enzyme was shown to be localized to the plasma mem¬ brane over the dorsal surface of the mouse-sperm acrosome (69).

Loss of Sperm-Surface Components Sperm-surface proteins also are apparently lost during epididymal maturation. A 110,000-dalton glycoprotein was the major surface component of rat testicular spermatozoa labeled with the glucose-oxidase-tritium-borohydride pro¬ cedure, but it was not labeled on spermatozoa from the cauda epididymis (189,190). In other studies on the rat, 94,000-, 72,000-, and 59,000-dalton components were iodinated on caput sperm but not on cauda sperm (221). In a study of mouse spermatozoa, a 28,000-dalton antigen was seen to be present over the entire surface of spermatozoa from the testis and caput epididymis, whereas those from the cauda epididymidis were labeled only over the anterior acrosome (97). However, it was not determined in these studies whether the components were (a) shed from the sperm surface, (b) modified in molecular weight because of processes such as limited proteolysis or addition of new moieties, or (c) still present but no longer accessible to surface labeling because of masking by other moieties. Im¬ munological studies have suggested that terminal A-acetylglucosamine on testicular spermatozoa was lost during ep¬ ididymal transit, probably as a result of galactosylation (27).

40

/ Chapter 2

Modification of the Sperm Plasma Membrane During Ejaculation and Capacitation Other alterations of sperm-surface composition may occur upon ejaculation (Table 4). These include: changes in sur¬ face charge (155,255-257) and in lectin binding (19,157); and possibly changes in lipid composition (142,256). In addition, there is absorption of blood-group antigens (259,260), histocompatibility antigens (261), and immu¬ nosuppressive factors (262). Sperm also become coated with proteins produced by accessory glands and absorbed from the seminal plasma. Human spermatozoa have been reported to be coated with lactoferrin (263,264), ferrisplan (265), PP5 (266), HSP-5 (267), pgl2 (268), and a basic 140,000dalton protein (269), all produced by the seminal vesicles. The latter protein cross-reacted immunologically with rat seminal-vesicle protein-IV (RSV-IV), a 17,000-dalton sem¬ inal-vesicle protein found in the rat (270). Another rat sem¬ inal-vesicle protein (50,000 dalton) was present in ejacu¬ lated, but not in epididymal, rat sperm (271). Similarly, a 20,000-dalton protein was present in ejaculated, but not in epididymal, sperm from the rabbit (272), and a 40,000dalton collagen-binding protein was present in ejaculated, but not in epididymal, rabbit sperm (273). It has been re¬ ported that 25,000- and 14,000-dalton proteins bind to bull sperm during ejaculation (198). Also, a 34,000-dalton com¬ ponent was present in ejaculated mouse sperm, in seminalvesicle fluid, and in sperm from the distal vas deferens, but not in epididymal sperm (78). Some of the proteins that bind to sperm upon ejaculation have been localized to specific sperm-surface domains. A 6,400-dalton proteinase inhibitor produced by mouse sem¬ inal vesicles was bound to a 15,000-dalton plasma-mem¬ brane component on the acrosome of ejaculated mouse sperm (274) . A 30,000-dalton prostate protein found in the dog was shown to be a proteolytic enzyme and to be present on the tail and postacrosomal region of ejaculated dog sperm (275) . Some of the components in the seminal plasma also appear to interact with each other and the sperm surface. Transglutaminase promoted covalent linking of spermidine to RSV-IV in vitro, and the protein was then able to bind

TABLE 4. Alterations of sperm surface upon ejaculation Alteration

References

Changes in sperm surface charge Changes in lectin binding Changes in lipid composition Integration of lipoproteins Absorption of blood group antigens Absorption of histocompatibility antigens Absorption of immunosuppressive factors Absorption of accessory gland secretions

155, 255-257 19, 157 142 258 259, 260 261 262 78, 263-276

to rat sperm (276). During ejaculation, a calcium-dependent transglutaminase from the prostate may modify the seminalvesicle protein to enhance its binding to the sperm surface (276). The plasma membrane undergoes additional modifica¬ tions in the female reproductive tract as spermatozoa gain the capacity to fertilize the ovum. Although the initial ob¬ servations were on changes that occurred in vivo (277,278), subsequent studies found that, in most species, comparable changes could be produced in vitro in defined media (279283). This allowed experimental studies to be carried out which identified modifications in sperm-surface charge (255,284), lectin binding (21,25,28,30,285), intramembranous particle distribution (10,30,50,51,61,286,287), membrane fluidity (77,288-290), lipid composition (61,286289,291,292), protein composition (272,273, 293-299), and antibody binding (206,226,298,300,301) during capacitation (Table 5). These membrane modifica¬ tions occur concomitantly with essential functional changes involving sperm motility, ion fluxes, and metabolism (7). Capacitation is a multistep process, but one aspect of it appears to be removal or modification of extrinsic compo¬ nents added to the sperm surface in the male reproductive tract. An example of this type of change is the loss of a mouse-sperm maturation antigen (SMA 4). This antigen is secreted by the epididymal epithelium and bound to the surface of the flagellum, but is shed by sperm as they ap¬ proach the surface of the egg (226). Another type of change with capacitation is the apparent release of constraints form¬ ing domain boundaries. An antigen present on the posterior tail of guinea-pig spermatozoa (PT-1) migrates onto the middle piece following capacitation, indicating that it is no longer constrained by the annulus (141). Following the ac¬ rosome reaction, another antigen (PH-20) migrates from the postacrosomal region to the inner acrosomal membrane (302). TABLE 5. Alteration of sperm plama membrane during capacitation Alteration Reduction in negative surface charge Modification in lectin¬ binding ability Changes in intramembranous particle distribution Changes in membrane fluidity Changes in lipid composition Changes in protein composition Changes in antibody binding Release of constraints forming domain boundaries

References 255, 284 21, 25, 28, 30, 285 10, 30, 50, 51, 61, 286, 287

77, 288-290 61, 286-292 226, 272, 273, 293-299 206, 226, 298, 300, 301 141, 302

The Spermatozoon

/

41

The serrated band at the posterior margin of the equatorial segment apparently does not pose a barrier to lateral dif¬ fusion at that time, but the antigen does not migrate to the midpiece, suggesting that the posterior ring remains a barrier to lateral diffusion.

sperm nucleus is unique, both in the amount of DNA present and in the composition of its nucleoproteins. The two meiotic divisions that occur during spermatogenesis produce a hap¬ loid genome, with only one member of each chromosome pair being present in the sperm nucleus.

THE SPERM HEAD

Nuclear Proteins

The head of the mammalian spermatozoon is occupied mostly by the nucleus and acrosome, but also contains cytoskeletal components and a small amount of cytoplasm (Figs. 3-6). The acrosome lies at the anterior end of the head, just beneath the plasma membrane, and is deeply indented posteriorly by the nucleus. Cytoskeletal compo¬ nents lie in the narrow space between the inner acrosomal membrane and the nuclear membrane and also just below the plasma membrane. Spermatozoa in most species have a spatulate head (Fig. 3), and the nucleus and acrosome are flattened in the plane of the anterior-posterior axis. The acrosome and nucleus are usually symmetrical structures; however, in some animals, protrusions of the acrosome extend perpendicularly to the flattened plane of the sperm head (Fig. 4). In rodent spermatozoa, which have a falci¬ form-shaped head, the acrosome overlies the convex margin of the nucleus. Although spermatozoa are uniform in size and shape in most species, in the human there is often variability in the size and shape of the head of some sperm, even in fertile individuals (303,304).

The major nuclear proteins associated with sperm DNA are protamines (305). These are relatively small (27-65 amino acids) highly basic proteins rich in arginine and cys¬ teine (306,307). The mRNA encoding for mouse protamines are synthesized in spermatids, indicating that protamines are products of the haploid genome (308). The highly con¬ densed protamine-DNA complex is stabilized by disulfide bonds between the protamines. Most mammals have only one protamine, but mice and humans are reported to have two (309-311). There are two general models showing how protamines associate with DNA (312). One suggests that protamines are present in an extended configuration and lie in the major or minor groove of the DNA double helix. They are presumed to cross-link the chromatin by forming covalent disulfide linkages with protamines on nearby DNA (313). The other model suggests that protamine is packaged into a-helical cylinders (314). These are thought to lie in the major or minor DNA groove and to facilitate orderly DNA condensation and subsequently to cross-link with neighboring cylinders to effect stabilization. Although both models are based on physical data and presume the lack of nucleosomes in sperm nuclei, some morphological studies have suggested that nucleosomes are present (315,316). In addition, studies using freeze-fracture (31,34,37), birefrin¬ gence (317), and physical methods (318) have suggested

The Sperm Nucleus The volume of the sperm nucleus is less than that of somatic cells, and its chromatin is highly condensed. The

FIG. 3. Shape of the head of sper¬ matozoa from different species. The falciform heads of mouse, rat, and hamster sperm are viewed laterally, whereas the spatulate heads of hu¬ man, rabbit, and guinea-pig sperm are viewed dorsoventrally. (Adapted from ref. 184.)

GUINEA PIG

42

/ Chapter 2

FIG. 4. Cross-sectional views of the heads of sper¬ matozoa from different species. Guinea pig, rabbit, and human sperm heads are seen in the sagittal plane, whereas the hamster sperm head is seen in the coronal plane. The acrosome and nucleus of rabbit and human spermatozoa are symmetrical, but those of guinea pig and hamster are asymmetrical in these planes.

RABBIT

that the chromatin in the spermatozoa of some mammals is stacked in lamellar plates. However, the chromatin of mouse and human spermatozoa appears to have a random fibrogranular pattern (44,319). Additional studies will be re¬ quired to fully understand how protamines and other nucleoproteins (305,312) interact with DNA in sperm nuclei. Nuclear Envelope The sperm nucleus is enclosed by an unusual nuclear envelope. Over most of the nucleus, nuclear pores are absent and the two membranes of the nuclear envelope lie only 7 to 10 nm apart. However, caudal to the posterior ring, in the so-called “redundant nuclear envelope,” nuclear pores are abundant and arranged in a hexagonal pattern; the two membranes are 40 to 60 nm apart as in most other cells (31,34,44,52,348). The membranes of the anterior part of the nuclear envelope contain a rich array of random intramembranous particles (320), whereas the closely apposed membranes of the nuclear envelope in the implantation fossa contain large closely packed particles surrounding particlefree areas (31). Nuclear Lamina The nuclear lamina is a protein meshwork lining the inner surface of the nuclear envelope. It is thought to form the

skeletal framework of the nuclear envelope and to serve as an anchoring site for chromatin (321-325). The nuclear lamina contains three closely related proteins: lamins A, B, and C. Lamins A (70,000 dalton) and C (60,000 dalton) have similar peptide maps but different antigenic epitopes. However, some antibodies recognize lamin B (67,000 dal¬ ton) as well as lamins A and C (326-328), indicating that all three share sequence homologies. Earlier immunohistochemical studies indicated that mouse spermatozoa did not contain proteins recognized by antibodies to lamins (329,330). However, a more recent study found that several other antibodies to lamins did react with nuclei of mouse spermatozoa (331). Immunostaining patterns were variable and were dependent on fixation and extraction conditions. The results indicated that an epitope recognized by antibody to lamin B was present over most of the sperm nucleus, that epitopes of lamin G (identified with an antibody to clam nuclei) were present on the convex margin of the nucleus, and that an epitope of lamins A and C was present on the convex margin, the anterior part of the concave margin, and adjacent to the implantation fossa (331). It was established recently that the lamins have extensive sequence homology with intermediate filament proteins (332,333) and appar¬ ently belong to the same family of structural proteins. A male germ-cell-specific lamin has been identified in Xenopus (334), and it has been suggested that a germ-cellspecific lamin may be present in mouse sperm (331). Al-

The Spermatozoon

/

43

though biochemical evidence for the presence of lamins in the mammalian sperm nucleus is lacking, it appears likely that lamins or related proteins contribute to the structure of the sperm nucleus.

Cytoskeleton of Sperm Head

FIG. 5. Cytoskeleton of head of the hamster spermatozoon. The subacrosomal cytoskeleton (perforatorium) lies between the acrosome and the anterior part of the nucleus and extends to the tip of the falciform head. The para-acrosomal cyto¬ skeleton lies between the plasma membrane and the acro¬ some on the concave margin of the sperm head. The postacrosomal cytoskeleton surrounds the posterior part of the nucleus, lying between the nuclear envelope and the plasma membrane of the postacrosomal domain.

Some cytoskeletal components of spermatogenic cells were discussed earlier in this chapter. Other cytoskeletal struc¬ tures have been identified in the head of mammalian sper¬ matozoa (Table 6) lying between the nucleus and the ac¬ rosome (subacrosomal cytoskeleton), between the plasma membrane and the postacrosomal portion of the nucleus (postacrosomal cytoskeleton), and between the plasma membrane and the anterior tip and convex surface of the acrosome of falciform sperm (para-acrosomal cytoskeleton) (Fig. 5). The subacrosomal and postacrosomal cytoskeletons can be isolated together (319) and have been referred to as the perinuclear theca (312). This complex resembles the nuclear matrix that has been isolated from somatic cells, but, unlike the nuclear matrix, it lies external to the nuclear envelope (319). The isolated perinuclear theca contains a diverse population of proteins ranging from 8,000 to 80,000 daltons, but the major proteins present were 13,000, 15,000, 16,500 and 25,000 daltons (319). The isolated perinuclear theca retains the shape of the sperm nucleus in the absence of DNA and the acrosome. It has been suggested that the perinuclear theca may be an extrinsic determinant of nuclear shape (312). The subacrosomal cytoskeleton (subacrosomal material, perinuclear material, perforatorium) is composed of amor-

FIG. 6. Structure of the head of guinea-pig and human sper¬ matozoa. The plasma membrane in the anterior acrosomal and equatorial segment domains overlies the outer acrosomal membrane. The inner acrosomal membrane, in turn, overlies

the nuclear envelope. The acrosome is thinner in the equatorial segment region than in the anterior acrosome region. The pos¬ terior ring separates the postacrosomal domain of the head from the middle-piece domain of the flagellum.

44

/ Chapter 2 TABLE 6. Sperm cytoskeletal components Component

Head Subacrosomal cytoskeleton (perforatorium) Postacrosomal cytoskeleton (postacrosomal sheath, postnuclear sheath, postacrosomal dense lamina, postnuclear cap, postnuclear body) Para-acrosomal cytoskeleton (filamentous cytoskeletal complex) Connecting piece Capitulum Segmented columns Tail Axoneme Outer dense fibers

Satellite fibers Fibrous sheath

References

335-346

34, 37, 41, 42, 44, 52, 121, 131, 319, 336, 346352

121, 122, 341, 342, 349, 352

52, 425, 426, 429 52, 418, 429 308, 433, 434, 436, 437 52, 337, 362, 425, 429, 450, 454-465, 467472 52, 336, 459, 474, 476 2, 303, 336, 337, 344, 457, 458, 473, 477-486

phous, electron-dense material situated in the narrow space between the inner acrosomal membrane and the outer mem¬ brane of the nuclear envelope (335). It is an extensive feature in rodents such as the rat, mouse, and hamster, which pos¬ sess falciform sperm heads (336), but is a minor component in sperm heads of other mammalian species (337,338). It has been referred to as the perforatorium in rodents because of its similarity in appearance to a structure in toads (339) and birds (340), thought to have a mechanical role in egg penetration (341,342). Earlier studies determined that the subacrosomal cytoskeleton was highly resistant to solubi¬ lization (343), apparently because of extensive disulfide bond formation (337). Advantage was taken of the insolubility of the subacrosomal cytoskeleton to devise a procedure for its isolation from rat spermatozoa (335,344). This allowed confirmation of earlier findings that the subacrosomal cy¬ toskeleton became more resistant to solubilization during epididymal transit (335,337). It also allowed the isolated subacrosomal cytoskeleton to be characterized biochemi¬ cally, and it was found to consist of a 13,000-dalton protein containing 6.5% cysteine (335). The subacrosomal cyto¬ skeleton is first visible in early spermatids as a dense layer between the forming acrosomal granule and the nuclear envelope. It extends peripherally over the anterior pole of the nucleus, just ahead of the advancing acrosome, as the sperm head elongates during spermiogenesis (345,346).

The postacrosomal cytoskeleton (postacrosomal sheath, postnuclear sheath, postacrosomal dense lamina, postnuclear cap, postnuclear body) lies between the nuclear en¬ velope and the plasma membrane of the postacrosomal seg¬ ment of the sperm head (34,52,336,347,348). It abuts the posterior end of the subacrosomal cytoskeleton anteriorly, at the posterior margin of the acrosome. and extends caudally to the posterior ring. It has a periodic substructure (34,39,121,131,348,349) and, in bull sperm, is composed of closely associated 10- to 12-nm filamentous elements aligned parallel to the long axis of the head (350). In ad¬ dition, the basal region contains coarse striations, formed by short rows of large particles (31,34,37,41,42,44,52,131). The basal region corresponds to a zone of close adhesion between the plasma membrane, the postacrosomal cyto¬ skeleton, and the nuclear envelope (350). Postacrosomal cytoskeleton formation begins in late spermiogenesis (346), with dense material appearing just caudal to the posterior margin of the acrosome, coincident with the formation of the manchette (351). The postacrosomal cytoskeleton ap¬ pears to assemble as the manchette moves caudally to form the posterior ring (352). This portion of the sperm-head cytoskeleton has not been isolated successfully, but it pre¬ sumably contains some of the perinuclear theca proteins that have been identified (319). The para-acrosomal cytoskeleton (filamentous cytoske¬ letal complex) has been identified recently in hamster sperm as a tripartite structure, consisting of a cone at the anterior tip and a bifurcated sheet on the convex surface of the head, lying between the acrosome and the plasma membrane (122). It is formed of filaments similar in size to intermediate filaments. Filaments have also been seen along the convex side of the acrosome of vole sperm (121), and striations were noted along the ventral surface of the acrosome of rat sperm (349), suggesting that the para-acrosomal cytoskel¬ eton may be present in these species. Sperm-head cytoskeletal structures have been suggested to have mechanical (341,342) and functional roles (350,352) in fertilization, and other roles are possible. The subacro¬ somal and para-acrosomal cytoskeletons are well developed in sperm possessing a falciform head and might serve as scaffolding to form or maintain this shape. However, the postacrosomal cytoskeleton is slightly more prominent in sperm containing a spatulate head and may provide stiff¬ ening and support for the posterior part of the sperm head and prevent its flexure during sperm motion. The function of the sperm-head cytoskeletal structures may be better understood once their composition is better defined. The major proteins forming cytoskeletal structures include tub¬ ulin, actin, and intermediate filament proteins. Microtubules are not present in these structures and although actin has been identified with antibodies in the postacrosomal region of testicular sperm, it was not detected in epididymal sperm (115,116,119,120). Structures similar in appearance to in¬ termediate filaments have been identified in the postacro¬ somal (350) and the paracrosomal (122) cytoskeleton. How-

The Spermatozoon

ever, antibodies to intermediate filaments do not react with these regions of the sperm head (127,350). This suggests that the sperm-head cytoskeletal structures are composed of unique proteins or of germ-cell-specific isoforms of cyto¬ skeletal proteins that lack antigenic determinants present on related proteins in somatic cells. However, sperm-head cy¬ toskeletal proteins may be related to flagellar cytoskeletal proteins. In sterile mutant mice, it is often the case that both the head and flagellum of sperm are defectively formed (353,354).

The Acrosome The acrosome originates from the Golgi complex in the spermatid and contains enzymes necessary for the sperm to penetrate to and/or fuse with the plasma membrane of the egg to achieve fertilization. This membranous structure sits as a cap over the nucleus in the anterior part of the sperm head.

Structure of Acrosome The inner acrosomal membrane is closely applied to the anterior part of the nuclear envelope; the outer acrosomal membrane underlies the plasma membrane. The acrosome consists of two segments, the anterior acrosome (acrosomal cap) and the equatorial segment (posterior acrosome), which immediately underlie the plasma membrane domains with the same names (Fig. 6). During the acrosome reaction the outer acrosomal membrane and the plasma membrane fuse and vesiculate, and most of the acrosomal contents are dis¬ charged. The inner acrosomal membrane and the equatorial segment persist until sperm-egg fusion in most species (7). The acrosome varies widely in size and shape in different species (Figs. 3 and 4), and the distribution and relative prominence of these two segments differ accordingly (336). The equatorial segment forms a band that approximately overlies the equator of the head of spatulate spermatozoa. In sperm possessing a falciform head, the equatorial segment may cover much of the lateral surface of the head. However, in species containing a discoid sperm head flattened in the plane perpendicular to the axis of the tail (“carpet tack¬ shaped head”), such as the wooly opossum (355), an equa¬ torial segment may not be identifiable. The portion of the anterior acrosome which extends beyond the anterior margin of the nucleus has been referred to as the apical segment, and the portion overlying the nucleus has been referred to as the principal segment (52). In the human, monkey, bull, boar, rabbit, and bat the acrosome is relatively small, with no appreciable extension beyond the nucleus, whereas in guinea pig, chinchilla, and ground squirrel the acrosome has a large apical segment (112,140,336,356). Electron microscopy revealed (a) that the acrosome, par¬ ticularly the apical segment, often has a much more complex

/

45

shape than is obvious by examining sperm smeared onto a slide and (b) that the shape of the acrosome is characteristic of the species (336,356). The final shape of the acrosome may be influenced by (a) extrinsic forces generated by cy¬ toskeletal elements in the spermatid and/or Sertoli-cell cy¬ toplasm (122,134) or (b) by forces intrinsic to the nucleus (357). However, in some species, it also appears that forces intrinsic to the acrosome may be involved. Acrosomes of guinea-pig and chinchilla sperm continue to undergo mor¬ phological differentiation after spermatogenesis, and the de¬ finitive shape is not achieved until sperm reaches the distal portion of the epididymis (112,356). As might be expected from species differences, genetic factors also influence ac¬ rosome formation and shape. Structurally defective acro¬ somes form in pink-eyed sterile mutant mice (358,359), whereas acrosomes fail to form in blind sterile mutant mice, even though proacrosomal granules, the manchette, and fla¬ gellar structures form and some nuclear elongation and chro¬ matin condensation occurs (354). The acrosome sometimes shows an internal lamellar or crystalline structure (31), an ordered substructure (360), or a cobblestone-like pattern (42,131). A 4.2-nm periodicity was present in the cortical region of the acrosome of rat sperm, lying just deep to the outer acrosomal membrane on the convex surface (31,361). A similar pattern was also reported in the acrosome of human sperm (362). Other evi¬ dence that the acrosome has a substructure comes from studies on hamster spermatozoa disrupted by nitrogen cav¬ itation. This treatment resulted in loss of much of the ac¬ rosomal matrix, but components immediately underlying the outer acrosomal membrane remained intact (122). These components were present in two areas: One was a larger and looser layer of fibrous material over the dorsal and lateral surfaces of the acrosome, whereas the other was a more compact fibrous layer adjacent to the anterior margin of the acrosome. The membrane of the acrosome, particularly the equa¬ torial segment, contains particles that form a crystalline array, giving the membrane a highly regular, granular ap¬ pearance. These may be an indication of ordered structure in the underlying acrosomal components. Such features have been reported in rabbit (34), bull (39,131), human (44), rat (31,363), mouse (320), guinea pig (31,39,363), degu (39), and rhesus monkey (39). This pattern has been seen by freeze-etch, by freeze-fracture, and on replicas of air-dried and critical-point-dried spermatozoa. Although the outer ac¬ rosomal membrane appears to be fragile and easily displaced or disrupted at the time of the acrosome reaction, it also has a thickened appearance because of an electron-dense coating on the inner surface (364-368). This inner surface coat of the outer acrosomal membrane has been isolated from bull sperm and shown to be composed of three highmolecular-weight glycoproteins (290,000, 280,000 and 260,000 daltons) as well as 115,000-, 81,000-, 58,000-, and 46,000-dalton proteins. In addition, there was a set of proteins between 34,000 and 12,000 daltons (368). Lectins

46

/ Chapter 2

were found to bind to the inner surface of the membrane; WGA was observed to bind to the 46,000-dalton compo¬ nent. It was suggested that glycosylated molecules at this site may help to stabilize the membrane or play a functional role in the membrane fusion events of the acrosome reaction (368). The 200,000- and 58,000-dalton components were phosphorylated in a cAMP-independent manner, whereas the proteins between 34,000 and 12,000 daltons appeared to include calmodulin-binding proteins (368). Inner acrosomal membrane development begins in early spermatids when the membrane of the proacrosomal granule abuts and then flattens against the nuclear envelope. This attachment spreads over the apical end of the nucleus during acrosomal development and nuclear elongation (369,370). The inner acrosomal membrane becomes exposed at the sperm surface and continuous with the plasma membrane when the acrosome reaction occurs. The inner acrosomal membrane in mouse and rabbit spermatozoa is quite resistant to chemical and physical disruption, including treatment with nonionic detergents and sonication (371,372). How¬ ever, boar-sperm inner acrosomal membrane is sensitive to proteinase treatment (373), and lectins bind to the inner acrosomal membrane of sperm from hamster (30,374) and guinea pig (25), indicating that glycoproteins are present. Bridges 7 nm wide and with 7-nm center-to-center spacing were reported to be present between the inner and outer acrosomal membranes in boar sperm, apparently holding these structures together (373). However, the inner acro¬ somal membrane appears to be fluid because antigens rec¬ ognized by monoclonal antibody PH-20 migrate from the plasma membrane of the postacrosomal region of the guineapig sperm head to the inner acrosomal membrane following the acrosome reaction (302,375). It has been suggested that the inner acrosomal membrane is associated with an exten¬ sive scaffolding network, possibly transmembrane in nature, in the equatorial segment (338).

Contents of Acrosome The acrosome is a large zymogen-secreting granule whose contents are discharged during the acrosome reaction. How¬ ever, it also contains enzymes typically found in primary lysosomes (Table 7). It has been described as a specialized lysosome (150,376), even though it apparently does not serve the usual lysosomal role of breaking down materials scavenged from the cytoplasm or entering the cell. The enzymes that are unique to the mammalian sperm acrosome are acrosin and hyaluronidase, and both are major constit¬ uents of the acrosome. Acrosin is a trypsin-like serine proteinase, but it differs from similar enzymes in other tissues in molecular weight, substrate, and inhibitor specificities, indicating that it is probably a spermatogenic cell-specific isozyme (377-381). It was observed that fertilization was decreased by soybean trypsin inhibitor (382) and that pancreatic and seminal plasma

TABLE 7. Enzymes in the acrosome Enzyme p-/V-Acetylglucosaminadase Acid phosphatase Acrosin Arylamidase Arylsulfatase A Aspartylamidase Calpain II Cathepsin-D-like protease Collagenase-like peptidase Esterases, nonspecific (3-Glucuronidase Hyaluronidase Neuraminidase Phospholipase A

References 416 415 184, 377-398 412 414 413 409 408 407 411 417 399-406 410 376

trypsin inhibitors block fertilization in vitro (383), indicating that acrosin is required for fertilization. The enzyme is present in the acrosome as proacrosin, which is con¬ verted to the active form during the acrosome reaction (377— 379,381,384). Immunolocalization studies have suggested that acrosin is present mainly in the anterior acrosome in human, boar, bull, and rabbit spermatozoa (385), but others have reported that it is localized to the inner acrosomal membrane (386-388). However, acrosin is rapidly released following the acrosome reaction, and it has been suggested that the bulk of the acrosin may be in the soluble acrosomal matrix (184). Although acrosins from various species are quite similar functionally (389-391), there are species differences in proacrosin and acrosin at the molecular level. Boar proac¬ rosin was present in 55,000- and 53,000-dalton forms; ac¬ rosin was present in 49,000-, 34,000-, and 25,000-dalton forms (377,392). Hamster-sperm proacrosin was present in forms between 56,000 and 51,000 daltons acrosin was pres¬ ent in multiple forms in two groups between 56,000 and 49,000 daltons and between 40,000 and 30,000 daltons (393). Rabbit proacrosin was present as a 27,300-dalton dimer (381), and a 38,000-dalton acrosin has been isolated from ram sperm (394). A monoclonal antibody (C 11 H) to human-sperm acrosin reacted with a 50,000-dalton protein and with several other proteins in the 34,000 to 24,000dalton range extracted from human sperm (395). The an¬ tibody also reacted with the 55,000- and 53,000-dalton forms of acrosin isolated from boar sperm and was used to deter¬ mine that acrosin is first detectable in step-9 spermatids during mouse spermatogenesis (396). Other studies have indicated that acrosin first appears in spermatids during sper¬ matogenesis in the bull, ram, rabbit, boar, and human (395,397,398). The hyaluronidase present in the acrosome has been re¬ ported to be distinct from the common lysosomal form and, like acrosin, appears to be a spermatogenic cell-specific isozyme (399^401). This glycosidase is abundant in aero-

The Spermatozoon

somes (402) and is located predominantly in the anterior acrosome in bull (403,404) and ram sperm (405). However, ram sperm denuded of the plasma membrane and outer acrosomal membrane still have half the hyaluronidase of an intact acrosome (406), and it has been suggested that some of the enzyme may be bound to the inner acrosomal mem¬ brane (405). Ram-sperm hyaluronidase was found to be a 62,000-dalton protein (400), whereas bull-sperm hyaluroni¬ dase was a 60,000-dalton protein and had a pH optimum of 3.8 (399). Other hydrolytic enzymes have been detected in the ac¬ rosome (Table 7). Proteinases reported include a 110,000dalton collagenase-like peptidase, with a pH optimum of 7.5 (407), and a cathepsin D-like protease (408). Antiserum to a calcium-activated neutral proteinase, calpain II, reacted with porcine-sperm acrosomes by indirect immunofluores¬ cence and recognized an 80,000-dalton subunit of the en¬ zyme on immunoblots (409). A neuraminidase (410), non¬ specific esterases (411), arylamidase (412), aspartylamidase (413), arylsulfatase A (414), acid phosphatase (415), (3-7Vacetylglucosaminidase (416), (3-glucuronidase (417), and phospholipase A (376) have also been reported to be present in the acrosome.

THE FLAGELLUM The flagellum of the mammalian spermatozoon consists of four distinct segments: the connecting piece (neck), the middle piece, the principal piece, and the end piece (Fig. 1). The main structural features of the flagellum of the mammalian spermatozoon are the axoneme, the mitochon¬ drial sheath, the outer dense fibers, and the fibrous sheath (Table 6). The axoneme is composed of a “9 + 2” complex of microtubules which extends the full length of the fla¬ gellum. The middle-piece segment contains the mitochon¬ drial sheath, while the principal-piece segment contains the fibrous sheath. The mitochondria underlie the plasma mem¬ brane in the middle piece, while the fibrous sheath underlies the plasma membrane in the principal piece. The outer dense fibers extend from the connecting piece into the posterior part of the principal piece and are situated between the mitochondria and the axoneme in the middle piece, and between the fibrous sheath and the axoneme in the principal piece. The base of the flagellum abuts the nucleus at the junction between connecting piece and head (140,418,419). The flagellum provides the motile force necessary for the sperm to reach the egg surface and achieve fertilization. The different elements of the flagellum are involved (a) in generating and shaping the waves of bending that produce this force and (b) in propagating the waves from the base to the tip. The flagellum contributes most of the length of the mammalian spermatozoon. For example, the sperma¬ tozoon of the human is about 60 p.m long, and the head is only 4 to 5 |xm of this length (303). However, the length of the spermatozoon varies considerably between species.

/

47

The spermatozoon in the rabbit is 46 |xm long; that in the mouse is 120 p.m long, that in the rat is 190 p,m long, and that in the Chinese hamster is 250 p,m long (420). The human-sperm flagellum, which is more than 1 p,m in di¬ ameter in the connecting-piece segment, tapers progres¬ sively toward its posterior tip (303).

Connecting Piece The main structural components of the connecting piece (Fig. 7) are the capitulum (a dense fibrous plate-like struc¬ ture that conforms to the shape of the implantation fossa) and the segmented columns (52). The implantation fossa is formed by a specialized region of the nuclear envelope and a dense plaque of material on the cytoplasmic surface of the nuclear envelope, the basal plate. The interspace be¬ tween the two membranes of the nuclear envelope in this region is traversed by regular periodic densities 6 nm wide and 6 nm apart (52). Freeze-fracture studies indicated that the membrane of the nuclear envelope lining the implan¬ tation fossa contains a dense population of large regularly spaced intramembranous particles surrounding a particlefree region of membrane (31). Fine filaments traversing the narrow region between capitulum and basal plate presum¬ ably are responsible for attaching the capitulum of the fla¬ gellum to the basal plate of the head (320,362). This region between capitulum and basal plate appears to be the site of cleavage of heads from tails following trypsin treatment (23,421), but decapitation of sperm with primary amines or sodium dodecyl sulfate usually results in cleavage between inner and outer nuclear membranes, next to the basal plate (422). Heads and tails can also be separated by sonication, but the cleavage site is not predictable (423,424). The basal plate and capitulum are composed of proteins that are soluble in ionic detergent containing a disulfide-bond reducing agent (425,426). Although the composition of these structures has not been determined, they may be related to ciliary rootlets (52), which contain a 250,000- and 230,000-dalton protein dimer called ankyrin (303). A genetic defect is sometimes seen in bulls in which the flagella of most mature sperm are detached from the heads (427). Detachment begins during late spermatogenesis and continues in the epididymis; the resulting detached flagella are motile, metabolically active, and able to penetrate cervical mucus (428). Extending posteriorly from the capitulum are two major and five minor segmented columns 1 to 2 p,m in length. The two major columns split into two columns each and, along with the other columns, fuse to the nine outer dense fibers extending throughout much of the remaining length of the flagellum (Fig. 9). However, the segmented columns and outer dense fibers have different origins, and the con¬ tinuity between them develops late in spermiogenesis (418,429). The segmented columns of the connecting piece are cross-striated, with a periodicity of 6.65 nm between segments; and each segment, in turn, has nine or 10 hori-

48

/ Chapter 2

FIG. 7. Connecting piece of the flagellum. The basal plate is adherent to the nuclear envelope, defining the implantation fossa and forming the site of attachment of the flagellum to the sperm head. The connecting piece is topped by the capitulum, with the segmented columns extending from it caudally to fuse with the outer dense fi¬ bers. (Modified from ref. 432.)

zontal bands (429). During development of the flagellum, a transversely or obliquely oriented proximal centriole lies between the longitudinally oriented distal centriole and a depression in the capitulum (429). The distal centriole is continuous with the axoneme and is surrounded by other accessory structures of the connecting piece. In many spe¬ cies, the distal centriole disintegrates late in spermatogenesis and the proximal centriole is retained (140,362,418,430,431). It has been suggested that the proximal centriole is the center of sperm motility (418). However, the proximal centriole disintegrates during the latter part of spermiogenesis in some species with normal motility (429,432), indicating that it is not required for generation of the flagellar beat. It appears that the centrioles serve as organizing centers for the for¬ mation of the axoneme and segmented columns, but are not required for initiation or propagation of waves of bending along the tail (52). The connecting piece joins the middle piece distally at the beginning of the mitochondrial sheath.

Axoneme The axoneme or axial filament complex of the mammalian sperm tail (Fig. 8) has the same organization as that in cilia and flagella, present in most plants and animals. It consists

of two central microtubules surrounded by nine microtubule doublets (433). Each doublet consists of a complete A mi¬ crotubule, onto which is attached a “C-shaped” B micro¬ tubule. Two arms extend from the A microtubule toward the B microtubule of the adjacent doublet. When axonemes are viewed from base to tip, the arms project clockwise (434). In the rat spermatozoon, the central pair of micro¬ tubules extend into the connecting piece to the capitulum, whereas the other microtubules appear to end on the rem¬ nants of the base of the distal centriole (432,435). Spokes radiate from the central pair of microtubules to the outer doublets in a double helix around the central tubules (436). The doublets are numbered one through nine, with number one being the doublet situated on a plane perpendicular to that bisecting both microtubules of the central pair. Doublet number two is adjacent to the arms of doublet number one, and so on. The microtubules are composed of “a-tubulin” and “13tubulin,” closely related proteins of approximately 56,000and 54,500 daltons respectively (437). At least one new form of both a- and (3-tubulin first appears during sper¬ matogenesis in spermatids (437,438), indicating that postmeiotic tubulin gene expression occurs in these haploid cells (308). A cDNA for a-tubulin appears to differ in sequence from other a-tubulin genes, suggesting that a unique sper-

The Spermatozoon

/

49

FIG. 8. The axoneme in the end-piece of the sperm flagellum. Nine outer doublets of microtubules sur¬ round an inner doublet of microtubules. The outer doublets consist of A and B microtubules. Inner and outer dynein arms extend from the A microtubule toward the B microtubule of the adjacent pair.

matogenic cell a-tubulin isoform is present in spermatids (438). It has not yet been determined whether this tubulin is present in the microtubules of the flagellum and might be involved in determining some of the special properties of the sperm axoneme. “Kinesin” is another interesting pro¬ tein associated with microtubules that has been identified recently (439). It is a 600,000-dalton complex, composed of 110,000- and 65,000- to 70,000-dalton subunits, which interacts with the surface of microtubules to effect move¬ ment. Isolated microtubules move over the surface of a slide coated with kinesin, and latex beads are moved along mi¬ crotubules in the presence of the protein. Kinesin has been shown to cause anterograde translocation of cell organelles along axonal microtubules, and recently a retrograde tran¬ slocator was characterized partially (440). Both transloca¬ tors require ATP, but differ from myosin or dynein in their structural and enzymatic characteristics (439,440). It will be interesting to learn whether kinesin is present in the mammalian sperm flagellum and whether it has a role in generating or propagating the flagellar wave. Other proteins originally identified in the axoneme of sea urchin sperm are probably also present in mammalian sperm. The arms of the microtubule doublets contain “dynein,” an approximately 500,000-dalton protein with ATPase activity (441). The ATPase activity is believed to be responsible for the sliding forces generated between adjacent doublets of microtubules during flagellar movement (442). ATPase has been extracted from mammalian spermatozoa, and ATPase activity has been identified by histochemistry on the arms of the doublets and at the junction between radial spokes and central tubules in human sperm (303,443). The links between adjacent microtubule doublets in sea urchin sperm contain “nexin,” a 165,000- to 150,000-dalton protein thought to constitute about 2% of the axonemal proteins (444). In addition, the sperm doublet microtubules of the sea urchin sperm flagellum can be dissociated with various treatments to produce ribbons of two to four pro¬ tofilaments (445) that have been further fractionated with chemical agents (446). The ribbons contain proteins that

have been named “tektins,” and the filaments that they form are 2 to 3 nm in diameter, highly insoluble, and have phys¬ ical and chemical properties similar to intermediate fila¬ ments (447,448). The filaments contain 55,000-, 51,000-, and 47,000-dalton proteins, and an antiserum to this set of proteins reacted with the entire length of the axoneme (449). They also appear to be laterally associated with the doublet microtubules, are probably structural components of the A microtubule wall, and may be involved in assembly or func¬ tion of microtubules (449).

Mitochondrial Sheath The mitochondria are helically wrapped around the outer dense fibers in the middle piece of the sperm tail (Fig. 1). They are generally arranged end-to-end, but the number of parallel helices, the number of gyres, and the length of the middle piece vary between species. In the mouse, the mi¬ tochondria are usually arranged in two parallel helices, with an average of 87 windings around the flagellum (450). The morphogenesis of this arrangement in the spermatid was found to occur by: (a) the migration of the annulus from the connecting piece down the flagellum to the beginning of the fibrous sheath, (b) the formation of a dextral helix of elongated mitochondria around the flagellum, (c) the division of these mitochondria into spherical mitochondria, and (d) finally the elongation and end-to-end apposition of these mitochondria into two tight sinestral helices (450). In the mouse, the mitochondria are usually of variable length and abut end-to-end at random along the helix. It was pos¬ sible by a genetic selection program to produce strains of mice with sperm midpieces longer and shorter than the 21p,m average midpiece length (451). In some species there is a precise order in the mitochon¬ drial sheath. In the common brown bat, there are two mi¬ tochondria of identical size in each turn of the gyre, and their ends always meet on a plane passing through the central pair of microtubules in the axoneme (132). Mitochondrial

50

/ Chapter 2

sheaths composed of a precise number of mitochondria of identical size and shape are present in the little brown bat (132), woolly opossum (452), and Chinese hamster (453). In rhesus monkey sperm there is a tendency for the de¬ marcation between mitochondria to occur in longitudinal register in alternate gyres (453). The length of the middle piece and the disposition of mitochondria in spermatozoa was determined for several species using surface replica methods (453). The middle piece in bull sperm was 12 |xm long, the number of mito¬ chondrial gyres was about 64, and there were usually three helices. Some mitochondria were found to extend parallel with the long axis of the flagellum into the connecting piece. The rabbit sperm midpiece was 8.5 p,m long and contained about 41 gyres of mitochondria in a quadruple or quintuple helix. The middle piece of rhesus monkey spermatozoa was about 10 p.m long with 40 gyres of mitochondria arranged in a single or double helix. The middle piece of rat sperm was 64 |i,m long and contained about 362 mitochondrial gyres. The large outer dense fibers in rat sperm produced a broad middle piece, and the gently spiraling mitochondria fitted together in intricate patterns. The middle piece in Chinese hamster sperm was about 100 |xm long, with the mitochondria apparently disposed in a double helix, each wrapping one-third the circumference of the quite broad middle piece. The degu sperm middle piece was 7 p.m long, with about 33 gyres of mitochondria in concentric rows of single or double helices. Finally, the guinea-pig spermatozoan middle piece was determined to be 9 |xm long, and the mitochondria were found to be arranged in irregular concentric patterns (453).

Outer Dense Fibers The outer dense fibers surround the axoneme, forming a “9 + 9 + 2” complex throughout the length of the middle piece and throughout most or all of the principal piece of the flagellum of mammalian spermatozoa (Figs. 1 and 9). Although similar structures are present in spermatozoa of nonmammalian species (2), it has not been determined whether they are related in composition or function to the outer dense fibers in mammalian spermatozoa. The basic structural plan of the sperm flagellum shows little variation in mammals, but the outer dense fibers differ considerably in size and shape between species. The fibers are often tear-drop shaped in cross section, with a rounded outer contour and an inner margin tapering toward a doublet of the axoneme (52). The dense fibers also differ among themselves in shape and size, with fibers 1,5, and 6 (and sometimes 9) being larger than the others. They are numbered corresponding to the adjacent microtubule doublet. The fibers are thickest in the proximal part of the middle piece and gradually taper in thickness toward the distal tip. They terminate in the proximal half of the principal piece in the human, macaque, and bat, but extend farther in the principal piece in the rat, hamster,

FIG. 9. Cytoskeletal components of the sperm flagellum. The axoneme extends from the connecting piece of the flagellum to the distal tip. In the principal piece of the flagellum, the axoneme is surrounded by the fibrous sheath, composed of longitudinal columns connected by circumferential ribs. The outer dense fibers extend from the connecting piece to the posterior part of the principal piece. They lie between the mitochondrial sheath and the axoneme in the middle piece and between the fibrous sheath and the axoneme in the principal piece.

guinea pig, and ground squirrel (52). In the human (454), bull, and rat (455), the dense fibers occupy 60% of the length of the principal piece. In most species, fibers 3 and 8 terminate in the first part of the principal piece, and their place is taken by inward extensions of the longitudinal col¬ umns of the fibrous sheath. The larger fibers (1,5, and 6) are usually the last to terminate (456). The formation of the outer dense fibers begins in sper¬ matids, with the appearance of a small dense fiber imme¬ diately adjacent to the outer aspect of each microtubule doublet (429,457,458). In the rat, fibers first appear in the most proximal part of the future middle piece in step 8 and extend into the principal piece by step 14 (459). During step 15 of spermiogenesis the outer dense fibers increase in diameter and change shape, with fibers 1,5, and 6 devel¬ oping ahead of the other fibers. During step 16 the fibers enlarge rapidly along their entire length and take on their nearly final form, with slight growth continuing during steps 17 to 19 (459). Radioautography following incorporation of labeled proline and cysteine suggested that rapid protein synthesis accompanied the rapid growth of the outer dense fiber during step 16 (459). On electron micrographs the fibers usually appear dense, with a slightly less dense cortical layer. This cortical layer

The Spermatozoon

stained intensely with phosphotungstic acid (460) and tannic acid (52) and, when fixed in the presence of ruthenium red, appeared to be composed of a single lamina of 6- to 7-nm globular particles (461). Surface replicas of rat- and mousesperm outer dense fibers showed striations in the outer con¬ tour of the cortex, with a periodicity of 40 nm (461,462); these striations had the same 70 to 80° “sinistral obliquity” as the mitochondrial helix in the middle piece (450). On replicas each striation appeared to be composed of a double linear array of 6- to 8-nm diameter globular subunits (461), and a central depression in each striation probably accounts for an apparent periodicity of 20 nm seen on negatively stained specimens (461,463). In the bull, 20-nm (464) and 50-nm (465) periodicity has been reported, and a 16-nm repeat was reported for human-sperm outer dense fibers (362). Studies on the rat have indicated that the periodic substructure is confined to the cortex (461); however, this substructure is reported to be present in both cortical and medullary elements in human and bull sperm (362,465). It has been possible to take advantage of the relative insolubility of fibrous components of the flagellum in the absence of disulfide bond-reducing agents to isolate outer dense fibers for biochemical analysis. Early studies reported that tail components contain keratin-like proteins rich in cysteine (466) and that disulfide bond cross-linking of pro¬ teins in the outer dense fiber increased during epididymal maturation (337,425). Initial studies on outer dense fibers isolated from rat sperm and analyzed by gel electrophoresis in the presence of disulfide-bond-reducing agents indicated that they contained 40,000-, 25,000-, 12,000-, and 11,000dalton proteins (467). A subsequent study indicated that the main proteins in the rat were 75,000, 35,000, 25,000, and 15,000 daltons (468). Another study reported that rat outer dense fibers contained major proteins of 87,000, 25,000, 19,000, and 12,000 daltons, as well as minor proteins of 71,000, 64,000, and 55,000 daltons (461). A more recent study indicated that the major proteins in the rat outer dense fibers were 87,000, 30,400, 26,000, 18,400, 13,000, and 11,500 daltons (469). Similar studies on bull-sperm outer dense fibers identified 55,000-, 30,000-, and 15,000-dalton proteins (465), later revised to be 75,000- to 72,000-, 57,000to 54,000-, and 27,000- to 24,000-dalton proteins (463). Several studies have confirmed that the outer dense fibers have a high cysteine content (461,463,469). In addition, it has been shown that there are differences between individual outer dense fiber proteins in amino acid composition, aminoterminal amino acid, net charge, and phosphorylation (461,463,469). The 87,000-dalton protein in rat outer dense fibers contained relatively large amounts of glutamic acid, lysine, alanine, and leucine, but had relatively small amounts of serine, glycine, and cysteine. However, the 30,400-, 26,000-, 18,400-, and 11,500-dalton proteins contained high amounts of cysteine, tyrosine, and proline plus glycine, but less glutamic acid. The 13,000-dalton component had a high content of glutamic acid and no histidine (469). The 30,400and 26,000-dalton proteins had similar amino acid com¬

/

51

position, histidine in the amino-terminal position, and sim¬ ilar peptide maps, suggesting that they share a common peptide chain (469). Other differences have been reported for the proteins that compose the outer dense fibers. Infrared spectrum analysis suggested that the higher-molecular-weight components have an alpha-helical configuration (463), whereas other studies indicated that the lower-molecular-weight components are capable of binding zinc to suifhydryl groups (468). Zinc is required for spermatogenesis (470), is incorporated into the flagellum in late spermatids (471), and is localized in the outer dense fibers (463). Isoelectric focusing indicated that the proteins have large charge heterogeneities, indicative of posttranslational modifications (469,472). Some studies have suggested that the outer dense fiber proteins are glycosylated (465), but other studies indicated that carbohydrates were either absent (461) or barely detectable, suggesting that they may be present as contaminants (469). All of the major polypeptides contained appreciable amounts of phosphate bound as phosphoserine, which could account for the charge heterogeneities, particularly in the higher-molecular-weight polypeptides (469). Early studies also suggested that tri¬ glycerides may be associated with the outer dense fibers (465,467). Satellite fibers are present in the flagellar matrix between the outer dense fibers (52,336,476). These fibers form in step 19 of spermiogenesis in the rat, just before spermatozoa are released from the seminiferous epithelium (459). They are present in limited numbers in most species, but are highly developed in the ground squirrel (336) and bandicoot (474), which have thick sperm tails (336). It has been sug¬ gested that they may be accessory tensile elements of the motor apparatus (336). The satellite fibers appear to arise by exfoliation from the free edge of the cortex on the sides of the outer dense fibers and have the same resistance to solubilization as the outer dense fibers (52). However, they have not been isolated separately from the outer dense fibers, and their composition is not known. An early speculation was that the outer dense fibers might be contractile because of their close association with the axoneme, their coincident appearance during phylogeny with internal fertilization, and a concomitant increase in the size of the mitochondrial sheath (336). This appeared to be sup¬ ported by (a) a report that the outer dense fibers were antigenically similar to actin (475) and (b) histochemical stud¬ ies that suggested that an ATPase was associated with the outer dense fibers (460). However, subsequent biochemical studies did not indicate a similarity between the composition of outer dense fibers and actin, myosin, or tubulin (461); also, other investigators did not detect ATPase activity in outer dense fibers (465,476). It now appears unlikely that the outer dense fibers play an active role in flagellar motion. However, stabilization of outer dense fiber proteins by abun¬ dant disulfide cross-linking may give them significant pas¬ sive elastic properties that serve to stiffen or provide elastic recoil for the sperm tail (52). By use of high-speed cine-

52

/ Chapter 2

matography, spermatozoa of mouse, human, rabbit, and opossum appeared relatively flexible, and as they beat they formed arcs with a small radius of curvature. Spermatozoa of rat and Chinese hamster appeared very stiff while beating and had a large radius of curvature (420). There was a correlation between the radius of curvature and the size of the dense fibers in these species, suggesting that the dense fibers might influence the form of the beat by determining the elastic properties of the sperm tail (420).

Fibrous Sheath The fibrous sheath defines the extent of the principal piece (Figs. 1 and 9), which is the longest segment of the flagellum and is the most effective in locomotion (336). It appears to be a cytoskeletal structure of the flagellum that is unique to spermatozoa in mammals and some birds. It closely un¬ derlies, but is not attached to, the plasma membrane (2). The fibrous sheath is a tapering cylinder formed by two longitudinal columns connected by circumferential ribs (Fig. 9). The columns are formed by longitudinally oriented, loosely packed filamentous structures 15 to 20 nm in diameter. The longitudinal columns run peripheral to microtubule doublets 3 and 8 of the axoneme and are attached to outer dense fibers 3 and 8 in the proximal part of the principal piece (473). Distal to the termination of these two dense fibers, the tapered central edges of the longitudinal columns attach to ridges projecting from microtubule doublets 3 and 8 (336). The columns lie approximately in the plane of the central pair of axoneme mjcrotubules throughout the principal piece segment and have been referred to as dorsal and ventral columns in sperm containing a falciform head (336). The size and shape of the longitudinal columns vary between species. They are narrow and inconspicuous in the common brown bat and guinea pig, but in the Chinese hamster and opossum they are large and result in prominent ridges that give the principal piece an elliptical profile in cross section (336). The ribs of the fibrous sheath are composed of closely packed, circumferentially oriented filaments (336). The ribs broaden toward their ends, where they merge with the lon¬ gitudinal columns and with each other. They are closely spaced and may bifurcate to anastomose with adjacent ribs. This occurs frequently in the mouse, resulting in broad bands instead of slender separate ribs. The ribs may also vary in shape and size, being broad and flat in the bat and greatly expanded at their ends in the opossum. The thickness of the ribs diminishes toward the distal end of the principal piece, and the ribs end abruptly at the posterior margin of the principal piece (52,336). In human spermatozoa, the ribs are 10 to 20 nm apart and about 50 nm thick (303). A study using electron microscopy and radioautography indicated that formation of the fibrous sheath takes place throughout much of spermiogenesis in the rat and proceeds

in a distal to proximal direction (477). The longitudinal columns appeared in the distal end of the flagellum in step 2 of spermiogenesis, as thin rods joined to microtubule doublets 3 and 8. The columns increased in length during steps 2 to 10; in step 11, evenly spaced and circumferentially arranged pairs of spines formed against the plasma mem¬ brane in the distal part of the flagellum. During steps 12 to 14, these structures associated with the columns in a distal to proximal direction. In early step 15, additional material was deposited between the pairs of spines to form the de¬ finitive ribs, and the longitudinal columns became thick¬ ened. During steps 15 to 17, additional ribs were formed from posterior to anterior, along the remaining proximal segment of the principal piece (477). Radioautography fol¬ lowing injection of labeled amino acids indicated that the fibrous sheath proteins were synthesized during the entire 15-day period of fibrous sheath formation (477). These stud¬ ies supported earlier suggestions that filamentous material present in the cytoplasm of step-11 to -15 spermatids was rib precursor in human and marmoset sperm (457,478,479). They confirmed and extended the general understanding of fibrous sheath formation that had come from earlier studies on bandicoot sperm (458). A monoclonal antibody that reacts with the mouse sperm fibrous sheath (480) was used in an immunohistochemical study on fibrous sheath formation (481). The antigen was first detected in the cytoplasm of stage-14 spermatids, and staining intensity increased at stage 15. The positive reaction was observed first in the proximal portion of the principal piece and was present throughout the fibrous sheath by step 16 (481). This proximal-to-distal appearance of the antigen was opposite to that reported previously in the rat (477). Early studies indicated that the fibrous sheath was highly resistant to acid solubilization (482) and that the proteins of the fibrous sheath are stabilized by disulfide bonds (337). More recent studies took advantage of these characteristics to devise a differential solubilization procedure for isolating the fibrous sheath of rat spermatozoa (344,483). Isolated fibrous sheaths were analyzed by polyacrylamide gel elec¬ trophoresis under disulfide-bond-reducing conditions and were found to consist of a major protein of 80,000-daltons and minor proteins of 90,000-, 25,000-, and 11,000-daltons (483). In another study, the major insoluble tail component syn¬ thesized during spermiogenesis in the mouse was a 74,000dalton protein (484), which may be a fibrous sheath com¬ ponent. Monoclonal antibodies to the fibrous sheath which have been reported recently should be useful for further defining its composition (481,485,486). The antigens rec¬ ognized by two of the antibodies have not yet been iden¬ tified, but one antibody appears to recognize a 67,000-dalton protein in the mouse, a 65,000-dalton protein in the rat, and a 66,000-dalton protein in the hamster (486). This antibody also reacted with late spermatids in all three species, but did not react with spermatids or the fibrous sheath in rabbit and guinea pig.

The Spermatozoon

It has been suggested that a function of the fibrous sheath might be to modulate the plane of the flagellar beat. The attachment of doublets 3 and 8 to the longitudinal columns might restrict their participation in microtubule sliding and axoneme bending during flagellar motion. The longitudinal columns themselves might also limit bending of the flagel¬ lum in this same plane, whereas flagellar bending perpen¬ dicular to this plane would not be restricted by these features (52). Although sperm appear to swim with a planar effective stroke and recovery stroke somewhat like cilia, they also rotate (420), and the propagated waves have a three-di¬ mensional component in the distal portion of the tail (52). However, it is not known how the fibrous sheath might influence flagellar movement in the female reproductive tract. Abnormal Flagella Spermatozoa with abnormal flagella are present in low numbers in most, or perhaps all, mammals but are relatively common in humans. One study indicated that 12.5 ± 5.5% of sperm from 20 fertile men had abnormal flagella and that an even higher number of sperm (24.4 ± 7.7%) had ab¬ normal axonemes; 41.6 ± 15.3% of sperm were motile (487). Infertile men have been reported to have 18% (488) or over 30% (489) sperm with abnormal tails. It is generally assumed that abnormal sperm have little chance of achieving fertilization. However, completely immotile human sper¬ matozoa have been reported to undergo capacitation, the acrosome reaction, and fusion with the vitelline membrane of zona-free hamster oocytes in vitro (490). Also, a sperm tail lacking four doublets and the outer dense fibers has been observed in the cytoplasm of a fertilized mouse egg (491). Anecdotal reports of infertile men with sperm that are immotile or have abnormal flagella are not uncommon (492-495). Although the causes of these abnormalities are seldom known, a hereditary condition has been identified, referred to as immotile-cila syndrome or Kartagener’s syn¬ drome, which results in male sterility, situs inversus in 50% of cases, and chronic respiratory problems (496). It appears that these problems are directly or indirectly a consequence of an autosomal recessive trait for inability of cilia and flagella to move or to perform normal and coordinated movements (497). The sperm from 12 males who have the syndrome had defective axonemes with a partial or complete lack of dynein arms, a lack of outer dynein arms but a presence of the inner arms, a disorganized axoneme lacking inner arms, or abnormal short spokes and a lack of the central sheath (498). Because of the different types of de¬ fects seen in sperm and cilia, the failure to locate a single gene by linkage analysis, and the relatively high prevalence of the condition, it has been suggested that many genes, when mutated, may produce the syndrome (496). Several mutations that affect formation of the flagellum have been identified in mice. All of the mutations described so far are pleiotropic, affecting processes in addition to

/

53

spermatogenesis, and most affect formation of the sperm head as well as the flagellum (353,354). Homozygous mice with the Wobbler mutation apparently produced normal numbers of sperm, but few sperm were motile (499). The sperm appeared to have tails of normal length, and they formed heads of normal appearance, but 70% of sperm in the ductus deferens had ultrastructural defects in the fla¬ gellum. These defects included the absence of from one to four outer doublets and their corresponding dense fibers, most commonly those from position 4 through 7. Other defects seen less frequently were supernumerary tubules or absence of central pair tubules (499). Defective sperm were common in T/t mice, with the most frequent flagellar defects seen in sterile mice being a lack of four outer dense fibers and doublets (500). The male-sterile mutation, hydroce¬ phalic-polydactyl (hpy), resulted in an absence of flagella or in partially assembled axonemal structures and/or poorly organized aggregates of other tail components (501). The axoneme was usually absent or abnormally formed; the outer dense fibers and fibrous sheath were morphologically atyp¬ ical, when present. It was suggested that this axoneme dys¬ genesis might be due to failure to form stabile structures because of defective subunits (501). The sperm from four infertile men with flagellar dyski¬ nesia had normal axonemes, but abnormal periaxonemal structures. These included abnormal extension of individual dense fibers along the axoneme, altered order of termination of these structures, and a modified number and location of longitudinal columns of the fibrous sheath (502). The outer dense fibers were found to have abnormal positions with respect to each other, being placed symmetrically with re¬ spect to a plane that passes through microtubular doublet 1 and between doublets 5 and 6. The most common defect of the longitudinal columns of the fibrous sheath was the pres¬ ence of only one column adjacent to doublet 3 or 8 (502). It was suggested that these abnormalities might be due to a defect of the components of the wall of A microtubules of the outer nine doublets, at the site of formation or assembly of the outer dense fibers and longitudinal columns (503). A defect common to abnormal or degenerating flagella is the absence of microtubule doublets and dense fibers 4 through 7 (499,500,504). The same alteration has been produced by adding ATP to detergent-extracted rat sperm tails; doublets and fibers 8 through 3 appear to remain firmly attached to the fibrous sheath, while doublets 4 through 7 are extruded from the sperm tail (505). This suggests that some sperm seen with this defect may be undergoing degeneration. It has also been noted in some studies that the flagellar axo¬ neme may appear to form normally and then become pro¬ gressively disorganized. This appears to occur during spermiogenesis in quaking (506) and in hpy/hpy mice (501). However, in Wobbler mice, the percentage of abnormal sperm increases along the male tract. Only about 5% of sperm tails were abnormal in the testis, while nearly 70% of sperm from the ductus deferens were abnormal (499).

54

/

Chapter 2

Flagellar Motion The flagellar wave is reported to propagate in a plane perpendicular to the central pair of microtubules of the axoneme and to pass through doublet 1 and between doublets 5 and 6, with the active stroke being toward doublet 1 (507). The flagellum appears to twist during bend propagation (508,509), with the plane of bending moving toward doublet 2 (508). It has been suggested that the part of the axoneme containing doublets 1 through 5 is active during this phase of flagellar motion (508,510). The outer dense fibers and fibrous sheath appear to play a passive role in flagellar motion (511-514). It has been suggested that the tapering of the dense fibers results in decreasing local resistance to bending (454,502) and might explain the progressive in¬ crease in the amplitude of curvature observed during fla¬ gellar wave propagation (513,515,516). The small flagellar amplitude observed in certain abnormal human sperm (517) may be due to the constraint imposed by the abnormal ar¬ rangement of dense fibers in those sperm (502). Sperm acquire the capacity for progressive motility during epididymal maturation. Sperm from the cauda epididymis are immotile in epididymal fluid, and it is not until they are diluted into physiological medium that the capacity for mo¬ tility is expressed. It is likely that in most species, immotility is enforced by the viscoelastic drag produced by a highmolecular-weight glycoprotein (immobilin) present in epi¬ didymal fluid (518,519). However, rabbit sperm show mo¬ tility in their native fluid (520), and epididymal fluid in the bull is not particularly viscous (521). Sperm from the caput epididymis in most mammals have a vibratory or slow and ineffective beat that often results in circular swimming pat¬ terns (183). In contrast, sperm from the cauda epididymis usually move with a vigorous motion that results in rapid forward movement. In correlation with this change in swim¬ ming pattern, the flagellum appears to become more rigid and to beat with a reduced arc of curvature (183). This may be due to the increased disulfide bond formation that occurs in outer dense fibers and the fibrous sheath during epidi¬ dymal maturation (337). However, cauda sperm from dif¬ ferent species show different patterns of motility, flagellar beat, and flagellar rigidity (420). Sperm undergo additional changes in the pattern of motility in the female reproductive tract. They become hyperactivated as capacitation proceeds, moving with a vigorous whip-like beating of the flagellum at the time of fertilization (7,184). A variety of factors appear to be involved in the initiation and regulation of sperm motility (Table 8), but a unifying hypothesis for how these factors interact to accomplish this is lacking. It is generally accepted that cyclic adenosine monophosphate (cAMP) mediates sperm motility, because cAMP levels increase in sperm during epididymal transit (522,523). Also, a number of cAMP phosphodiesterase in¬ hibitors that elevate levels of cAMP in sperm induce a poorly coordinated pattern of motility in caput sperm (524). This pattern is converted to the effective pattern comparable to

TABLE 8. Factors involved in initiation and regulation of sperm motility Factor ATP required for sperm motility ATP interacts with dynein ATPase to promote sliding of outer doublets of axoneme Calcium-dependent and calmodulin-regulated adenyl cyclase converts ATP to cAMP cAMP levels in sperm increase during epididymal transit Phosphodiesterse inhibitors elevate cAMP levels and induce partial motility in caput sperm Increase in internal pH and cAMP induces motility cAMP-dependent protein kinases present cAMP stimulates phosphorylation of sperm proteins Phosphoprotein phosphatase present cAMP phosphodiesterase present Inhibitor of cAMP-dependent protein kinase present Low-molecular-weight factor in seminal plasma stimulates sperm adenyl cyclase Adenosine elevates cAMP levels and initiates motility in caput sperm High levels of adenosine inhibit sperm adenyl cyclase activity Adenosine increases level of Sadenosylhomocysteine, which inhibits sperm motility Adenyl cyclase inhibitory guaninenucleotide-binding protein present Inhibitor of cAMP-independent protein kinase present in seminal plasma Inhibitor of calcium uptake present in seminal plasma Forward motility protein in epididymal fluid modulates pattern of sperm motility Motility inhibitor present in seminal plasma pH-dependent sperm motility quiescence factor present in epididymal fluid Immotility of epididymal sperm enforced by viscoelastic drag produced by high-molecularweight glycoprotein in epididymal fluid

References 530-532 529

534-537

522, 523 524

527, 528 536-538 533, 542, 554-556 539 540 542 544

545

546-548 562-564

551

558, 559

560, 561 525, 526

557 521, 565

578, 579

that seen in caudal sperm by addition of a “forward motility protein” present in epididymal fluid (525,526). Another change that occurs as sperm transit the epididymis is an increase in internal pH; bicarbonate ions, in the presence of calcium, markedly elevate cAMP levels in cauda sperm

The Spermatozoon

(527), and conditions that elevate both pH and cAMP in caput sperm cause them to swim like cauda sperm (528). ATP is required for motility, and one of its roles is to interact with ATPases associated with the dynein arms to promote sliding of the outer doublets (529). Sperm that have been permeabilized by detergent treatment are immotile but can be reactivated with ATP, and addition of cAMP causes an increase in motility (530-532). The major role of cAMP in sperm is probably to mediate cAMP-stimulated phosphorylation of proteins essential for initiation or maintenance of motility (533). ATP is converted to cAMP in sperm by a calcium-dependent adenylate cyclase that may be controlled partly by calmodulin (534,535). The 55,000- and 49,000-dalton RI and RII regulatory subunits as well as the 40,000-dalton catalytic (C) subunits of cAMPdependent protein kinase are present in sperm, and most of the cAMP-binding activity is in the tail (536,537). Some investigators have suggested that protein kinase may exist on the surface of the sperm and that cAMP may be a first message (537), but others report that the enzyme is internal (536,538). Other enzymes needed for protein phosphory¬ lation and dephosphorylation are also present in sperm, in¬ cluding phosphoprotein phosphatase (539) and cAMP phos¬ phodiesterase (540,541). In addition, sperm have been reported to contain an inhibitor of cAMP-dependent protein kinase (542), and a 32,000-dalton calmodulin-binding pro¬ tein that inhibits phosphodiesterase has been identified in the testis (543). Seminal plasma is rich in prostaglandins and steroid hor¬ mones known to alter cAMP levels in other tissues, but until recently no external regulators had been shown to affect sperm cAMP levels, adenylate cyclase activity, or motility (533). However, it has been reported that a low-molecularweight factor in porcine seminal plasma stimulates sperm adenylate cyclase activity (544). In addition, adenosine and its analogs have been shown to elevate cAMP levels and to initiate motility in immature caput sperm in a pH-dependent manner (545). Some of the analogs used do not appear to enter cells, suggesting that adenosine may act on sperm through external adenosine receptors. There have also been reports that high levels of adenosine inhibit sperm adenylate cyclase activity, possibly through an inhibitory “P” site as¬ sociated with the adenylate cyclase catalytic subunit (546-548). Earlier studies indicated that sperm adenylate cyclase is unaffected by fluoride, guanine nucleotides, forskolin, or cholera toxin plus NAD (533,549), suggesting that the enzyme is not regulated by guanidine-nucleotide¬ binding regulatory proteins. Although stimulatory (Gs) and inhibitory (Gj) guanine-nucleotide-binding regulatory pro¬ teins for adenylate cyclase were not detected in sperm in one recent study (550), another study identified the 41,000dalton ai subunit and the 35,000-dalton (3-subunit of Gj in detergent extracts of mouse, bovine, and human sperm (551). However, Gs was not found and adenylate cyclase did not reconstitute with exogenous Gs regulatory protein (551), suggesting that adenylate cyclase may have unique regu¬

/

55

latory properties in sperm (550). Receptor-mediated regu¬ lation of guanylate cyclase activity by a peptide released from eggs has been reported in invertebrate sperm (552), but a comparable system has not been identified in vertebrate sperm. Adenylate cyclase activation in most cells results in phos¬ phorylation of specific proteins by cAMP-dependent protein kinases (553). It has been reported that radioactive phos¬ phate was incorporated into a 55,000-dalton protein in sperm from the cauda but not into those from the caput epididymis in the rat (554). In addition, a 55,000-dalton protein in bull sperm was more heavily phosphorylated in motile than in nonmotile sperm (555). Another study reported that sperm tubulin was phosphorylated in a cAMP-dependent manner and that the amount of phosphorylation was correlated with an increase in sperm motility (542). Tubulin is approxi¬ mately 55,000 daltons, but the phosphorylated protein in bull sperm did not bind colchicine (555). A more recent study reported that the stimulatory effect of cAMP on reac¬ tivation of detergent-extracted dog, human, and sea-urchin sperm required the phosphorylation of a soluble 56,000dalton protein, “axokinin” (556). Axokinin was present in extracts of immature testis, indicating that it was synthesized during spermatogenesis. Other modulators of sperm motility have been reported. A motility inhibitor in seminal plasma was highest in the fluid from the seminal vesicles, but was also present in prostatic fluid from bull, rat, and rabbit (557). This 15,000dalton modulator inhibited the reactivation of motility in detergent-extracted sperm and reduced the motility of pre¬ viously reactivated sperm and may itself be regulated by a dialyzable activator (557). An inhibitor of cAMP-independent protein kinase was also present in human seminal plasma (558,559). It was a high-molecular-weight, heat-labile, trypsin-insensitive protein. It did not appear to act via an enzymatic mechanism and was present in seminal plasma of vasectomized men, indicating that it was produced by one of the male accessory organs. Seminal plasma has also been reported to contain low-molecular-weight components that inhibit the pattern (560) or uptake of calcium by sperm (561) , and calcium is important for sperm motility (531-533). Adenosine has been shown to increase the level of S-adenosylhomocysteine, a competitive inhibitor of S-adenosylmethionine protein carboxymethylation (562). Carboxymethylation occurs in motile sperm (563,564), and agents that elevate S-adenosylhomocysteine inhibit sperm motility (562) . Finally, a pH-dependent sperm motility quiescence factor has been reported to be present in cauda epididymal fluid but has not been characterized (521,565).

SUMMARY There recently has been a considerable increase in knowl¬ edge about the sperm surface and the composition, organ¬ ization, and function of the plasma membrane. Proteins,

56

/

Chapter 2

glycoproteins, and lipids, which make up the plasma mem¬ brane, have been identified and characterized. However, the most important advance has been the realization that spermsurface components often are segregated into specific re¬ gions or domains of the plasma membrane. The regional differentiation in composition of the sperm surface results in components associated with specialized functions being located in specific domains. For example, molecules in¬ volved in the acrosome reaction are present over the anterior acrosome (85), molecules involved in fusion of the sperm and egg are present over the posterior acrosome (103), and molecules involved in flagellar activity are associated with the plasma membrane of the flagellum (551). It also has been found that the composition of the plasma membrane is modified after spermatozoa leave the testis. Changes in the sperm surface occur during maturation in the epididymis and exposure to accessory gland secretory products during ejaculation. The sperm surface is further modified in the female reproductive tract. During capacitation, some sperm-surface components are lost and others migrate laterally out of their domains. These changes in composition and organization indicate that although the do¬ mains are constant in distribution, the components of the sperm surface are dynamic features of the cell. These are exciting discoveries, but much remains to be learned about the composition, organization, and function of the sperm surface. Some proteins present in specific do¬ mains have been identified, but their amino acid sequences and native structures remain to be determined. The definition of the primary structure of functionally significant spermsurface molecules in laboratory animals may allow com¬ parable proteins and their genes to be identified in humans. Knowing the secondary and tertiary structures of these pro¬ teins will help to determine how they associate with other molecules in the cytoplasm and plasma membrane, as well as how they participate in sperm functions. In addition, the lipids present in the sperm plasma membrane are being characterized (143), but how specific lipids are distributed among different domains remains to be determined. Al¬ though it is generally accepted that the sperm surface is organized in domains, little is known about how and when domains are established, what mechanisms are responsible for targeting or segregating molecules into specific domains, and how the boundaries of domains are maintained. The morphology of spermatozoa from different species has been thoroughly described, and the main structural fea¬ tures of the head and flagellum have been well characterized. Although the time and sequence of formation of different cytoplasmic structures during spermatogenesis are known, much remains to be learned about the mechanisms of mor¬ phogenesis of spermatozoa, the regulation of these mech¬ anisms, and the composition and function of sperm-specific structures. Perhaps a good example of this is the cytoskeletal architecture of the head and flagellum (Table 6). The cytoskeleton of the sperm head includes structures lying be¬

tween the acrosome and the nucleus, between the acrosome and plasma membrane, and surrounding the nucleus pos¬ terior to the acrosome. Some of these structures are more distinguishable in*sperm containing a falciform head, whereas others are more obvious in sperm possessing a spatulate head. Several of the proteins in these structures have been partially characterized, but we do not know when they are synthesized or how they interact to form the cytoskeleton of the sperm head. It can only be hypothesized that the role of these features is to help determine or maintain the shape of the sperm head. The cytoskeleton of the flagellum includes the axoneme, outer dense fibers, fibrous sheath, and satellite fibers. The microtubules of the axoneme contain tubulin, but little is known about the composition and function of other parts of the complex and highly organized flagellum. Work is just beginning on the identi¬ fication of the proteins in this structure, but the time of synthesis, distribution, organization, and function of spe¬ cific proteins remain to be determined. Because the indi¬ vidual components of the flagellum are not well character¬ ized, there is little understanding of how they function together to produce the effective flagellar beat. Similar observations can be made about other components and activities of the spermatozoon. Some of the enzymes of the acrosome have been well characterized in vitro, but their function in vivo is not well understood. Surprisingly little is known about the nature and role of most of the other enzymes in the acrosome. The acrosome reaction to release these enzymes occurs at the end of capacitation, a process involving poorly understood metabolic changes in the sperm and in the structure and function of the plasma membrane (7). Other such changes during epididymal maturation give sperm the ability to perform coordinated flagellar motion. This is likely to involve components of the cAMP-dependent second message system in spermatozoa that are used in other cells to respond to hormonal first message stimuli (533). However, neither the first message(s) that influences fla¬ gellar motion nor the sperm receptor for the message(s) has been identified (551). These specialized features of the spermatozoon serve a common purpose of delivering new genetic material to the egg- It remains to be determined how chromatin is organized in the mammalian spermatozoon, how paternally derived genes required during development are imprinted in sperm, and whether sperm have other roles in initiation of devel¬ opment besides awakening the egg and donating a haploid genome to the next generation. It is clear that substantial questions remain about the composition, organization, and function of the spermato¬ zoon. However, many of them may be answered with new research methods now available that are leading to rapid progress in knowledge in cell and molecular biology. Mon¬ oclonal antibodies already have proven to be valuable tools for studying the structure and function of sperm-surface domains, because of their specificity, sensitivity, and use-

The Spermatozoon

fulness as morphological, biochemical, and physiological probes. It is likely that they will be equally useful for dis¬ secting the nature and role of the cytoskeleton of the sper¬ matozoon. The advances of molecular genetics are having profound effects on the understanding of the structure and function of genes and cells. Probes for genes identified in other cells are being used to determine when these genes are active in spermatogenesis (see, e.g., ref. 437) and can be used to study their regulation. As more functionally sig¬ nificant sperm-specific proteins are identified, the ap¬ proaches of molecular genetics can be used to identify and characterize the nature and function of their genes. In ad¬ dition, genetic mutations that affect spermatogenesis and sperm function may provide valuable clues for identifying genes important to spermatozoa (353). Furthermore, it is likely that there will soon be a better understanding of the regulation of sperm function. There have been substantial gains in understanding the components and activities of the second message system, and this knowledge is beginning to be applied to spermatozoa (551). Other powerful physical and chemical approaches that have not been used to study sperm before, such as use of fluorescent membrane-inter¬ calating agents and fluorescence redistribution after photobleaching, are now providing important new insights into the properties of the plasma membrane (65) and the behavior of specific surface antigens (141). Because the spermato¬ zoon is highly polarized and can be studied in vitro, it is proving to be the cell type of choice for such investigations. The use of these and other new approaches should provide a substantially better understanding of the composition, or¬ ganization, and function of the spermatozoon in the near future.

REFERENCES 1. Baccetti, B. (1986): Evolutionary trends in sperm structure. Comp. Biochem. Physiol., 85A:29-36. 2. Baccetti, B., and Afzelius, B. A. (1976): The Biology of the Sperm Cell, Monographs in Developmental Biology, Vol. 10, edited by A. Wolsky. S. Karger, Basel. 3. Roosen-Runge, E. (1977): The Process of Spermatogenesis in Mam¬ mals, Developmental and Cell Biology Series, Vol. 10, edited by M. Abercrombie, D. R. Newth, and J. G. Torrey. Cambridge University Press, Cambridge. 4. Segal, S. (1985): Sexual differentiation in vertebrates. In: The Origin and Evolution of Sex, MBL Lectures in Biology, Vol. 7, edited by H. O. Halvorson and A. Monroy, pp. 263-270. Alan R. Liss, New York. 5. Anderegg, C., and Markert, C. L. (1986): Successful rescue of microsurgically produced homozygous uniparental mouse embryos via production of aggregation chimeras. Proc. Natl. Acad. Sci. USA, 83:6509-6513. 6. de Kretser, D. M., and Kerr, J. B. (1987): Cytology of spermato¬ genesis. Chapter 20, this volume. 7. Yanagimachi, R. (1987): Fertilization. Chapter 5, this volume. 8. Pedersen, R. A. (1987): Early embryogenesis. Chapter 6, this vol¬ ume. 9. Holt, W. V. (1984): Membrane heterogeneity in the mammalian spermatozoon. In: International Review of Cytology, Vol. 87, edited by G. H. Bourne and J. F. Danielli, pp. 159-194. Academic Press, New York.

/

57

10. Friend, D. S. (1984): Membrane organization and differentiation in the guinea-pig spermatozoon. In: Ultrastructure of Reproduction, edited by J. Van Blerkom and P. M. Motta, pp. 75-85. Martinus Nijhoff, The Hague. 11. Bangham, A. D. (1961): Electrophoretic characteristics of ram and rabbit spermatozoa. Proc. R. Soc. Lond. {Biol.), 155:292-305. 12. Nevo, A. C., Michaeli, I., and Schindler, H. (1961): Electrophoretic properties of bull and of rabbit spermatozoa. Exp. Cell Res., 23:69-83. 13. Bedford, J. M. (1963): Changes in the electrophoretic properties of rabbit spermatozoa during passage through the epididymis. Nature, 200:1178-1180. 14. Cooper, G. W., and Bedford, J. M. (1971): Acquisition of surface charge by the plasma membrane of mammalian spermatozoa during epididymal maturation. Anat. Rec., 169:300-301. 15. Yanagimachi, R., Noda, Y. D., Fujimoto, M., and Nicholson, G. (1972): The distribution of negative surface charges on mammalian spermatozoa. Am. J. Anat., 135:497-520. 16. Sharon, N., and Lis. H. (1974): Use of lectins for the study of membranes. In: Methods in Membrane Biology, Vol. 3, edited by E. D. Korn, pp. 147-199. Academic Press, New York. 17. Nicolson, G. (1974): The interactions of lectins with animal cell surfaces. In: International Review of Cytology, Vol. 39, edited by G. H. Bourne and J. F. Danielli, pp. 89-190. Academic Press, New York. 18. Kashiwahara, T., Tanaka, R., and Matsomoto, T. (1965): Tail to tail agglomeration of bull spermatozoa by phytoagglutinins present in soy beans. Nature, 207:831-832. 19. Nicolson, G., and Yanagimachi, R. (1972): Terminal saccharides on sperm plasma membranes. Identification by specific agglutinins. Sci¬ ence, 177:276-279. 20. Nicolson, G., Poste, G., and Ji, T. H. (1977): The dynamics of cell membrane organization. In: Dynamic Aspects of Cell Surface Or¬ ganization, edited by G. Poste and G. Nicolson, pp. 1-73. NorthHolland, Amsterdam. 21. Koehler, J. K. (1981): Lectins as probes of the spermatozoon surface. Arch. Androl., 6:197-217. 22. Nicolson, G., Lacorbiere, M., and Yanagimachi, R. (1972): Quan¬ titative determination of plant agglutinin membrane sites on mam¬ malian spermatozoa. Proc. Soc. Exp. Biol. Med., 141:661-663. 23. Edelman, G. M., and Millette, C. F. (1971): Molecular probes of spermatozoon structures. Proc. Natl. Acad. Sci. USA, 68:2436-2440. 24. Millette, C. F. (1977): Distribution and mobility of lectin binding sites on mammalian spermatozoa. In: Immunobiology of Gametes, edited by M. Edidin and M. H. Johnson, pp. 51-71. Cambridge University Press, Cambridge. 25. Schwarz, M. A., and Koehler, J. K. (1979): Alterations in lectin binding to guinea pig spermatozoa accompanying in vitro capacitation and the acrosome reaction. Biol. Reprod., 21:1295-1307. 26. Koehler, J. K. (1982): The mammalian sperm surface: An overview of structure with particular reference to mouse spermatozoa. In: Pros¬ pects for Sexing Mammalian Sperm, edited by R. P. Amann and G. E. Seidel, Jr., pp. 23-42. Colorado Associated University Press, Boulder. 27. Fenderson, B. A., O’Brien, D. A., Millette, C. F., and Eddy, E. M. (1984): Stage-specific expression of three cell surface carbo¬ hydrate antigens during murine spermatogenesis detected with monoclonal antibodies. Dev. Biol., 103:117-128. 28. Gordon, M., Dandekar, P. V., and Bartoszewicz, W. (1975): The surface coat of epididymal, ejaculated and capacitated sperm. J. Ultrastruct. Res., 50:199-207. 29. Nicolson, G., and Yanagimachi, R. (1974): Mobility and restriction of mobility of plasma membrane lectin-binding components. Science 184:1294-1296. 30. Kinsey, W. H., and Koehler, J. K. (1976): Fine structural locali¬ zations of concanavalin A binding sites on hamster spermatozoa. J. Supramol. Struct., 5:185-189. 31. Friend, D. S., and Fawcett, D. W. (1974): Membrane differentiations in freeze-fractured mammalian sperm. J. Cell Biol., 63:641-664. 32. Suzuki, F., and Nagano, T. (1980): Epididymal maturation of rat spermatozoa studied by thin sectioning and freeze-fracture. Biol. Reprod., 22:1219-1231. 33. Suzuki, F. (1981): Changes in intramembranous particle distribution in epididymal spermatozoa of the boar. Anat. Rec., 199:361-376.

58

/

Chapter 2

34. Koehler, J. K. (1970): A freeze-etch study of rabbit spermatozoa with particular reference to head structures. J. Ultrastruct. Res., 33:598-614. 35. Schulte-Wrede, S., and Wetzstein, R. (1972): Raster-Elektronenmikroskopie von Spermien des Hausschaufs (Ovis ammon aries, L.), Z. Zellforsch., 134:105-127. 36. Eddy, E. M., and Koehler, J. K. (1982): Restricted domains of the sperm surface. In: Scanning Electron Microscopy, Vol. 3, edited by O. Johari, pp. 1313-1323. SEM Inc., Chicago. 37. Koehler, J. K. (1966): Fine structure observations in frozen-etched bovine spermatozoa. J. Ultrastruct. Res., 16:359-375. 38. Koehler, J. K. (1978): The mammalian sperm surface: Studies with specific labeling techniques. In: International Review of Cytology, Vol. 54, edited by G. H. Bourne and J. F. Danielli, pp. 73-108. Academic Press, New York. 39. Phillips, D. M. (1977): Surface of the equatorial segment of mam¬ malian acrosome. Biol. Reprod., 16:128-137. 40. Toshimori, K., Higashi, R., and Oura, C. (1986): Distribution of intramembranous particles and filipin-sterol complexes in mouse sperm membranes: Polyene antibiotic filipin treatment. Am. J. Anat., 174:455-470. 41. Koehler, J. K. (1973): Studies on the structure of the postnuclear sheath of water buffalo spermatozoa. J. Ultrastruct. Res., 44:355-368. 42. Flechon, J.-E. (1974): Freeze-fracturing of rabbit spermatozoa. J. Submicrosc. Cytol., 19:59-64. 43. Bradley, M. P., Ryans, D. G., and Forrester, I. T. (1980): Effects of filipin, digitonin, and polymyxin B on plasma membrane of ram spermatozoa—An EM study. Arch. Androl., 4:195-204. 44. Koehler, J. K. (1972): Human sperm head ultrastructure: A freeze¬ etching study. J. Ultrastruct. Res., 39:520-539. 45. Gresson, R. A. R., and Zlotnik, I. (1945): A comparative study of the cytoplasmic components of the male germ-cells of certain mam¬ mals. Proc. R. Soc. Edinb. B, 62:137-170. 46. Hancock, J. L., and Trevan, D. J. (1957): The acrosome and postnuclear cap of bull spermatozoa. J. R. Microsc. Soc., 76:77-83. 47. Piko, L. (1969): Gamete structure and sperm entry in mammals. In: Fertilization, Vol. 2, edited by C. B. Metz and A. Monroy, pp. 325-403. Academic Press, New York. 48. Woolley, D. M. (1970): A posterior ring in the spermatozoa of species of muridae. J. Reprod. Fertil., 23:361-363. 49. Koehler, J. K. (1983): Structural heterogeneity of the mammalian sperm flagellar membrane. J. Submicrosc. Cytol., 15:247-253. 50. Friend, D. S., and Rudolf, I. (1974): Acrosomal disruption in sperm. Freeze-fracture of altered membranes. J. Cell. Biol., 63:466-479. 51. Koehler, J. K., and Gaddum-Rosse, P. (1975): Media induced al¬ terations of the membrane associated particles of the guinea pig sperm tail. J. Ultrastruct. Res., 51:106-118. 52. Fawcett, D. W. (1975): The mammalian spermatozoon. Dev. Biol., 44:394-436. 53. Gilula, N. B., and Satir, P. (1972): The ciliary necklace: A ciliary membrane specialization. J. Cell Biol., 53:494-509. 54. Olson, G. E., Lifsics, M., Hamilton, D. W., and Fawcett, D. W. (1977): Structural specializations in the flagellar plasma membrane of opossum spermatozoa. J. Ultrastruct. Res., 59:207-221. 55. Friend, D. S., Elias, P. M., and Rudolf, I. (1979): Disassembly of the guinea-pig sperm tail. In: The Spermatozoon, edited by D. W. Fawcett and J. M. Bedford, pp. 157-168. Urban and Schwarzenberg, Baltimore. 56. Friend, D. S., and Heuser, J. E. (1981): Orderly particle arrays on the mitochondrial outer membrane in rapidly-frozen sperm. Anat. Rec., 159:198-199. 57. Elias, P. M., Gorke, J., and Friend, D. S. (1978): Freeze-fracture identification of sterol-digitonin complexes in cell and liposome membrane. J. Cell Biol., 78:577-596. 58. Enders, G. C., Werb, Z., and Friend, D. S. (1983): Lectin binding to guinea-pig sperm zipper particles. J. Cell Sci., 60:303-329. 59. Elias, P. M., Friend, D. S., and Goerke, J. (1979): Membrane sterol heterogeneity. Freeze-fracture detection with saponins and filipin. J. Histochem. Cytochem., 27:1247-1260. 60. Friend, D. S. (1982): Plasma-membrane diversity in a highly polar¬ ized cell. J. Cell Biol., 93:243-249. 61. Friend, D. S. (1980): Freeze-fracture alterations in guinea-pig sperm membrane preceding gamete fusion. In: Membrane-Membrane In¬

62.

63.

64.

65.

66. 67.

68.

69.

70.

71. 72.

73.

74.

75.

76.

77. 78.

79.

80.

81.

82.

83.

84.

teractions, edited by N. B. Gilula, pp. 153—165. Raven Press, New York. Bearer, E. L., and Friend, D. S. (1980): Anionic lipid domains: Correlation and functional topography in a mammalian cell mem¬ brane. Proc. Nhtl. Acad. Sci. USA, 77:6601-6605. Friend, D. S., and Bearer, E. L. (1981): p-Hydroxysteroi distribution as determined by freeze-fracture cytochemistry. Histochem. J., 13:535-546. Bearer, E. L., and Friend, D. S. (1982): Modifications of anionic lipid domains preceding membrane fusion in guinea pig sperm. J. Cell Biol., 92:604-615. Wolf, D. E., and Voglmayr, J. K. (1984): Diffusion and regional¬ ization in membranes of maturing ram spermatozoa. J. Cell Biol., 98:1678-1684. Stemberger, L. A. (1979): Immunocytochemistry, John Wiley & Sons, New York. Koo, G. C., Stackpole, C. W., Boyse, E. A., Hammerling, U., and Lardis, M. P. (1973): Topographical location of H-Y antigen on mouse spermatozoa by immunoelectronmicroscopy. Proc. Natl. Acad. Sci. USA, 70:1502-1505. Fellous, M., Gachelin, G., Buc-Caron, M.-H., Dubois, P., and Jacob, F. (1974): Similar location of an early embryonic antigen on mouse and human spermatozoa. Dev. Biol., 41:331-337. Lopez, L. C., Bayna, E. M., Litoff, D., Shaper, N. L., Shaper, J. H., and Shur, B. D. (1985): Receptor function of mouse sperm surface galactosyltransferase during fertilization. J. Cell Biol., 101:1501-1510. O’Rand, M. G., and Romrell, L. J. (1980): Appearance of regional surface autoantigens during spermatogenesis: Comparison of anti¬ testis and anti-sperm antisera. Dev. Biol., 75:431-441. Koehler, J. K. (1974): Studies on the distribution of antigenic sites on the surface of rabbit spermatozoa. J. Cell Biol., 67:647-659. Millette, C. F., and Bellve, A. R. (1977): Temporal expression of membrane antigens during mouse spermatogenesis. J. Cell Biol., 74:86-97. Koehler, J. K., and Perkins, W. D. (1974): Fine structure obser¬ vations on the distribution of antigenic sites on guinea pig sperma¬ tozoa. J. Cell Biol., 60:789-795. Tung, K. S. K., Han, L.-B. P., and Evan, A. P. (1979): Differ¬ entiation autoantigen of testicular cells and spermatozoa in the guinea pig. Dev. Biol., 68:224-238. Tung, P. S., and Fritz, I. B. (1978): Specific surface antigens on rat pachytene spermatocytes and successive classes of germinal cells. Dev. Biol., 64:297-315. Koo, G. C., Boyse, E. A., and Wachtel, S. S. (1977): Immunogenetic techniques and approaches in the study of sperm and testicular cell surface antigens. In: Immunobiology of Gametes, edited by M. Edidin and M. H. Johnson, pp. 73-80. Cambridge University Press, Cambridge. O’Rand, M. J. (1977): The presence of sperm-specific isoantigens on the egg following fertilization. J. Exp. Zool., 202:267-273. Herr, J. C., and Eddy, E. M. (1980): Identification of mouse sperm surface antigens by a surface labeling and immunoprecipitation ap¬ proach. Biol. Reprod., 22:1263-1274. Villarroya, S., and Scholler, R. (1986): Regional heterogeneity of human spermatozoa detected with monoclonal antibodies. J. Reprod. Fertil., 76:435-447. Primakoff, P., and Myles, D. G. (1983): A map of the guinea pig sperm surface constructed with monoclonal antibodies. Dev. Biol., 98:417-428. Gaunt, S. J., Brown, C. R., and Jones, R. (1983): Identification of mobile and fixed antigens on the plasma membrane of rat spermatozoa using monoclonal antibodies. Exp. Cell Res., 144:275-284. Jones, R., Brown, C. R., von Glos, K. I., and Gaunt, S. J. (1985): Development of a maturation antigen on the plasma membrane of rat spermatozoa in the epididymis and its fate during fertilization. Exp. Cell Res., 156:31-44. Moore, H. D. M., and Hartman, T. D. (1984): Localization by monoclonal antibodies of various surface antigens of hamster sper¬ matozoa and the effect of antibody on fertilization in vitro. J. Reprod. Fertil., 70:175-183. Feuchter, F. A., Vernon, R. B., and Eddy, E. M. (1981): Analysis of the sperm surface with monoclonal antibodies: Topographically

The Spermatozoon

85.

86.

87.

88.

89.

90.

91.

92.

93.

94.

95.

96.

97.

98.

99.

100.

101.

102.

103.

104.

105.

106.

restricted antigens appearing in the epididymis. Biol. Reprod., 24:1099-1110. Saling, P. M. (1986): Mouse sperm antigens that participate in fer¬ tilization. IV. A monoclonal antibody prevents zona penetration by inhibition of the acrosome reaction. Dev. Biol., 117:511-519. Okabe, M., Katsuaki, T., Adachi, T., Kohama, T., and Mimura, T. (1986): Inconsistent reactivity of an anti-sperm monoclonal an¬ tibody and its relationship to sperm capacitation. J. Reprod. Im¬ munol., 9:67-70. Yan, C. Y., Wang, L. F., Sato, E., and Koide, S. S. (1983): Mon¬ oclonal antibody inducing human sperm agglutination. Am. J. Re¬ prod. Immunol., 4:111-115. Herr, J. C., Fowler, J. E., Howards, S. S., Sigman, M., Sutherland, W. M., and Koons, D. J. (1985): Human antisperm monoclonal antibodies constructed postvasectomy. Biol. Reprod., 32:695-712. Glassy, M. C., Surh, C. D., and Sarkar, S. (1984): Murine mon¬ oclonal antibodies that identify antigenically distinct subpopulations of human sperm. Hybridoma, 3:363-371. Vernon, R. B., Muller, C. H., and Eddy, E. M. (1987): Further characterization of a secreted epididymal glycoprotein in mice that binds to sperm tails. J. Androl. 8:123-128. Toshimori, K., and Eddy, E. M. (1987): Epididymal maturation produces a 31,000 molecular weight antigen on the surface of fla¬ gellum of the mouse spermatozoon (submitted). Schmell, E. D., Gulyas, B. J., Yuan, L. C., and August, J. T. (1982): Identification of mammalian sperm surface antigens: II. Char¬ acterization of an acrosomal cap protein and a tail protein using monoclonal anti-mouse sperm antibodies. J. Reprod. Immunol., 4:91106. Gaunt, S. J., Brown, C. R., and Jones, R. (1983): Identification of mobile and fixed antigens on the plasma membrane of rat spermatozoa using monoclonal antibodies. Exp. Cell Res., 144:275-284. Isahakia, M., and Alexander, N. J. (1984): Interspecies cross-reac¬ tivity of monoclonal antibodies directed against human sperm anti¬ gens. Biol. Reprod., 30:1015-1026. Hinrichsen-Kohane, A. C., Hinrichsen, M. J., and Schill, W.-B. (1985): Analysis of antigen expression on human spermatozoa by means of monoclonal antibodies. Fertil. Steril., 43:279-285. Crichton, D. N., and Cohen, B. B. (1983): Analysis of the murine sperm surface with monoclonal antibodies. J. Reprod. Fertil., 68:497-505. Gaunt, S. J. (1982): A 28K-dalton cell surface autoantigen of sper¬ matogenesis: Characterization using a monoclonal antibody. Dev. Biol., 89:92-100. Lee, G., C.-Y., Wong, E., and Teh, C.-Z. (1984): Analysis of mouse sperm isoantigens using specific monoclonal antibodies. Am. J. Re¬ prod. Immunol., 6:37-43. Saling, P. M., and Lakoski, K. A. (1985): Mouse sperm antigens that participate in fertilization. II. Inhibition of sperm penetration through the zona pellucida using monoclonal antibodies. Biol. Re¬ prod., 33:527-536. Myles, D. G., Primakoff, P., and Bellve, A. R. (1981): Surface domains of the guinea pig sperm defined with monoclonal antibodies. Cell, 23:433-439. Myles, D. G., and Primakoff, P. (1985): Sperm surface domains. In: Hybridoma Technology in the Biosciences and Medicine, edited by T. A. Springer, pp. 239-250. Plenum Press, New York. Naz, R. N., Saxe, J. M., and Menge, A. C. (1983): Inhibition of fertility in rabbits by monoclonal antibodies against sperm. Biol. Reprod., 28:249-254. Saling, P. M., Irons, G., and Waibel, R. (1985): Mouse sperm antigens that participate in fertilization. I. Inhibition of sperm fusion with the egg plasma membrane using monoclonal antibodies. Biol. Reprod., 33:515-526. Ellis, D. H., Hartman, T. D., and Moore, H. D. M. (1985): Mat¬ uration and function of the hamster spermatozoon probed with mon¬ oclonal antibodies. J. Reprod. Immunol., 7:299-314. Wolf, D. P., Sokoloski, J. E., Dandekar, P., and Bechtol, K. B. (1983): Characterization of human sperm surface antigens with mon¬ oclonal antibodies. Biol. Reprod., 29:713-723. Naz, R. K., Alexander, N. J., Isahakia, M., and Hamilton, M. S. (1984): Monoclonal antibody to a human germ cell membrane gly¬ coprotein that inhibits fertilization. Science, 225:342-344.

/

59

107. Bechtol, K. B., Brown, S. C., and Kennett, R. H. (1979): Recog¬ nition of differentiation antigens of spermatogenesis in the mouse by using antibodies from spleen cell-myeloma hybrids after syngeneic immunization. Proc. Natl. Acad. Sci. USA, 76:363-367. 108. Bechtol, K. B. (1984): Characterization of a cell-surface differen¬ tiation antigen of mouse spermatogenesis: Timing and localization of expression by immunohistochemistry using a monoclonal anti¬ body. J. Embryol. Exp. Morphol., 81:93-104. 109. Saxena, N. K., Russell, L. D., Saxena, N., and Peterson, R. N. (1986): Immunofluorescence antigen localization on boar sperm plasma membranes: Monoclonal antibodies reveal apparent new domains and apparent redistribution of surface antigens during sperm maturation and at ejaculation. Anat. Rec., 214:238-252. 110. Bedford, J. M. (1963): Morphological changes in rabbit spermatozoa during passage through the epididymis. J. Reprod. Fertil., 5:169-282. 111. Jones, R. C. (1971): Studies of the structure of the head of boar spermatozoa from the epididymis. J. Reprod. Fertil. (Suppl.), 13:51-64. 112. Fawcett, D. W., and Hollenberg, R. D. (1963): Changes in the acrosomes of guinea pig spermatozoa during passage through the epididymis. J. Reprod. Fertil. (Suppl.), 6:276-292. 113. Eddy, E. M., Vemon, R. B., Muller, C. H., Hahnel, A. C., and Fenderson, B. A. (1985): Immunodissection of sperm surface modifications during epididymal maturation. Am. J. Anat., 174:225237. 114. Burridge, K., and Feramisco, J. R. (1982): a-Actinin and vinculin from nonmuscle cells: Calcium-sensitive interactions with actin. In: Cold Spring Harbor Symposia on Quantitative Biology, Vol. XLVI, Part 2, pp. 587-597. Cold Spring Harbor, New York. 115. Welch, J. E., and O’Rand, M. G. (1985): Identification and distri¬ bution of actin in spermatogenic cells and spermatozoa of the rabbit. Dev. Biol., 109:411-417. 116. Campanella, C., Gabbiani, G., Baccetti, B., Burrini, A. G., and Pallini, V. (1979): Actin and myosin in the vertebrate acrosomal region. J. Submicrosc. Cytol., 11:53-71. 117. Clarke, G. N., Clarke, F. M., and Wilson, S. (1982): Actin in human spermatozoa. Biol. Reprod., 26:319-327. 118. Talbot, P., and Kleve, M. G. (1978): Hamster sperm cross-react with antiactin. J. Exp. Zool., 204:131-136. 119. Peterson, R. N., Russell, L. D., Bundman, D., and Freund, M. (1978): Presence of microfilaments and tubular structure in chemi¬ cally induced acrosome reactions of boar spermatozoa. Biol. Reprod., 19:459-465. 120. Tamblyn, T. M. (1980): Identification of actin in boar epididymal spermatozoa. Biol. Reprod., 22:727-734. 121. Koehler, J. K. (1978): Observations on the fine structure of vole spermatozoa with particular reference to cytoskeletal elements in the mature sperm head. Gamete Res., 1:247-257. 122. Olson, G. E., and Winfrey, V. P. (1985): Substructure of a cyto¬ skeletal complex associated with the hamster sperm acrosome. J. Ultrastruct. Res., 92:167-179. 123. Goodman, S. R., and Schiffer, K. (1983): The spectrin membrane skeleton of normal and abnormal human erythrocytes: A review. Am. J. Physiol., 244:C121-C141. 124. Lazerides, E., and Moon, R. T. (1984): Assembly and topogenesis of the spectrin-based membrane skeleton in erythroid development. Cell, 37:354-356. 125. Damjanov, I., Damjanov, A., Lehto, V.-P., and Virtanen, I. (1986): Spectrin in mouse gametogenesis and embryogenesis. Dev. Biol., 114:132-140. 126. Repasky, E. A., Granger, B. L., and Lazarides, E. (1982): Wide¬ spread occurrence of avian spectrin in nonerythroid cells. Cell, 29:821-833. 127. Virtanen, I., Badley, R. A., Paasivuo, R., and Lehto, V.-P. (1984): Distinct cytoskeletal domains revealed in sperm cells. J. Cell Biol., 99:1083-1091. 128. McIntosh, J. R., and Porter, K. R. (1967): Microtubules in the spermatids of the domestic fowl. J. Cell Biol., 35:153-173. 129. Fawcett, D. W., Anderson, W. A., and Phillips, D. M. (1971): Morphogenetic factors influencing the shape of the sperm head. Dev. Biol., 26:220-251. 130. Blom, E., and Birch-Anderson, A. (1965): The ultrastructure of the bull sperm. Nord, Vet. Med., 17:193-212.

60

/

Chapter 2

131. Plattner, H. (1971): Bull spermatozoa: A re-investigation by freeze etching using widely different cryofixation procedures. J. Submicrosc. Cytol., 3:19-32. 132. Fawcett, D. W., and Ito, S. (1965): The fine structure of bat sper¬ matozoa. Am. J. Anat., 116:567-610. 133. Fawcett, D. W., Eddy, E. M., and Phillips, D. M. (1970): Obser¬ vations on the fine structure and relationships of the chromatoid body in mammalian spermatogenesis. Biol. Reprod., 2:129-153. 134. Russell, L. D. (1980): Sertoli-germ cell interactions: A review. Ga¬ mete Res., 3:179-202. 135. Russell, L. D. (1984): Spermiation—The sperm release process: Ultrastructural observations and unresolved problems. In: Ultrastruc¬ ture of Reproduction, edited by J. Van Blerkom and P. M. Motta, pp. 46-66. Martinus Nijhoff, The Hague. 136. Ross, M. H., and Dobler, J. (1975): The Sertoli cell junctional specializations and their relationship to the germinal epithelium as observed after efferent ductule ligation. Anat. Rec., 183:267-292. 137. Russell, L. D., and Clermont, Y. (1976): Anchoring device between Sertoli cells and late spermatids in rat seminiferous tubules. Anat. Rec., 185:259-278. 138. Russell, L. D. (1979): Spennatid-Sertoli tubulobulbar complexes as devices for elimination of cytoplasm from the head region of late spermatids of the rat. Anat. Rec., 194:233-246. 139. Burgos, M. H., Blaquier, J., Cameo, M. S., and Gutierrez, L. (1972): Morphological maturation of spermatozoa in the epididymis. In: Biology of Reproduction, Symposium III, Pan American Congress of Anatomy, edited by J. T. Vilardo and B. A. Kasprow, pp. 367371. Pan American Association of Anatomy, New Orleans. 140. Fawcett, D. W. (1965): The anatomy of the mammalian spermato¬ zoon with particular reference to the guinea pig. Z. Zellforsch., 67:279-296. 141. Myles, D. G., Primakoff, P., and Koppel, D. E. (1984): A localized surface protein of guinea pig sperm exhibits free diffusion in its domain. J. Cell Biol., 98:1905-1909. 142. Langlais, J., and Roberts, K. D. (1985): A molecular membrane model of sperm capacitation and the acrosome reaction of mammalian spermatozoa. Gamete Res., 12:183-224. 143. Nikolopoulau, M., Soucek, D. A., and Vary, J. C. (1985): Changes in the lipid content of boar sperm plasma membranes during epididymal maturation. Biochem. Biophys. Acta, 815:486-498. 144. Murry, R. K., Narasimhan, R., Levine, M., Shirley, M., Lingwood, C. A., and Schachter, H. (1980): Galactoglycerolipids of mammalian testis, spermatozoa and nervous tissues. In: Cell Surface Glycolipids, ACS Symposium Series, No. 128, edited by C. Sweeley, pp. 105— 125. American Chemical Society Press, Washington D.C. 145. Komblatt, M. J., Knapp, A., Levine, M., Schachter, H., and Mur¬ ray, R. K. (1974): Studies on the structure and formation during spermatogenesis of the sulfoglycerogalactolipid of rat testis. Can. J. Biochem., 52:689-697. 146. Lingwood, C. A. (1985): Protein-glycolipid interactions during sper¬ matogenesis. Binding of specific germ cell proteins to sulfatoxygalactosylacylalkylglycerol, the major glycolipid of mammalian male germ cells. Can. J. Biochem. Cell Biol., 63:1077-1085. 147. Lingwood, C. A., and Schachter, H. (1981): Localization of sulfatoxygalactosylacylalkylglycerol at the surface of rat testicular ger¬ minal cells by immunocytochemical techniques: pH dependence of a nonimmunological reaction between immunoglobulin and germinal cells. J. Cell Biol., 89:621-630. 148. Eddy, E. M., Muller, C. H., and Lingwood, C. A. (1985): Prepa¬ ration of monoclonal antibody to sulfatoxygalactosylglycerolipid by in vitro immunization with a glycolipid-glass conjugate. J. Immunol. Methods, 81:137-146. 149. Langlais, J., Zollinger, M., Plante, L., Chapdelaine, A., Bleau, G., and Roberts, K. D. (1981): Localization of cholesteryl sulfate in human spermatozoa in support of a hypothesis for the mechanism of capacitation. Proc. Natl. Acad. Sci. USA, 78:7266-7270. 150. Mann, T., and Lutwak-Mann, C. (1981): Male Reproductive Func¬ tion and Semen, Springer-Verlag, New York. 151. Acott, T. S., Katz, D. F., and Hoskins, D. D. (1983): Movement characteristics of bovine epididymal spermatozoa: Effects of forward motility protein and epididymal maturation. Biol. Reprod., 29:389-399. 152. Cooper, T. G., Waites, G. M. H., and Nieschlag, E. (1986): The epididymis and male fertility. A symposium report. Int. J. Androl 9:81-90.

153. Saling, P. M. (1982): Development of the ability to bind zonae pellucidae during epididymal maturation: Reversible immobilization of mouse spermatozoa by lanthanum. Biol. Reprod., 26:429-436. 154. Orgebin-Crist, M.-C., and Foumier-Delpech, S. (1982): Sperm-egg interaction. Evidence for maturational changes during epididymal transit. J. Androl., 3:429-433. 155. Moore, H. D. M. (1979): The net negative surface charge of mam¬ malian spermatozoa as determined by isoelectric focusing. Changes following sperm maturation, ejaculation, incubation in the female tract, and after enzyme treatment. Int. J. Androl., 2:244-262. 156. Bedford, J. M., Calvin, H. I., and Cooper, G. W. (1973): The maturation of spermatozoa in the human epididymis. J. Reprod. Fertil. (Suppl.), 18:199-213. 157. Nicolson, G. L., Usui, N, Yanagimachi, R., Yanagimachi, H., and Smith, J. R. (1977): Lectin-binding sites on the plasma membranes of rabbit spermatozoa. Changes in surface receptors during epi¬ didymal maturation and after ejaculation. J. Cell Biol., 74:950962. 158. Holt, W. V. (1980): Surface-bound sialic acid on ram and bull sper¬ matozoa: Deposition during epididymal transit and stability during washing. Biol. Reprod., 23:847-857. 159. Flechon, J. E. (1975): Ultrastructural and cytochemical modifica¬ tions of rabbit spermatozoa during epididymal transport. In: The - Biology of Spermatozoa. Transport, Survival and Fertilizing Ability, edited by E. S. E. Hafez and C. G. Thibault, pp. 36-45. Karger, Basel. 160. Bedford, J. M., and Millar, R. P. (1978): The character of sperm maturation in the epididymis of the ascrotal hyrax, Procavia capensis, and armadillo, Dasypus novemcinctus. Biol. Reprod., 19:396-406. 161. Hammerstedt, R. H., Hay, S. R., and Amann, R. P. (1982): Mod¬ ification of ram sperm membranes during epididymal transit. Biol. Reprod., 27:745-754. 162. Foumier-Delpech, S., and Courot, M. (1980): Glycoproteins of ram sperm plasma membrane. Relationship of protein having affinity for Con A to epididymal maturation. Biochem. Biophys. Res. Commun , 96:756-761. 163. Lewin, L. W., Weissenberg, R., Sobel, J. S., Marcus, Z., and Nebel, L. (1979): Differences in Con-A-FITC binding to rat spermatozoa during epididymal maturation and capacitation. Arch. Androl., 2:279281. 164. Foumier-Delpech, S., Danzo, B. J., and Orgebin-Crist, M.-C. (1977): Extraction of concanavalin A affinity material from rat testicular and epididymal spermatozoa. Ann. Biol. Anim. Biochem. Biophys., 17:207213. 165. Dawson, R. M. C., and Scott, T. W. (1964): Phospholipid com¬ position of epididymal spermatozoa prepared by density gradient centrifugation. Nature, 202:292-293. 166. Quinn, P. J., and White, I. G. (1967): Phospholipid and cholesterol content ot epididymal and ejaculated ram spermatozoa and seminal plasma in relation to cold shock. Aust. J. Biol. Sci., 20:1205-1215. 167. Grogan, D. E. Mayer, D. T., and Sikes, J. D. (1966): Quantitative differences in phospholipids of ejaculated spermatozoa and sper¬ matozoa from three different levels of the epididymis of the boar. J. Reprod. Fertil., 12:431-436. 168. Poulos, A., Voglmayr, J. K.. and White, I. G. (1973): Phospholipid changes in spermatozoa during passage through the genital tract of the bull. Biochim. Biophys. Acta, 306:194-202. 169. Poulos, A., Brown-Woodman, P. D. C., White, I. G., and Cox, R. I. (1975): Changes in phospholipids of ram spermatozoa during migration through the epididymis and possible origin of prostaglan¬ dins F2a in testicular and epididymal fluid. Biochim. Biophys Acta, 388:12-21. 170. Temer, C., MacLaughlin, J., and Smith, B. R. (1975): Changes in lipase and phospholipase activities of rat spermatozoa in transit from the caput to the cauda epididymis. J. Reprod. Fertil., 45:1-8. 171. Evans, R. W., and Setchell, B. P. (1979): Lipid changes in boar spermatozoa during epididymal maturation with some observations on the flow and composition of boar rete testis fluid. J Reprod Fertil., 57:189-196. 172. Scott, T. W., Voglmayr, J. K., and Setchell, B. P. (1967): Lipid composition and metabolism in testicular and ejaculated ram sper¬ matozoa. Biochem. J., 102:456-461. 173. Bleau, G., and VandenHeuvel, W. J. A. (1974): Desmosteryl sulfate and desmosterol in hamster epididymis. Steroids, 24:549-556.

The Spermatozoon 174. Legault, Y., Bouthillier, M., Bleau, G., Chapdelaine, A., and Rob¬ erts, K. D. (1979): The sterol and sterol sulfate content of the male hamster reproductive tract. Biol. Reprod., 20:1213-1219. 175. Lalumiere, G., Bleau, G., Chapdelaine, A., and Roberts, K. D. (1976): Cholesterol sulfate and sterol sulphatase in the human re¬ productive tract. Steroids, 27:247-260. 176. Parks, J. E., and Hammerstedt, R. H. (1985): Developmental changes occurring in the lipids of ram epididymal spermatozoa plasma mem¬ brane. Biol. Reprod., 32:653-668. 177. Hammerstedt, R. H., Keith, A. D., Hay, S., Deluca, N., and Amann, R. P. (1979): Changes in ram sperm membranes during epididymal transit. Arch. Biochem. Biophys., 196:7-12. 178. Voglmayr, J. K., Scott, T. W., Setchell, B. P., and Waites, G. M. H. (1967): Metabolism of testicular spermatozoa and cha¬ racteristics of testicular fluid collected from conscious rams. J. Re¬ prod. Fertil., 14:87-99. 179. Scott, T. W., Voglmayr, J. K., and Setchell, B. P. (1967): Lipid composition and metabolism in testicular and ejaculated ram sper¬ matozoa. Biochem. J., 102:456—461. 180. Vijayasarathy, S., and Balaram, P. (1982): Regional differentiation in bull sperm plasma membranes. Biochem. Biophys. Res. Commun., 108:760-764. 181. Orgebin-Crist, M.-C., Danzo, B. J., and Davies, J. (1975): Endo¬ crine control of the development and maintenance of sperm fertilizing ability in the epididymis. In: Handbook of Physiology, Vol. 5, En¬ docrinology, Section 7, Male Reproductive System, edited by R. O. Greep, pp. 319-338. American Physiological Society, Washington D.C. 182. Hamilton, D. W. (1975): Structure and function of the epithelium lining the ductuli efferentes, ductus epididymis, and ductus deferens in the rat. In: Handbook of Physiology, Vol. 5, Endocrinology, Sec¬ tion 7, Male Reproductive System, edited by R. O. Greep, pp. 259301. American Physiological Society, Washington D.C. 183. Bedford, J. M. (1975): Maturation, transport and fate of spermatozoa in the epididymis. In: Handbook of Physiology, Vol. 5, Endocri¬ nology, Section 7, Male Reproductive System, edited by R. O. Greep, pp. 303-317. American Physiological Society, Washington D.C. 184. Yanagimachi, R. (1981): Mechanisms of fertilization in mammals. In: Fertilization and Embryonic Development in Vitro, edited by L. Mastroianni and J. D. Biggers, pp. 81-182. Plenum Press, New York. 185. Austin, C. R. (1985): Sperm maturation in the male and female genital tracts. In: Biology of Fertilization, Vol. 2, Biology of the Sperm, edited by C. B. Metz and A. Monroy, pp. 121-155. Aca¬ demic Press, Orlando, Florida. 186. Olson, G. E., and Hamilton, D. W. (1978): Characterization of the surface glycoproteins of rat spermatozoa. Biol. Reprod., 19:26-35. 187. Olson, G. E., and Danzo, B. J. (1981): Surface changes in rat spermatozoa during epididymal transit. Biol. Reprod., 24:431^443. 188. Toowicharanount, P., and Chulavatnatol, M. (1983): Characteri¬ zation of sialoglycoproteins of rat epididymal fluid and spermato¬ zoa by periodate-tritiated borohydride. J. Reprod. Fertil., 67:133141. 189. Brown, C. R. von Glos, K. I., and Jones, R. (1983): Changes in plasma membrane glycoproteins of rat spermatozoa during matura¬ tion in the epididymis. J. Cell Biol., 96:256-264. 190. Jones, R., Phorpramool, C., Setchell, B. P., and Brown, C. R. (1981): Labelling of membrane glycoproteins on rat spermatozoa collected from different regions of the epididymis. Biochem. J., 200:457-460. 191. Jones, R., von Glos, K. I., and Brown, C. R. (1981): Characteri¬ zation of hormonally regulated secretory proteins from the caput epididymidis of the rabbit. Biochem. J., 196:105-114. 192. Zaheb, R., and Orr, G. A. (1984): Characterization of a maturationassociated glycoprotein on the plasma membrane of rat caudal epi¬ didymal sperm. J. Biol. Chem., 259:839-848. 193. Faye, J. C., Duguet, L., Mazzuca, M., and Bayard, F. (1980): Purification, radioimmunoassay, and immunohistochemical locali¬ zation of a glycoprotein produced by the rat epididymis. Biol. Re¬ prod., 23:423-432. 194. Voglmayr, J. K., Fairbanks, G., Jakowitz, M. A., and Colella, J. R. (1980): Post-testicular developmental changes in the ram sperm cell surface and their relationship to luminal fluid proteins of the reproductive tract. Biol. Reprod., 22:655-667.

/

61

195. Voglmayr, J. K., Fairbanks, G., Vespa, D. B., and Colella, J. R. (1982): Studies on mechanisms of surface modifications in ram sper¬ matozoa during the final stages of differentiation. Biol. Reprod., 26:483-500. 196. Voglmayr, J. K., Fairbanks, G., and Lewis, R. G. (1983): Surface glycoprotein changes in ram spermatozoa during epididymal matu¬ ration. Biol. Reprod., 29:767-775. 197. Dacheaux, J. L., and Voglmayr, J. K. (1983): Sequence of sperm cell surface differentiation and its relationship to exogenous fluid proteins in the ram epididymis. Biol. Reprod., 29:1033-1046. 198. Vieurla, M., and Rajaniemi, H. (1980): Radioiodination of surface proteins of bull spermatozoa and their characterization by sodium dodecyl sulfate-polyacrylamide gel electrophoresis. J. Reprod. Fer¬ til., 58:483-489. 199. Nicolson, G. L., Bronginski, A. B., Beattie, G., and Yanagimachi, R. (1979): Cell surface changes in the proteins of rabbit spermatozoa during epididymal passage. Gamete Res., 2:153-162. 200. Russell, L. D., Peterson, R. N., Hunt, W., and Strack, L. E. (1984): Post-testicular surface modifications and contributions of reproduc¬ tive tract fluids to the surface polypeptide composition of boar sper¬ matozoa. Biol. Reprod., 30:959-978. 201. Young, L. G., Hinton, B. T., and Gould, K. G. (1985): Surface changes in chimpanzee sperm during epididymal transit. Biol. Re¬ prod., 32:399-412. 202. Hunter, A. G. (1969): Differentiation of rabbit sperm antigens from those of seminal plasma. J. Reprod. Fertil., 20:413^418. 203. Barker, L. D. S., and Amann, R. P. (1970): Epididymal physiology. I. Specificity of antisera against bull spermatozoa and reproductive fluids. J. Reprod. Fertil., 22:441-452. 204. Barker, L. D. S., and Amann, R. P. (1971): Epididymal physiol¬ ogy. II. Immunofluorescent analysis of epithelial secretion and absorp¬ tion, and of bovine sperm maturation. J. Reprod. Fertil., 26:319332. 205. Killian, G. J., and Amann, R. P. (1973): Immunoelectrophoretic characterization of fluid and sperm entering and leaving the bovine epididymis. Biol. Reprod., 9:489^499. 206. Dravland, E., and Joshi, M. S. (1981): Sperm-coating antigens se¬ creted by the epididymis and seminal vesicle of the rat. Biol. Reprod., 25:649-658. 207. Lea, O. A., Petrusz, P., and French, F. (1978): Purification and localization of acidic epididymal glycoprotein (AEG): A sperm coat¬ ing protein secreted by the rat epipidymis. Int. J. Androl. (Suppl.), 2:592-607. 208. Cameo, M. S., and Blaquier, J. A. (1976): Androgen-controlled specific protein in rat epididymis. J. Endocrinol., 69:47-55. 209. Pholpramol, C., Lea, O. A., Burrow, P. V., Dott, H. M., and Setchell, B. P. (1983): The effects of acidic epididymal glycoprotein (AEG) and some other proteins on the motility of rat epididymal spermatozoa. Int. J. Androl., 6:240-248. 210. Garberi, J. C., Kohane, A. C., Cameo, M. S., and Blaquier, J. A. (1979): Isolation and characterization of specific rat epididymal pro¬ teins. Mol. Cell. Endocrinol., 13:73-82. 211. Kohane, A. C., Gonzales Echeverna, F. M. C., Pineiro, L., and Blaquier, J. A. (1980): Distribution and site of production of specific proteins in rat epididymis. Biol. Reprod., 23:181-187. 212. Kohane, A. C., Gonzales Echeverna, F. M. C., Pineiro, L., and Blaquier, J. A. (1980): Interactions of proteins of epididymal origin with spermatozoa. Biol. Reprod., 23:737-742. 213. Cuasnicu, P. S., Gonzales Echeverna, F., Piazza, A., Cameo, M. S., and Blaquier, J. A. (1984): Antibodies against epididymal glycoproteins block fertilizing ability in rat. J. Reprod. Fertil., 72:461471. 214. Brooks, D. E., and Higgins, S. J. (1980): Characterization and an¬ drogen-dependence of proteins associated with luminal fluid and sper¬ matozoa in the rat epididymis. J. Reprod. Fertil., 59:262-375. 215. Brooks, D. E. (1981): Secretion of proteins and glycoproteins by the rat epididymis: Regional differences, androgen-dependence, and ef¬ fects of protease inhibitors, procaine, and tunicamycin. Biol. Re¬ prod., 25:1099-1117. 216. Brooks, D. E. (1981): Metabolic activity in the epididymis and its regulation by androgens. Physiol. Rev., 61:515-555. 217. Brooks, D. E. (1983): Selective binding of specific rat epididymal secretory proteins to spermatozoa and erythrocytes. Gamete Res., 4:367-376.

62

/ Chapter 2

218. Brooks, D. E., and Tiver, K. (1983): Localization of epididymal secretory proteins on rat spermatozoa. J. Reprod. Fertil., 69:651-657. 219. Rifkin, J., and Olson, G. E. (1985): Characterization of maturationdependent extrinsic proteins of the rat sperm surface. J. Cell Biol., 100:1582-1591. 220. Hamilton, D. W., Wenstrom, J. C., and Baker, J. B. (1986): Mem¬ brane glycoproteins from spermatozoa: Partial characterization of an integral Mr = ~ 24,000 molecule from rat spermatozoa that is glycosylated during spididymal maturation. Biol. Reprod., 34:925936. 221. Olson, G. E., and Orgebin-Crist, M.-C. (1982): Sperm surface changes during epididymal maturation. In: The Cell Biology of the Testis, Vol. 383, Annals of the New York Academy of Sciences, edited by C. W. Bardin and R. J. Sherins, pp. 372-390. New York Academy of Sciences, New York. 222. Moore, H. D. M. (1980): Localization of specific glycoproteins se¬ creted by the rabbit and hamster epididymis. Biol. Reprod., 22:705-718. 223. Gonzales Echiverria, F., Cuasnicu, P. S., and Blaquier, J. A. (1982): Identification of androgen-dependent glycoproteins in the hamster epididymis and their association with spermatozoa. J. Reprod. Fertil., 64:1-7. 224. Tezon, J. G., Ramella, R., Cameo, M. S., Vazquez, M. H., and Blaquier, J. A. (1985): Immunochemical localization of secretory antigens in the human epididymis and their association with sper¬ matozoa. Biol. Reprod., 32:591-597. 225. Vernon, R. B., Muller, C. H., Herr, J. C., Feuchter, F. A., and Eddy, E. M. (1982): Epididymal secretion of a mouse sperm surface component recognized by a monoclonal antibody. Biol. Reprod., 26:523-535. 226. Vernon, R. B., Hamilton, M. S., and Eddy, E. M., (1985): Effects of in vivo and in vitro fertilization environments on the expression of a surface antigen of the mouse sperm tail. Biol. Reprod., 32:669-680. 227. Fox, N., Damjanov, I., Knowles, B. B., and Solter, D. (1982): Teratocarcinoma antigen is secreted by epididymal cells and coupled to maturing sperm. Exp. Cell Res., 137:485-488. 228. Jones, R. (1978): Comparative biochemistry of mammalian epidi¬ dymal plasma. Comp. Biochem. Physiol. (B), 61:365-370. 229. Zaneveld, L. J. D., and Chatterton, R. T. (1982): Biochemistry of Mammalian Reproduction. John Wiley & Sons, New York. 230. Conchie, J., Findlay, J., and Levvy, G. A. (1959): Mammalian glycosidases. Distribution in the body. Biochem. J., 71:318-325. 231. Kemp, W. R., and Killian, G. J. (1978): Glycosidase activity in epididymal epithelial cells isolated from normal and a-chlorohydrin treated male rats. Contraception, 17:93-101. 232. Chapman, D. A., and Killian, G. J. (1984): Glycosidase activities in principal cells, basal cells, fibroblasts and spermatozoa isolated from the rat epididymis. Biol. Reprod., 31:627-636. 233. Skudlarek, M. D., and Orgebin-Christ, M.-C. (1986): Glycosidases in cultured rat epididymal cells: Enzyme activity, synthesis and se¬ cretion. Biol. Reprod., 35:167-178. 234. Grandmont, A.-M., Chapdelaine, P., and Tremblay, R. R. (1983): Presence of a-glucosidases in the male reproductive system of the rat and hormonal influences. Can. J. Biochem. Cell Biol., 61:764-769. 235. Jones, R. (1974): Absorption and secretion in the cauda epididymidis of the rabbit and the effects of degenerating spermatozoa on epidi¬ dymal plasma after castration. J. Endocrinol., 63:157-165. 236. Poirier, G. R., and Jackson, J. (1981): Isolation and characterization of two proteinase inhibitors from the male reproductive tract of mice. Gamete Res., 4:555-569. 237. Wenstrom, J. C., and Hamilton, D. W. (1980): Dolichol concen¬ tration and biosynthesis in rat testis and epididymis. Biol. Reprod., 23:1054-1069. 238. Iusem, N. B., de Larminant, M. A., Tezon, Blaquier, J. A., and Belocopitow, E. (1984): Androgen dependence of protein A-glycosylation in rat epididymis. Endocrinology, 114:1448-1458. 239. Bemal, A., Torres, J., Reyes, A., and Rosada, A. (1980): Presence and regional distribution of sialyl transferase in the epididymis of the rat. Biol. Reprod., 23:290-293. 240. Cossu, G., and Boitani, C. (1984): Lactosaminoglycans synthesized by mouse male germ cells are fucosylated by an epididymal fucosyltransferase. Dev. Biol., 102:402-408. 241. Letts, P. J., Meistrich, M. R., Bruce, W. R., and Schachter H. (1974): Glycoprotein glycosyltransferase levels during spermatogen¬ esis in mice. Biochim. Biophys. Acta, 343:192-207.

242. Reddy, P. R. K., Tadolini, B., Wilson, J., and Williams-Ashman, H. G. (1976): Glycoprotein glycosyltransferase in male reproductive organs and their hormonal regulations. Mol. Cell. Endocrinol., 5:23—31. 243. Tadolini, B., Wilson, J., Reddy, P. R. K., and Williams-Ashman, H. G. (1977): Characteristics and hormonal control of some glyco¬ protein glycosyltransferase reactions in male reproductive organs. Adv. Enzymol. Regul., 15:319-336. 244. Hamilton, D. W. (1980): UDP-galactose: V-acetylglucosamine glalctosyltransferase in fluids from rat testis and epididymis. Biol. Reprod., 23:377-385. 245. Hamilton, D. W., and Gould, R. P. (1982): Preliminary observations on enzymatic galactosylation of glycoproteins on the surface of rat caput epididymal spermatozoa. Int. J. Androl. (Suppl.), 5:73-80. 246. Hamilton, D. W. (1981): Evidence for a-lactalbumin-like activity in reproductive tract fluids of the male rat. Biol. Reprod., 25:385-392. 247. Durr, R., Shur, B., and Roth, S. (1977): Sperm-associated sialyltransferase activity. Nature, 265:547-548. 248. Shur, B. D., and Bennett, D. (1979): A specific defect in galactosyltransferase on sperm bearing mutant alleles of the T/t locus. Dev. Biol., 71:243-259. 249. Shur, B. D., and Hall, N. G. (1982): Sperm surface galactosyltransferase activities during in vitro capacitation. J. Cell Biol., 95:567-573. 250. Klinefelter, G. R., and Hamilton, D. W. (1985): Synthesis and secretion of proteins by perfused caput epididymal tubules, and as¬ sociation of secreted proteins with spermatozoa. Biol. Reprod., 33:1017-1027. 251. Ensrude, K., Wenstrom, J. C., Baker, J. B., and Hamilton, D. W. (1985): A monoclonal antibody against rat epididymal a-lactalbuminlike 24Kd polypeptide recognizes rat cauda sperm surface. J. Androl. (Suppl.) 6:54. 252. Quasba, P. K., Hewlett, I. K., and Byers, S. (1983): The presence of the milk protein, a-lacatalbumin and its mRNA in the rat epidi¬ dymis. Biochem. Biophys. Res. Commun., 117:306-312. 253. Brobek, U., Denton, W. L., Tanahashi, N. andEbner, K. E. (1967): The isolation and identification of the B protein of lactose synthetase as a-lactalbumin. J. Biol. Chem., 242:1391-1397. 254. Shur, B. D., and Hall, N. G. (1982): A role for mouse sperm surface galactosyltransferase in sperm binding to the egg zona pellucida. J. Cell Biol., 95:574-579. 255. Vaidya, R. A., Glass, R. W., Dandekar, P., and Johnson, K. (1971): Decrease in electrophoretic mobility of rabbit spermatozoa following intra-uterine incubation. J. Reprod. Fertil., 24:299-301. 256. Rosado, A., Valezquez, A., and Lara-Ricalde, R. (1973): Cell polarography. II. Effect of neuraminidase and follicular fluid upon the surface characteristics of human spermatozoa. Fertil. Steril., 24:349354. 257. Moore, H. D. M., and Hibbits, K. G. (1975): Isoelectric focusing of boar spermatozoa. J. Reprod. Fertil., 44:329-332. 258. Clegg, E. D., and Foote, R. H. (1973): Phospholipid composition of bovine sperm fractions, seminal plasma and cytoplasmic droplet. J. Reprod. Fertil., 34:379-383. 259. Edwards, R. G., Ferguson, L. C., and Coombs, R. R. A. (1964): Blood group antigens on human spermatozoa. J. Reprod. Fertil., 7:153-161. 260. Boettcher, B. (1968): Correlation between human ABO blood group antigens in seminal plasma and on seminal spermatozoa. J. Reprod. Fertil., 16:49-54. 261. Kerek, G., Biberfeld, P., and Afzelius, B. A. (1973): Demonstration of HL-A antigens, “species,” and “semen”-specific antigens on hu¬ man spermatozoa. Int. J. Fertil., 18:145—155. 262. James, K., and Hargreave, T. B. (1984): Immunosuppression by seminal plasma and its possible clinical significance. Immunol. To¬ day, 5:357-363. 263. Hekman, A., and Rumke, P. (1969): The antigens of human seminal plasma with special reference to lactoferrin, a spermatozoa coating antigen. Fertil. Steril., 20:312-323. 264. Roberts, T. K., and Boettcher, B. (1969): Identification of human sperm coating antigen. J. Reprod. Fertil., 18:347-350. 265. Koyama, Y., Takuda, Y., Takamura, T., and Isojima, S. (1983): Localization of human seminal plasma No. 7 antigen (ferrisplan) in accessory glands of the male genital tract. J. Reprod. Immunol., 5:135-143. 266. Wahlstrom, T., Bohn, H., and Seppala, M. (1982): Immunohistochemical demonstration of placental protein 5 (PP5)-like material in

The Spermatozoon

267.

268.

269.

270.

271.

272.

273.

274.

275. 276.

277. 278. 279.

280.

281.

282. 283.

284.

285.

286.

287.

288.

289.

290.

the seminal vesicle and the ampullar part of the vas deferens. Life Sci., 31:2723-2725. Evans, R. J., and Herr, J. C. (1986): Immunohistochemical locali¬ zation of the MHS-5 antigen in principal cells of human seminal vesicle epithelium. Anat. Rec., 214:372-377. Saji, J., Minagawa, Y., Ohashi, K., Negoro, T., and Tanizawa, O. (1986): Further characterization of a human sperm coating antigen (gpl2). Am. J. Reprod. Immunol., 12:13-16. Abrescia, P., Lombardi, G., De Rosa, M., Quagliozzi, L., Guardiola, J., and Metafora, S. (1985): Identification and preliminary characterization of sperm-binding protein in normal human semen. J. Reprod. Fertil., 73:71-77. Ostrowski, M. C., Kistler, M. K., and Kistler, W. S. (1979): Pu¬ rification and cell-free synthesis of a major protein from rat seminal vesicle secretion. J. Biol. Chem., 254:4007-4021. Draveland, E., and Joshi, M. S. (1981): Sperm-coating antigens secreted by the epididymis and seminal vesicle of the rat. Biol. Reprod., 25:649-658. Oliphant, G., and Singhas, C. A. (1979): Iodination of rabbit sperm plasma membrane: Relationship of specific surface proteins to epididymal function and sperm capacitation. Biol. Reprod., 21:937-944. Koehler, J. K., Nudelman, E. D., and Hakomori, S. (1980): A collagen-binding protein on the surface of ejaculated rabbit sper¬ matozoa. J. Cel! Biol., 86:529-536. Irwin, M., Nicholson, N., Haywood, J. T., and Pourier, G. R. (1983): Immunoflourescent localization of a murine seminal vesicle proteinase inhibitor. Biol. Reprod., 28:1201-1206. Isaacs, W., and Coffey, D. S. (1984): The predominant protein of canine seminal plasma is an enzyme. J. Biol. Chem., 259:11520-11526. Paonessa, G., Metafora, G., Tajana, G., Abrescia, P., De Santis, A., Gentile, V., and Porta, R. (1984): Transglutaminase-mediated modifications of the rat sperm surface in vitro. Science, 226:852-855. Chang, M. C. (1951): Fertilizing capacity of spermatozoa deposited into fallopian tubes. Nature, 168:697-698. Austin, C. R. (1951): Observations on the penetration of the sperm into the mammalian egg. Aust. J. Sci. Res. B, 4:581-596. Bedford, J. M. (1972): Sperm transport, capacitation and fertiliza¬ tion. In: Reproductive Biology, edited by H. Balin and S. Glasser, pp. 338-392. Excerpta Medica, Amsterdam. Barros, C. (1974): Capacitation of mammalian spermatozoa. In: Physiology and Genetics of Reproduction, Part B, edited by E. M. Coutinho and F. Fuchs, pp. 3-24. Plenum Press, New York. Chang, M. C., and Hunter, R. H. F. (1975): Capacitation of mam¬ malian sperm: Biological and experimental aspects. In: Handbook of Physiology, Vol. 5, Endocrinology, Section 7, Male Reproduction, edited by R. O. Greep, pp. 339-351. American Physiological So¬ ciety, Washington D.C. Rogers, B. J. (1978): Mammalian sperm capacitation and fertilization in vitro: A critique of methodology. Gamete Res., 1:165-223. O’Rand, M. J. (1979): Changes in sperm surface properties correlated with capacitation. In: The Spermatozoon, edited by D. W. Fawcett and J. M. Bedford, pp. 195-204. Urban & Schwarzenberg, Balti¬ more. Courtens, J. L., and Foumier-Delpech, S. (1979): Modifications in the plasma membranes of epididymal ram spermatozoa during mat¬ uration and incubation in utero. J. Ultrastruct. Res., 68:136-148. Talbot, P., and Franklin, L. E. (1978): Surface modification of guinea pig sperm during in vitro capacitation: An assessment using lectininduced agglutination of living sperm. J. Exp. Zool., 203:1-14. Friend, D. S. (1977): The organization of the sperm membrane. In: Immunobiology of Gametes, edited by M. Edidin and M. H. Johnson, pp. 5-30. Cambridge University Press, Cambridge. Friend, D. S., Orci, L., Perrelet, A., and Yanagimachi, R. (1977): Membrane particle changes attending the acrosome reaction in guinea pig spermatozoa. J. Cell Biol., 74:561-577. Davis, B. K. (1980): Interaction of lipids with the plasma membrane of sperm cells. I. The antifertilization action of cholesterol. Arch. Androl., 5:249-254. Davis, B. K., Byrne, R., and Hungund, B. (1979): Studies on the mechanism of capacitation. II. Evidence for lipid transfer between plasma membrane of rat sperm and serum albumin during capacitation in vitro. Biochim. Biophys. Acta, 558:257-266. O’Rand, M. J. (1979): Changes in sperm surface properties correlated with capacitation. In: The Spermatozoon, edited by D. W. Fawcett

291.

292.

293.

294.

295. 296.

297.

298.

299.

300.

301.

302.

303.

304.

305. 306.

307.

308.

309.

310.

311.

/

63

and M. J. Bedford, pp. 195-204, Urban & Schwarzenberg, Balti¬ more. Clegg, E. D., Morre, D. J., and Lunstra, D. D. (1975): Porcine sperm membrane: In vivo phospholipid changes, isolation and elec¬ tron microscopy. In: The Biology of Male Gametes, edited by J. G. Duckett and P. A. Racey, pp. 321-335. Academic Press, London. Snider, D. R., and Clegg, E. D. (1975): Alteration of phospholipids in procine spermatozoa during in vivo uterus and oviduct incubation. J. Anim. Sci., 40:269-274. Oliphant, G., and Brackett, B. G. (1973): Capacitation of mouse spermatozoa in media with elevated ionic strength and reversible decapacitation with epididymal extracts. Fertil. Steril., 24:948-955. Aonuma, S., Mayumi, T., Suzuki, K., Noguchi, T., Iwai, M., and Okabe, M. (1973): Studies on sperm capacitation. I. The relationship between a guinea-pig sperm-coating antigen and a sperm capacitation phenomenon. J. Reprod. Fertil., 35:425-432. Brackett, B. G., and Oliphant, G. (1975): Capacitation of rabbit spermatozoa in vitro. Biol. Reprod., 12:260-274. Johnson, M. H. (1975): The macromolecular organization of mem¬ branes and its bearing on events leading up to fertilization. J. Reprod. Fertil., 44:167-184. Schill, W. B., Heimburger, N., Schiessler, H., Stolla, R., and Fritz, H. (1975): Reversible attachment and localization of acid-stable sem¬ inal plasma acrosin-trypsin inhibitors on boar spermatozoa as re¬ vealed by the indirect immunofluorescent staining technique. Biol. Chem. Hoppe Seyler, 356:1473-1476. Koehler, J. K. (1976): Changes in antigenic site distribution on rabbit spermatozoa after incubation in “capacitating” media. Biol. Reprod., 9:444-456. Eng, L. A., and Oliphant, G. (1978): Rabbit sperm reversible ca¬ pacitation by membrane stabilization with a highly purified glyco¬ protein from seminal plasma. Biol. Reprod., 19:1083-1094. Johnson, W. L., and Hunter, A. G. (1972): Seminal antigens: Their alteration in the genital tract of female rabbits and during partial in vitro capacitation with beta amylase and beta glucuronidase. Biol. Reprod., 7:332-340. Oliphant, G., and Brackett, B. G. (1973): Immunological assessment of surface changes of rabbit sperm undergoing capacitation. Biol. Reprod., 9:404-414. Myles, D. G., and Primakoff, P. (1984): Localized surface antigens of guinea pig sperm migrate to new regions prior to fertilization. J. Cell Biol., 99:1634-1641. Baccetti, B. (1984): The human spermatozoon. In: Ultrastructure of Reproduction, edited by J. Van Blerkom and P. M. Motta, pp. 110126. Martinus Nijhoff, The Hague. Wyrobek, A. J., Gordon, L. A., Burkhart, J. G., Francis, M. W., Kapp, R. W. Jr., Letz, G., Mailing, H. V., Topham, J. C., and Whorton, M. D. (1983): An evaluation of human sperm as indicators of chemically induced alterations of spermatogenic function. A report of the U.S. Environmental Protection Agency Gene-Tox Program. Mutat. Res., 115:73-148. Grimes, S. R. Jr. (1986): Nuclear proteins in spermatogenesis. Comp. Biochem. Physiol., 83D:495-500. Coelingh, J. P., Monfoort, C. H., Rozijin, T. H., Gevers-Leuven, J. A., Shiphof, K., Steyn-Parve, E. P., Brauntizer, G., Shrank, B., and Ruhfus, A. (1972): The complete amino acid sequence of the basic nuclear protein of bull spermatozoa. Biochim. Biophys. Acta, 285:1-14. Kistler, W. S., Keim, P. S., and Heinrickson, R. L. (1976): Partial structural analysis of the basic chromosomal protein of rat sperma¬ tozoa. Biochim. Biophys. Acta, 427:931-954. Hecht, N. B., Bower, P. A., Waters, S. H., Yelick, P. C., and Distel, R. J. (1986): Evidence for haploid expression of mouse tes¬ ticular genes. Exp. Cell Res., 164:183-190. Bellve, A. R., and Carraway, R. (1978): Characterization of two basic chromosomal proteins isolated from mouse spermatozoa. J. Cell Biol., 79:177a. Mayer, J. F., Chang, T. S. K., and Zirkin, B. R. (1981): Sper¬ matogenesis in the mouse. 2. Amino acid incorporation into basic nucleoproteins of mouse spermatids and spermatozoa. Biol. Reprod., 25:1041-1051. Balhom, R., Weston, S., Thomas, C., and Wyrobek, A. J. (1984): DNA packaging in mouse spermatids. Synthesis of protamine var¬ iants and four transition proteins. Exp. Cell Res., 150:298-308.

64

/ Chapter 2

312. Bellve, A. R., and O’Brien, D. A. (1983): The mammalian sper¬ matozoon: Structure and temporal assembly. In: Mechanisms and Control of Animal Fertilization, edited by J. F. Hartmann, pp. 55137. Academic Press, Orlando, Florida. 313. Balhom, R. (1982): A model for the structure of chromatin in mam¬ malian sperm. J. Cell Biol., 93:298-305. 314. Warrent, R. W., and Kim. S.-H. (1978): a-Helix-double helix in¬ teraction shown in the structure of a protamine-transfer RNA complex and a nucleoprotamine model. Nature, 271:130-135. 315. Gusse, M., and Chevaillier, P. (1980): Electron microscopic evi¬ dence for the presence of globular structures in different sperm chro¬ matins. J. Cell Biol., 87:280-284. 316. Tsanev, R., and Avramova, Z. (1981): Nonprotamine nucleoprotein ultrastructures in mature ram sperm nuclei. Eur. J. Cell Biol., 24:139145. 317. Bendet, I. J., and Bearden, J., Jr. (1972): Birefringence of bull sperm. II. Form birefringence of bull sperm. J. Cell Biol., 55:501-510. 318. Sipski, M. R., and Wagner, T. E. (1977): The total structure and organization of chromosomal fibers in eutherian sperm nuclei. Biol. Reprod., 16:428-440. 319. Bellve, A. R. (1982): Biogenesis of the mammalian spermatozoon. In: Prospects for Sexing Mammalian Sperm, edited by R. P. Amann and G. E. Seidel, Jr., pp. 69-102. Colorado Associated University Press, Boulder. 320. Stackpole, C. W., and Devorkin, D. (1974): Membrane organization in mouse spermatozoa revealed by freeze-etching. J. Ultrastruct. Res., 49:167-187. 321. Gerace, L., Comeau, C., and Benson, M. (1984): Organization and modulation of nuclear lamina structure. J. CellSci. (Suppl.), 1:137-160. 322. Krohne. G., and Benavente, R. (1986): The nuclear lamins. A mul¬ tigene family of proteins in evolution and differentiation. Exp. Cell Res., 162:1-10. 323. Gerace, L., Blum, A., and Blobel, G. (1978): Immunocytochemical localization of the major polypeptides of the nuclear pore complex lamina fraction. Interphase and mitotic distribution. J. Cell Biol., 79:546-566. 324. Hancock, R., and Baulikis, T. (1982): Functional organisation of the nucleus. In: International Review of Cytology, Vol. 79, edited by G. H. Bourne and J. F. Danielli, pp. 165-214. Academic Press, New York. 325. Lebkowski, Y. S., and Laemmli, U. K. (1982): Non-histone proteins and long-range organization of HeLa interphase DNA. J. Mol. Biol., 156:121-141. 326. Gerace, L., and Blobel, G. (1980): The nuclear envelope lamina is reversibly depolymerized during mitosis. Cell, 34:13-23. 327. Burke, B., Tooze, J., and Warren, G. (1983): A monoclonal antibody which recognizes each of the nuclear lamin polypeptides in mam¬ malian cells. EMBO J., 1:1621-1628. 328. Krone, G., Debus, E., Osborn, W., and Franke, W. W. (1984): A monoclonal antibody against nuclear lamina proteins reveals cell type-specificity in Xenopus laevis. Exp. Cell Res., 150:47-59. 329. Stick, R., and Schwarz, H. (1982): The disappearance of the nuclear lamina during spermatogenesis: An electron microscopic and im¬ munofluorescence study. Cell Differ., 11:235-243. 330. Hogner, D., Telling, A., Lepper, K., and Jost, E. (1984): Patterns of nuclear lamins in diverse animal and plant cells and in germ cells as revealed by immunofluorescence microscopy with polyclonal and monoclonal antibodies. Tissue Cell, 16:693-703. 331. Maul, G. G., French, B. T., and Bechtol, K. B. (1986): Identification and redistribution of lamins during nuclear differentiation in mouse spermatogenesis. Dev. Biol., 115:68-77. 332. McKeon, F. D., Kirschner, M. W., and Caput, D. (1986): Homol¬ ogies in both primary and secondary structure between nuclear en¬ velope and intermediate filament proteins. Nature, 319:463-468. 333. Fisher, D. Z., Chaudhary, N., and Blobel, G. (1986): cDNA se¬ quencing of nuclear lamins A and C reveals primary and secondary structural homology to intermediate filament proteins. Proc. Natl. Acad. Sci. USA, 83:6450-6454. 334. Benavente, R., and Krohne, G. (1985): Changes of karyoskeleton during spermatogenesis of Xenopus: Expression of lamin LIV, a nuclear lamina protein specific for the male germ line. Proc. Natl. Acad. Sci, USA, 82:6176-6180. 335. Olson, G. E., Hamilton, D. W., and Fawcett, D. W. (1976): Isolation and characterization of the perforatorium of rat spermatozoa. J. Reprod. Fertil., 47:293-297.

336. Fawcett, D. W. (1970): A comparative view of sperm ultrastructure. Biol. Reprod. (Suppl.) 2:90-127. 337. Calvin, H. I., and Bedford, J. M. (1971): Formation of disulfide bonds in the nucleus and accessory structures of mammalian sper¬ matozoa durirfg maturation in the epididymis. J. Reprod. Fertil. (Suppl.), 13:65-75. 338. Huang, T. T. F., and Yanagimachi, R. (1985): Inner acrosomal membrane of mammalian spermatozoa: Its properties and possible functions in fertilization. Am. J. Anat., 174:249-268. 339. Burgos, M. H., and Fawcett, D. W. (1956): An electron microscopic study of spermatid differentiation in the toad, Bufo arenarum Hensel. J. Biophys. Biochem., Cytol., 2:223-240. 340. Nagano, T. (1962): Observations on the fine structure of the devel¬ oping spermatid in the domestic chicken. J. Cell Biol., 14:193-205. 341. Clermont, Y., Einberg, E., Leblond, C. P., and Wagner, S. (1955): The perforatorium—An extension of the nuclear membrane of the rat spermatozoon. Anat. Rec., 121:1-12. 342. Yanagimachi, R., and Noda, Y. D. (1970): Ultrastructural changes in the hamster sperm head during fertilization. J. Ultrastruct. Res., 31:465-485. 343. Austin, C. R., and Bishop, M. W. H. (1958): Some features of the acrosome and perforatorium in mammalian spermatozoa. Proc. R. Soc. Lond. (Biol.), 149:234-240. 344. Olson, G. E. (1979): Isolation of the fibrous sheath and perforatorium of rat spermatozoa. In: The Spermatozoon, edited by D. W. Fawcett and J. M. Bedford, pp. 395-400. Urban & Schwarzenberg, Balti¬ more. 345. Courtens, J. L., Courot, M., and Flechon, J. E. (1976): The per¬ inuclear substance of boar, bull, ram and rabbit spermatozoa. J. Ultrastruct. Res., 57:54-64. 346. Lalli, M., and Clermont, Y. (1981): Structural changes in the head component of the rat spermatid during late spermatogenesis. Am. J. Anat., 160:419-434. 347. Nicander, L., and Bane, A. (1966): Fine structure of the sperm head in some mammals with particular reference to the acrosome and subacrosomal substance. Z. Zelforsch., 72:496-515. 348. Pedersen, H. (1972): The postacrosomal region of man and Macaca artoides. J. Ultrastruct. Res., 40:366-377. 349. Phillips, D. M. (1975): Cell surface structure of rodent sperm heads. J. Exp. Zool., 191:1-8. 350. Olson, G. E., Noland, T. D., Winfrey, V. P., and Garbers, D. L. (1983): Substructure of the postacrosomal sheath of bovine sper¬ matozoa. J. Ultrastruct. Res., 85:204-218. 351. Maxwell, W. L. (1982): The acrosomal zonule. Tissue Cell, 14:283288. 352. Czaker, R. (1985): Morphogenesis and cytochemistry of the posta¬ crosomal dense lamina during mouse spermiogenesis. J. Ultrastruct. Res., 90:26-39. 353. Searle, A. G. (1982): The genetics of sterility in the mouse. In: Genetic Control of Gamete Production and Function, edited by P. G. Crosignai, B. L. Rubin, and M. Fraccaro, pp. 93-114. Grune & Stratton, New York. 354. Sotomayor, R. E., and Handel, M. A. (1986): Failure of acrosome assembly in a male sterile mutant. Biol. Reprod., 34:171-182. 355. Phillips, D. M. (1970): Development of spermatozoa in the woolly opossum with special reference to the shaping of the sperm head. J. Ultrastruct. Res., 33:369-380. 356. Fawcett, D. W., and Phillips, D. M. (1969): Observations on the release of spermatozoa and on changes in the head during passage through the epididymis. J. Reprod. Fertil., (Suppl.), 6:405-418. 357. Fawcett, D. W., Anderson, W. A., and Phillips, D. M. (1971): Morphogenetic factors influencing the shape of the sperm head. Dev Biol., 26:220-251. 358. Hunt, D. M., and Johnson, D. R. (1971): Abnormal spermiogenesis in two pink-eyed sterile mutants in the mouse. J. Embryol. Exp. Morphol., 26:111-121. 359. Bryan, J. H. D. (1977): Spermatogenesis revisited: III. The course of spermatogenesis in a male-sterile pink-eyed mutant type in the mouse. Cell Tissue Res., 180:173-186. 360. Wooding, F. B. P. (1973): The effect of Triton X-100 on the ultra¬ structure of ejaculated bovine sperm. J. Ultrastruct. Res., 42:502-516. 361. Phillips, D. M. (1972): Substructure of the mammalian acrosome. J. Ultrastruct. Res., 38:591-604. 362. Pedersen, H. (1972): Further observations on the fine structure of the human spermatozoon. Z. Zellforsch., 123:305-315.

The Spermatozoon

363. Koehler, J. K. (1975): Periodicities in the acrosome or acrosomal membrane: Some observations on mammalian spermatozoa. Biol. J. Linnean Soc. (Suppl.), 1:337-342. 364. Zahler, W. L., and Doak, G. A. (1975): Isolation of the outer ac¬ rosomal membrane from bull spermatozoa. Biochim. Biophys Acta 406:479-488. 365. Russell, L., Peterson, R., and Freund, M. (1979): Direct evidence for formation of hybrid vesicles by fusion of plasma and outer ac¬ rosomal membranes during the acrosome reaction in boar sperma¬ tozoa. J. Exp. Zool., 208:41-56. 366. Noland, T. D., Olson, G. E., and Garbers, D. L. (1983): Purification and partial characterization of plasma membranes from bovine sper¬ matozoa. Biol. Reprod., 29:987-998. 367. Topfer-Petersen, E., and Schill, W. B. (1981): A new separation method of subcellular fractions of boar spermatozoa. Andrologia, 13:174-176. 368. Olson, G. E., Winfrey, V. P., Garbers, D. L., and Noland, T. D. (1985): Isolation and characterization of a macromolecular complex associated with the outer acrosomal membrane of bovine sperma¬ tozoa. Biol. Reprod., 33:761-779. 369. Burgos, M. H., and Fawcett, D. W. (1955): Studies on the fine structure of the mammalian testis. I. Differentiation of the spermatids in the cat (Felis domestica). J. Biophys. Biochem. Cytol., 1:287-299. 370. Hermo, L., Rambourg, L. A., and Clermont, Y. (1980): Threedimensional architecture of the cortical region of the Golgi apparatus in rat spermatids. Am. J. Anat.. 157:357-373. 371. Thakkar, J. K., East, J., Seyler, D., and Fanson, R. C. (1983): Surface-active phospholipase A2 in mouse spermatozoa. Biochim. Biophys. Acta, 754:44-50. 372. Rahi, H., Sheikhnejade, G., and Srivastava, P. N. (1983): Isolation of the inner acrosomal-nuclear membrane complex from rabbit sper¬ matozoa. Gamete Res., 7:215-225. 373. Russell, L., Peterson, R. N., and Freund, M. (1980): On the presence of bridges linking the inner and outer acrosomal membranes of boar spermatozoa. Anat. Rec., 198:449-459. 374. Yanagimachi, R. (1977): Specificity of sperm-egg interaction. In: Immunobiology of Gametes, edited by M. Edidin and M. H. Johnson, pp. 255-296. Cambridge University Press, London. 375. Cowen, A. E., Primakoff, P., and Myles, D. G. (1986): Sperm exocytosis increases the amount of PH-20 antigen on the surface of guinea pig sperm. J. Cell Biol., 103:1289-1297. 376. Allison, A. C., and Hartree, E. F. (1970): Lysosomal enzymes in the acrosome and their possible role in fertilization. J. Reprod. Fertil., 21:501-515. 377. Polakoski, K. L., and Parrish, R. F. (1977): Boar proacrosin. Pu¬ rification and preliminary activation studies of proacrosin isolated from ejaculated boar sperm. J. Biol. Chem., 252:1888-1894. 378. Tobias, P. S., and Schumacher, G. F. B. (1977): Observation of two proacrosins in extracts of human spermatozoa. Biochem. Bio¬ phys. Res. Commun., 74:434-439. 379. Brown, C. R., and Harrison, R. A. P. (1978): The activation of proacrosin in spermatozoa from ram, bull and boar. Biochim. Bio¬ phys. Acta, 526:202-217. 380. Mukerji, S. K., and Meizel, S. (1979): Rabbit testis proacrosin. Purification, molecular weight estimation, and amino acid and car¬ bohydrate composition of the molecule. J. Biol. Chem., 254:1172111728. 381. Miiller-Esterl, W., and Fritz, H. (1981): Sperm acrosin. In: Methods in Enzymology, Vol. 80, edited by L. Lorand, pp. 621-632. Aca¬ demic Press, New York. 382. Stambaugh, R., and Buckley, J. (1969): Identification and subcellular localization of the enzymes effecting penetration of the zona pellucida by rabbit spermatozoa. J. Reprod. Fertil., 19:423-432. 383. Zaneveld, L. J. D., Robetson, R. T., Kessler, M., and Williams, W. L. (1971): Inhibition of fertilization in vivo by pancreatic and seminal plasma trypsin inhibitors. J. Reprod. Fertil., 25:387-392. 384. Mukerji, S. K., and Meizel, S. (1975): The molecular transformation of rabbit testis proacrosin into acrosin. Arch. Biochem. Biophys., 168:720-721. 385. Gamer, D. L., and Easton, M. P. (1977): Immunofluorescent lo¬ calization of acrosin in mammalian spermatozoa. J. Exp. Zool., 200:157-162. 386. Morton, D. B. (1975): Acrosomal enzymes: Immunochemical lo¬ calization of acrosin and hyaluronidase in ram spermatozoa. J. Re¬ prod. Fertil., 45:375-378.

/

65

387. Morton, D. B. (1977): Lysosomal enzymes in mammalian sper¬ matozoa. In: Immunobiology of Gametes, edited by M. Edidin and M. H. Johnson, pp. 115-155. Cambridge University Press, Lon¬ don. 388. Green, D. P. L., and Hockaday. A. R. (1978): The histochemical localization of acrosin in guinea-pig sperm after the acrosome re¬ action. J. Cell Sci., 32:177-184. 389. McRorie, R. A., and Williams, W. L. (1974): Biochemistry of mam¬ malian fertilization. Ann. Rev. Biochem., 43:777-803. 390. Stambaugh, R. (1978): Enzymatic and morphological events in mam¬ malian fertilization. Gamete Res., 1:65-85. 391. Meizel, S. (1978): The mammalian sperm acrosome reaction. In: Development in Mammals, Vol. 3, edited by M. H. Johnson, pp. 162. North-Holland, Amsterdam. 392. Parrish, R. F., and Polakoski, K. L. (1979): Mammalian sperm proacrosin-acrosin system. Int. J. Biochem., 10:391-395. 393. Siegel, M. S., Bechtold, D. S., Kopta, C. I., and Polakoski, K. L. (1986): Quantification and partial characterization of the hamster sperm proacrosin-acrosin system. Biol. Reprod., 35:485-491. 394. Brown, C. R., and Hartree, E. F. (1978): Studies on ram acrosin. Activation of proacrosin accompanying the isolation of acrosin from spermatozoa and purification of the enzyme by affinity chromatog¬ raphy. Biochem. J., 175:227-238. 395. Kallojoki, M., and Suominen, J. (1984): An acrosomal antigen of human spermatozoa and spermatogenic cells characterized with a monoclonal antibody. Int. J. Androl., 7:283-296. 396. Kallojoki, M., Parvinen, M., and Suominen, J. J. O. (1986): Expres¬ sion of acrosin during mouse spermatogenesis: A biochemical and immunocytochemical analysis by a monoclonal antibody C 11 H. Biol. Reprod., 35:157-165. 397. Phi-van, L., Miiller-Esterl, W., Florke, S., Schmid, M., and Engel, W. (1983): Proacrosin and the differentiation of the spermatozoa. Biol. Reprod., 29:479-486. 398. Florke, S., Phi-van, L., Miiller-Esterl, W., Scheuber, H.-P., and Engel, W. (1983): Acrosin in the spermiohistogenesis of mammals. Differentiation, 24:250-256. 399. Zaneveld, L. J. D., Polakoski, K. L., and Schumacher, G. F. B. (1973): Properties of acrosomal hyaluronidase from bull spermato¬ zoa. Evidence for its similarity to testicular hyaluronidase. J. Biol. Chem., 248:564-570. 400. Yang, C.-H., and Srivastava, P. N. (1975): Purification and prop¬ erties of hyaluronidase from bull sperm. J. Biol. Chem., 250:79-83. 401. Goldberg, E. (1977): Isozymes in testes and spermatozoa. In: Iso¬ zymes: Current Topics in Biological and Medical Research, Vol. 1, edited by M. Ratazzi, J. Scandalios, and G. Whitt, pp. 79-124. Alan R. Liss, New York. 402. Brown, C. R. (1981): Distribution of hyaluronidase in the ram sper¬ matozoon. J. Reprod. Fertil., 45:537-539. 403. Mancini, R. E., Alonso, A., Barquet, J., andNemirovski, B. (1964): Histo-immunological localization of hyaluronidase in bull testis. J. Reprod. Fertil., 8:325-330. 404. Gould, S. F., and Bernstein, M. H. (1975): Localization of bovine sperm hyaluronidase. Differentiation, 3:123-132. 405. Morton, D. B. (1975): Acrosomal enzymes: Immunological locali¬ zation of acrosin and hyaluronidase in ram spermatozoa. J. Reprod. Fertil., 45:375-378. 406. Brown, C. R. (1975): Distribution of hyaluronidase in the ram sper¬ matozoa. J. Reprod. Fertil., 45:537-539. 407. Koren, E., and Milkovtc, S. (1973): “Collagenase-like” peptidase in human, rat and bull spermatozoa. J. Reprod. Fertil., 32:349-356. 408. Erickson, R. P., and Martin, S. R. (1974): The relationship of mouse spermatozoal to mouse testicular cathepsins. Arch. Biochem. Bio¬ phys., 165:114-120. 409. Schollmeyer, J. E. (1986): Identification of calpain II in porcine sperm. Biol. Reprod., 34:721-731. 410. Srivastava, P. N., and Abou-Issa, H. (1977): Purification and prop¬ erties of rabbit spermatozoal acrosomal neuraminidase. Biochem J 161:193-200. 411. Bryan, J. H. D., and Unithan, R. R. (1972): Non-specific esterase activity in bovine acrosomes. Histochem. J., 4:413-419. 412. Meizel, S., and Cotham, J. (1972): Partial characterization of a new bull sperm arylaminidase. J. Reprod. Fertil., 28:303-307. 413. Bhalla, V. K., Tillman, W. L., and Williams, W. L. (1973): Presence of (3-aspartyl A-acetylglucosamine amido hydrolase in mammalian spermatozoa. J. Reprod. Fertil., 34:137-139.

66 / Chapter 2

414. Dudkiewicz, A. B. (1984): Purification of boar acrosomal arylsulfatase A and possible role in the penetration of cumulus cells. Biol. Reprod., 30:1005-1014. 415. Gonzales, L. W., and Meizel, S. (1973): Acid phosphatases of rabbit spermatozoa. II. Partial purification and biochemical characterization of the multiple forms of rabbit spermatozoan acid phosphatase. Biochim. Biophys. Acta., 320:180-194. 416. Stambaugh, R., and Buckley, J. (1970): Comparative studies of the acrosomal enzymes of rabbit, rhesus monkey and human sperma¬ tozoa. Biol. Reprod., 3:275-282. 417. Bhattacharyya, A. K., and Zaneveld, L. J. D. (1982): The sperm head. In: Biochemistry of Mammalian Reproduction, edited by L. J. D. Zaneveld and R. T. Chatterton, pp. 119-152. John Wiley & Sons, New York. 418. Zamboni, L., and Stefanini, M. (1971): The fine structure of the neck of mammalian spermatozoa. Anat. Rec., 169:155-172. 419. Oura, C. (1971): The ultrastructure and development of the neck region of the golden hamster spermatozoon. Monitore Zool. Ital., 5:253-264. 420. Phillips, D. M. (1972): Comparative analysis of mammalian sperm motility. J. Cell Biol., 53:561-573. 421. Millette, C. F., Spear, P. G., Gall, W. E., and Edelman, G. M. (1973): Chemical dissection of mammalian spermatozoa. J. Cell Biol., 58:662-675. 422. Young, R. J., and Cooper, G. W. (1979): Separation of the head and tail of mammalian spermatozoa by primary amines: Evidence for their junction by Schiff bases. In: The Spermatozoon, edited by D. W. Fawcett and J. M. Bedford, pp. 391-394. Urban & Schwarzenberg, Baltimore. 423. Calvin, H. I. (1976): Isolation and subfractionation of mammalian sperm heads and tails. In: Methods in Cell Biology, Vol. 13, edited by D. M. Prescott, pp. 85-104. Academic Press, New York. 424. Calvin, H. I. (1979): Isolation of stable structures from rat sper¬ matozoa. In: The Spermatozoon, edited by D. W. Fawcett and J. M. Bedford, pp. 387-389. Urban & Schwarzenberg, Baltimore. 425. Bedford, J. M., and Calvin, H. I. (1974): Changes in the -S-S- linked structures of the sperm tail during epididymal maturation with com¬ parative observations in sub-mammalian species. J. Exp. Zool., 187:181-204. 426. Bellve, A. R., Anderson, E., and Hanley-Bowdoin, L. (1975): Syn¬ thesis and amino acid composition of basic proteins in mammalian sperm nuclei. Dev. Biol., 47:349-365. 427. Mann, T. (1964): The Biochemistry of Semen and of the Male Re¬ productive Tract, Methuen, London. 428. Blom, E., and Birch-Anderson, A. (1970): The ultrastructure of de¬ capitated sperm defect in Guernsey bulls. J. Reprod. Fertil., 23:67-72. 429. Fawcett, D. W., and Phillips, D. W. (1969): The fine structure and development of the neck region of the mammalian spermatozoon. Anat. Rec., 165:153-184. 430. Nicander, L., and Bane, A. (1962): Fine structure of boar sperma¬ tozoa. Z. Zellforsch., 57:390^405. 431. Illison, L. (1966): Fine structure of the mature spermatozoan head and neck of the mouse. J. Anat., 100:949-950. 432. Woolley, D. M., and Fawcett, D. W. (1973): The degeneration and disappearance of the centrioles during the development of the rat spermatozoon. Anat. Rec., 177:289-302. 433. Fawcett, D. W., and Porter, K. R. (1954): A study of the fine structure of ciliated epithelia. J. Morphol., 94:221-281. 434. Gibbons, I. R., and Grimstone, A. V. (1960): On flagellar structure in certain flagellates. J. Biophys. Biochem. Cytol., 7:697-716. 435. Olson, G. E., and Linck, R. W. (1977): Observations of the structural components of flagellar axonemes and central pair microtubules from rat sperm. J. Ultrastruct. Res., 61:21^43. 436. Bryan, J., and Wilson, L. (1971): Are cytoplasmic microtubules heteropolymers? Proc. Natl. Acad. Sci. USA, 8:1762-1766. 437. Hecht, N. B., Kleene, K. C., Distel, R. J., and Silver, L. M. (1984): The differential expression of the actins and tubulins during sper¬ matogenesis in the mouse. Exp. Cell Res., 153:275-280. 438. Distel, R. J., Kleene, K. C., and Hecht, N. B. (1984): Haploid expression of a mouse testis a-tubulin gene. Science, 224:68-70. 439. Vale, R. D., Reese, T. S., and Sheetz, M. P. (1985): Identification of a novel force generating protein, kinesin, involved in microtubulebased motility. Cell, 42:39-50. 440. Vale, R. D., Schnapp, B. J., Mitchison, T., Steuer, E., Reese, T. S., and Sheetz, M. P. (1985): Different axoplasmic proteins gener¬

ate movement in opposite directions along microtubules in vitro. Cell, 43:623-632. 441. Gibbons, I. R., and Rowe, A. J. (1965): Dynein: A protein with ATPase activity from cilia. Science, 149:424-426. 442. Gibbons, I. R., and Fronk, E. (1972): Some properties of bound and soluble dynein from sea urchin flagella. J. Cell Biol., 54:365-381. 443. Baccetti, B., Burrini, A. G., Pallini, V., and Renieri, T. (1981): Human dynein and sperm pathology. J. Cel! Biol., 88:102-107. 444. Stephens, R. E. (1974): Enzymatic and structural proteins of the axoneme. In: Cilia and Flagella, edited by M. A. Sleigh, pp. 3976. Academic Press, New York. 445. Linck, R. W. (1976): Flagellar doublet microtubules: Fractionation of minor components and a-tubulin from specific regions of the Atubule. J. Cell Sci., 20:405-539. 446. Linck, R. W., and Langevin, G. L. (1982): Structure and chemical composition of insoluble filamentous components of sperm flagellar microtubules. J. Cell Biol., 58:1-22. 447. Linck, R. W. (1982): The structure of microtubules. Ann. N.Y. Acad. Sci., 383:98-121. 448. Linck, R. W., Albertini, D. F., Kenny, D. M., and Langevin, G. L. (1982): Tektin filaments: Chemically unique filaments of sperm flagellar microtubules. Cell Motil. (Suppl.), 1:127-132. 449. Linck, R. W., Amos, L. A., and Amos, W. B. (1985): Localization of tektin filaments in microtubules of sea urchin flagella by immunoelectron microscopy. J. Cell Biol., 100:126-135. 450. Woolley, D. M. (1970): The midpiece of the mouse spermatozoon: Its form and development as seen by surface replication. J. Cell Sci., 6:865-879. 451. Woolley, D. M. (1970): Selection for the length of the spermatozoan midpiece in the mouse. Genet. Res., 16:225-228. 452. Phillips, D. M. (1970): Ultrastructure of spermatozoa of the woolly opossum Caluromys philander. J. Ultrastruct. Res., 33:381-397. 453. Phillips, D. M. (1977): Mitochondrial disposition in mammalian spermatozoa. J. Ultrastruct. Res., 58:144-154. 454. Serres, C., Escalier, D., and David, G. (1983): Ultrastructural mor¬ phometry of human spermatozoa flagellum with a sterological anal¬ ysis of the lengths of the dense fibers. Biol. Cell, 49:153-162. 455. Lindemann, C. B., Fentie, I., and Rikmenspoel, R. (1980): A se¬ lective effect of Ni2+ on wave initiation in bull sperm flagella. J. Cell Biol., 87:420-426. 456. Telkka, A., Fawcett, D. W., and Christensen, A. K. (1961): Further observations on the structure of the mammalian sperm tail. Anat. Rec., 141:231-246. 457. de Kretser, D. M. (1969): Ultrastructural features of human spermiogenesis. Z. Zellforsch., 98:229-236. 458. Sapsford, C. S., Rae, C. A., and Cleland, K. W. (1970): Ultrastruc¬ tural studies on the development and form of the principal piece sheath of the Bandicoot spermatozoon. Aust. J. Zool., 8:21-48. 459. Irons, M. J., and Clermont, Y. (1982): Formation of the outer dense fibers during spermiogenesis in the rat. Anat. Rec., 202:463-471. 460. Gordon, M., and Bensch, K. G. (1968): Cytochemical differentiation of the guinea pig sperm flagellum with phosphotungstic acid. J. Ultrastruct. Res., 24:33-50. 461. Olson, G. E., and Sammons, D. W. (1980): Structural chemistry of outer dense fibers of rat sperm. Biol. Reprod., 22:319-332. 462. Woolley, D. M. (1971): Striations in the peripheral fibers of rat and mouse spermatozoa. J. Cell Biol., 49:936-939. 463. Baccetti, B., Pallini, V., and Burrini, A. G. (1976): The accessory fibers of the sperm tail. III. High-sulfur and low-sulfur components in mammals and cephalopods. J. Ultrastruct. Res., 57:289-308. 464. Pihlaja, D. J., and Roth, L. E. (1973): Bovine sperm fractionation. II. Morphology and chemical analysis of tail segments. J. Ultrastruct Res., 44:293-309. 465. Baccetti, B., Pallini, V., and Burrini, A. G. (1973): The accessory fibers of the sperm tail. I. Structure and chemical composition of the bull coarse fibers. J. Submicrosc. Cytol., 5:237-256. 466. Zittle, C. W., and O’Dell, R. A. (1941): Chemical studies of bull spermatozoa. Lipid, sulfur, cystine, nitrogen, phosphorus, and nu¬ cleic acid content of whole spermatozoa and of the parts obtained by physical means. J. Biol. Chem., 140:899-907. 467. Price, J. M. (1973): Biochemical and morphological studies of outer dense fibers of rat spermatozoa. J. Cell Biol., 59:272a. 468. Calvin, H. I. (1979): Electrophoretic evidence for the identity of the major zinc-binding polypeptides in the rat sperm tail. Biol. Reprod 21:873-882.

The Spermatozoon 469. Vera, J. C., Brito, M., Zuvic, T., and Burzio, L. O. (1984): Poly¬ peptide composition of rat sperm outer dense fibers. J. Biol. Chem., 259:5970-5977. 470. Gunn, S. A., and Gould, T. C. (1970): Cadmium and other mineral elements. In: The Testis, Vol. Ill, edited by A. D. Johnson, W. R. Gomes, and N. L. Vandemark, pp. 377^481. Academic Press, New York. 471. Miller, M. J., Vincent, N. R., and Mawson, C. A. (1961): An autoradiographic study of the distribution of zinc-65 in rat tissues. J. Histochem. Cytochem., 9:111-125. 472. Bradley, F. M., Meth, B. M., and Bellve, A. R. (1981): Structural proteins of the mouse spermatozoan tail: An electrophoretic analysis. Biol. Reprod., 24:691-701. 473. Fawcett, D. W., and Phillips, D. M. (1970): Recent observations on the ultrastructure and development of the mammalian spermato¬ zoon. In: Comparative Spermatology, edited by B. Baccetti, pp. 1328. Academic Press, New York. 474. Cleland, K. W., and Lord Rothschild (1959): The bandicoot sper¬ matozoon: An electron microscopic study of the tail. Proc. R. Soc. B, 150:24-42. 475. Nelson, L. (1958): Cytochemical studies with the electron micro¬ scope. I. Adenosine triphosphatease in the rat spermatozoa. Biochem. Biophys. Acta, 27:634-641. 476. Nagano, T. (1965): Localization of adenosine triphosphatase activity in the rat sperm tail as revealed by electron microscopy. J. Cell Biol., 25:101-112. 477. Irons, M. J., and Clermont, Y. (1982): Formation of the outer dense fibers during spermiogenesis in the rat. Anat. Rec., 202:463-471. 478. Rattner, J. B., and Brinkley, B. R. (1970): Ultrastructure of mam¬ malian spermiogenesis. I. A tubular complex in developing sperm of the cottontop marmoset Sequinus oedipus. J. Ultrastruct. Res., 32:316-322. 479. Wartenberg, H., and Holstein, A. F. (1975): Morphology of the “Spindle-shaped body” in the developing tail of human spermatids. Cell Tissue Res., 159:435-443. 480. Koyama, Y., Shinomiya, T., Sakai, Y., Shiba, T., and Yanagisawa, K. O. (1984): Identification of sperm antigenic determinants with phylogenetically diverse and limited distribution using monoclonal antibodies. J. Reprod. Immunol., 6:141-150. 481. Sakai, Y., Koyama, Y.-I., Fujimoto, H., Nakamoto, T., and Yamashina, S. (1986): Immunocytochemical study of fibrous sheath formation in mouse spermiogenesis using a monoclonal antibody. Anat. Rec., 215:119-126. 482. Bradfield, J. R. G. (1955): Fibre patterns in animal flagella and cilia. Symp. Soc. Exp. Biol., 9:306-334. 483. Olson, G. E., Hamilton, D. W., and Fawcett, D. W. (1976): Isolation and characterization of the fibrous sheath of rat epididymal sper¬ matozoa. Biol. Reprod., 14:517-530. 484. O’Brien, D. O., and Bellve, A. R. (1980): Protein constituents of the mouse spermatozoon. II. Temporal synthesis during spermato¬ genesis. Dev. Biol., 75:405-418. 485. Jones, R., Brown, C. R., Cran, D. G., and Gaunt, S. J. (1983): Surface and internal antigens of rat spermatozoa distinguished using monoclonal antibodies. Gamete Res., 8:255-265. 486. Fenderson, B. A., Toshimori, K., Muller, C. H., Lane, T. F., and Eddy, E. M. (1987): Identification of a protein in the fibrous sheath of the sperm flagellum. Biol. Reprod. (submitted for pub¬ lication). 487. Hunter, D. G., and Kretzer, F. L. (1986): Abnormal axonemes in sperm of fertile men. Arch. Androl., 16:1-12. 488. Pelfrey, J. J., Overstreet, J. W., and Lewis, E. L. (1982): Abnor¬ malities of sperm morphology in cases of persistent infertility after vasectomy reversal. Fertil. Steril., 33:160-166. 489. Escalier, D., and David, G. (1984): Pathology of the cytoskeleton of the human sperm flagellum: Axonemal and peri-axonemal anom¬ alies. Biol. Cell, 50:37-52. 490. Aitken, R. J., Ross, A., and Lees, M. M. (1983): Analysis of sperm function in Kartagener’s syndrome. Fertil. Steril., 40:696-698. 491. Smith, D., Oura, C., and Zamboni, L. (1970): Fertilizing ability of structurally abnormal spermatozoa. Nature, 227:79-80. 492. Baccetti, B., Burrini, A. G., and Pallini, V. (1980): Spermatozoa and cilia lacking axoneme in an infertile man. Andrologia, 12:525-532. 493. Ross, A., Christie, S., and Edmond, P. (1973): Ultrastructural tail defects in the spermatozoa from two men attending a subfertility clinic. J. Reprod. Fertil., 32:243-251.

/

67

494. Williamson, R. A., Koehler, J. K., and Smith, W. D. (1984): Ul¬ trastructural sperm tail defects associated with sperm immotility. Fertil. Steril., 41:103-107. 495. Sauvalle, A., Le Bris, C., and Izard, J. (1983): Supernumerary microtubules and prolongation of the middle piece in two infertile patients. Int. J. Fertil., 28:173-176. 496. Afzelius, B. A. (1981): Genetical and ultrastructural aspects of the immotile-cilia syndrome. Am. J. Hum. Genet., 33:852-864. 497. Afzelius, B. A. (1976): A human syndrome caused by immotile cilia. Science, 193:317-319. 498. Afzelius, B. A., and Eliasson, R. (1979): Flagellar mutants in man: On the heterogeneity of the immotile-cilia syndrome. J. Ultrastruct. Res., 69:43-52. 499. Leestma, J. E., and Sepsenwol, S. (1980): Sperm tail axoneme alterations in the Wobbler mouse. J. Reprod. Fertil., 58:267-270. 500. Olds, P. J. (1971): Effect of the T locus on sperm ultrastructure in the house mouse. J. Anat., 109:31-37. 501. Bryan, J. H. D. (1977): Spermatogenesis revisited. IV. Abnormal spermiogenesis in mice homozygous for another male-sterility-in¬ ducing mutation, hpy (hydrocephalic-polydactyl). Cell Tissue Res., 180:187-201. 502. Serres, C., Feneux, D., and Jouannet, P. (1986): Abnormal distri¬ bution of the periaxonemal structures in a human sperm flagellar dyskinesia. Cell Motil. Cytoskel., 6:68-76. 503. Escalier, D., and Series, C. (1985): Aberrant distribution of the peri¬ axonemal structures in the human spermatozoon: Possible role of the axoneme in the spatial organization of the flagellar components. Biol. Cell, 53:239-250. 504. Cooper, T. G., and Hamilton, D. W. (1977): Observations on de¬ struction of spermatozoa in the cauda epididymides and proximal vas deferens of non-seasonal male animals. Am. J. Anat., 149:93—110. 505. Olson, G. E., and Linck, R. W. (1977): Observations of the structural components of flagellar axonemes and central pair microtubules from rat sperm. J. Ultrastruct. Res., 61:21-43. 506. Bennett, W. I., Gall, A. M. Southard, J. L., and Sidman, R. L. (1971): Abnormal spermiogenesis in quaking. A myelin-deficient mutant mouse. Biol. Reprod., 5:30-58. 507. Woolley, D. M. (1977): Evidence for twisted plane undulation in golden hamster sperm tails. J. Cell Biol., 67:159-170. 508. Yeung, C. H., and Woolley, D. M. (1984): Three-dimensional bend propagation in hamster sperm models and the direction of roll in free-swimming cells. Cell Motil., 4:215-226. 509. Woolley, D. M., and Osborn, I. W. (1984): Three-dimensional ge¬ ometry of motile hamster spermatozoa. J. Cell Sci., 67:159-170. 510. Mohri, H., and Yano, Y. (1982): Reactivation and microtubules sliding in rodent spermatozoa. Cell Motil. (Suppl.), 1:143-147. 511. Phillips, D. M., and Olson, G. E. (1975): Mammalian sperm mo¬ tility. Structure in relation to function. In: The Functional Anatomy of the Spermatozoon, edited by B. A. Afzelius, pp. 117-126. Pergamon Press, New York. 512. Lindemann, C. B. (1980): Requirements for motility in mammalian sperm. In: Testicular Development, Structure and Function, edited by A. Steinberger and E. Steinberger, pp. 473-479. Raven Press, New York. 513. Yeung, C. H., and Woolley, D. M. (1983): A study of bend formation in locally reactivated hamster sperm flagella. J. Muscle Res. Cell Motil., 4:625-645. 514. Rikmenspoel, R. (1984): Movements and active moments of bull sperm flagella as a function of temperature and viscosity. J. Exp. Biol., 108:205-230. 515. Gray, J. (1958): The movement of the spermatozoa of the bull. J. Exp. Biol., 35:96-108. 516. Serres, C., Feneux, D., Jouannet, P., and David, G. (1984): Influ¬ ence of the flagellar wave development and progagation on the human sperm movement in seminal plasma. Gamete Res., 9:183-195. 517. Feneux, D., Serres, C., and Jouannet, P. (1985): Sliding sperma¬ tozoa: A dyskinesia responsible for human infertility? Fertil. Steril., 44:508-511. 518. Turner, T. T., and Giles, R. D. (1982): A sperm motility inhibiting factor in the rat epididymis. Am. J. Physiol., 242:R199-R203. 519. Usselman, M. C., and Cone, R. A. (1983): Rat sperm are mechan¬ ically immobilized in the cauda epididymidis by “immobilin,” a high molecular weight glycoprotein. Biol. Reprod., 29:1241-1253. 520. Turner, T. T., and Reich, G. W. (1985): Cauda epididymal sperm motility: A comparison among five species. Biol. Reprod., 32:120-128.

68

/ Chapter 2

521. Carr, D. W., and Acott, T. S. (1984): Inhibition of bovine sper¬ matozoa by cauda epididymidal fluid: I. Studies of a sperm motility quiescence factor. Biol. Reprod., 30:913-925. 522. Hoskins, D. D., Stephens, D. T., and Hall, M. L. (1974): Cyclic adenosine 3',5'-monophosphate and protein kinase levels in devel¬ oping bovine spermatozoa. J. Reprod. Fertil., 37:131-133. 523. Amann, R. P., Hay, S. R., and Hammerstedt, R. H. (1982): Yield, characteristics, motility and cAMP content of sperm isolated from seven regions of ram epididymis. Biol. Reprod., 27:723-733. 524. Hoskins, D. D., Hall, M. L., and Munsterman, D. (1975): Induction of motility in immature bovine spermatozoa by cyclic AMP phos¬ phodiesterase inhibitors and seminal plasma. Biol. Reprod., 13:168176. 525. Brandt, H., Acott, T. S., Johnson, D. J., and Hoskins, D. D. (1978): Evidence for the epididymal origin of bovine sperm forward motility protein. Biol. Reprod., 19:830-835. 526. Acott, T. S.. Katz, D. F., and Hoskins, D. D. (1983): Movement characteristics of bovine epididymal spermatozoa: Effects of forward motility protein and epididymal maturation. Biol. Reprod., 29:389399. 527. Garbers, D. L. Tubb, D. J.. and Hyne, R. V. (1982): A requirement of bicarbonate for Ca2 +-induced elevation of cyclic AMP in guinea pig spermatozoa. J. Biol. Chem., 257:8980-8984. 528. Vijayaraghavan, S., Critchlow, L. M., and Hoskins, D. D. (1985): Evidence for a role for cellular alkalinization in the cyclic adenosine 3',5'-monophosphate-mediated initiation of motility in bovine caput spermatozoa. Biol. Reprod., 32:489-500. 529. Gibbons, I. R. (1981): Cilia and flagella of eukaryotes. J. Cell Biol., 91:107 s— 124s. 530. Lindemann, C. B. (1978): A cAMP-induced increase in the motility of demembranated bull sperm models. Cell, 13:9-18. 531. Mohri, H., and Yanagimachi, R. (1980): Characteristics of motor apparatus in testicular, epididymal and ejaculated spermatozoa. A study using demembranated sperm models. Exp. Cell Res., 127:191196. 532. White, I. G., and Voglmayr, J. K. (1986): ATP-induced reactivation of ram testicular, cauda epididymal, and ejaculated spermatozoa ex¬ tracted with Triton X-100. Biol. Reprod., 34:183-193. 533. Garbers, D. L., and Kopf, G. S. (1980): The regulation of sper¬ matozoa by calcium and cyclic nucleotides. In: Advances in Cyclic Nucleotide Research, edited by P. Greengard and G. A. Robison, pp. 251-306. Raven Press, New York. 534. Hyne, R. V., and Garbers, D. L. (1979): Regulation of guinea pig sperm adenylate cyclase by calcium. Biol. Reprod., 21:1135-1142. 535. Wasco, W. M., and Orr, G. A. (1984): Function of calmodulin in mammalian sperm: Presence of a calmodulin-dependent cyclic nu¬ cleotide phosphodiesterase associated with demembranated rat caudal epididymal sperm. Biochem. Biophys. Res. Commun., 188:632-642. 536. Horowitz, J. A., Toeg, H., and Orr, G. A. (1984): Characterization and localization of cAMP-dependent protein kinases in rat caudal epididymal sperm. J. Biol. Chem., 259:832-838. 537. Atherton, R. W. Khatoon, S., Schoff, P. K., and Haley, B. E. (1985): A study of rat epididymal sperm adenosine 3',5'-monophosphate-dependent protein kinases: Maturation differences and cel¬ lular location. Biol. Reprod., 32:155-171. 538. Noland, T. D., Corbin, J. D., and Garbers, D. L. (1986): Cyclic AMP-dependent protein kinase isozymes of bovine epididymal sper¬ matozoa: Evidence against the existence of an ectokinase. Biol. Re¬ prod., 34:681-689. 539. Tang, F. Y., and Hoskins, D. D. (1976): Phosphoprotein phosphatase of bovine epididymal spermatozoa. Biochem. Biophys. Res. Com¬ mun., 62:328-335. 540. Tash, J. S. (1976): Investigations on adenosine 3',5'-monophosphate phosphodiesterase in ram semen and initial characterization of a sperm-specific enzyme. J. Reprod. Fertil., 47:63-67. 541. Stephens, D. T., Wang, J. L., and Hoskins, D. D. (1979): The cyclic AMP phosphodiesterase of bovine spermatozoa: Multiple forms, ki¬ netic properties, and changes during development. Biol. Reprod., 20:483-491. 542. Tash, J. S., and Means, A. R. (1982): Regulation of protein phos¬ phorylation and motility of sperm by cyclic adenosine monophos¬ phate and calcium. Biol. Reprod., 26:745-763. 543. Ono, T., Koide, Y., Arai, Y., and Yamashita, K. (1985): Estab¬ lishment of an efficient purification method and further characteri¬

zation of 32K calmodulin-binding protein in testis. J. Biochem., 98:1455-1461. 544. Okamura, N., and Sugita, Y. (1983): Activation of spermatozoan adenylate cyclase by a low molecular weight factor in porcine seminal plasma. J. Biol. Chem., 258:13056-13062. 545. Vijayaraghavan, S., and Hoskins, D. D. (1986): Regulation of bovine sperm motility and cyclic adenosine 3',5'-monophosphate by aden¬ osine and its analogues. Biol. Reprod., 34:468-477. 546. Hyne, R. V., and Lopata, A. (1982): Calcium and adenosine affect human sperm adenylate cyclase activity. Gamete Res., 6:81-89. 547. Brown, M. A., and Casillas, E. R. (1984): Bovine sperm adenylate cyclase inhibition by adenosine and adenosine analogs. J. Androl., 5:361-368. 548. Henry, D., Ferino, F., Tomova, S., Ferry, N., Stengel, D., and Hanoune, J. (1986): Inhibition of the catalytic subunit of ram sperm adenylate cyclase by adenosine. Biochem. Biophys. Res. Commun., 137:970-977. 549. Forte, L. R., Bylund, D. B., and Zahler, W. L. (1983): Forskolin does not activate sperm adenylate cyclase. Mol. Pharmacol., 24:4247. 550. Hildebrandt, J. D., Codina, J., Tash, J. S., Kirchick, H. J., Lipschultz, L., Sekura, R. D., and Bimbaumer, L. (1985): The mem¬ brane-bound spermatozoal adenyl cyclase system does not share cou“ pling characteristics with somatic cell adenyl cyclases. Endocrinology, 116:1357-1366. 551. Kopf, G. S., Woolkalis, M. J., and Gerton, G. L. (1986): Evidence for a guanine nucleotide-binding regulatory protein in invertebrate and mammalian sperm. Identification by islet-activating protein-cat¬ alyzed ADP-ribosylation and immunochemical methods. J. Biol. Chem., 261:7327-7331. 552. Ramarao, C., and Garbers, D. L. (1985): Receptor-mediated regu¬ lation of guanylate cyclase activity in spermatozoa. J. Biol. Chem., 260:8390-8396. 553. Krebs, E. G., and Beavo, J. A. (1979): Phosphorylation-dephosphylation of enzymes. Annu. Rev. Biochem.,48:923-956. 554. Chulavatnatol, M., Panyim, S., and Wititsuwannakul, D. (1982): Comparison of phosphorylated proteins in intact rat spermatozoa from caput and cauda epididymidis. Biol. Reprod., 26:197-207. 555. Brandt, H., and Hoskins, D. D. (1980): A cAMP-dependent phos¬ phorylated motility protein in bovine epididymal sperm. J. Biol. Chem., 255:982-987. 556. Tash, J. S., Kakar, S. S., and Means, A. R. (1984): Flagellar motility requires the cAMP-dependent phosphorylation of a heat-stable NP40-soluble 56 kd protein, axokinin. Cell, 38:551-559. 557. de Lamirande, E., and Gagnon, C. (1984): Origin of a motility inhibitor within the male reproductive tract. J. Androl., 5:269-276. 558. Freedman, M. F., and Kopf, G. S. (1985): Characterization of a seminal plasma-associated inhibitor of human seminal plasma protein kinase. Biol. Reprod., 32:322-332. 559. Pliego, J. F., Van-Arsdalen, K., and Kopf, G. S. (1986): Distribution of a seminal plasma-associated protein kinase inhibitor in normal, oligozoospermic, and vasectomized men. Biol. Reprod., 34:885893. 560. Byrd, W., Sodoloski, J. E., and Wolf, D. P. (1983): Analysis of calcium uptake during incubation of human spermatozoa. Biol. Re¬ prod. (Suppl.), 28:103. 561. Rufo, G. A., Singh, J. P., Babcock, D. F., and Lardy, H. A. (1982): Purification and characterization of a calcium transport inhibitor from bovine seminal plasma. J. Biol. Chem., 257:4627-4632. 562. Goh, P., and Hoskins, D. D. (1985): The involvement of methyl transfer reactions and S-adenosylhomocysteine in the regulation of bovine sperm motility. Gamete Res., 12:399-409. 563. Bouchard, P., Penningroth, S. M., Cheung, A., Gagnon, C., and Bardin, C. W. (1981): Erythro-9-[3-(2-hydroxynonyl)] adenine is an inhibitor of sperm motility that blocks dynein ATPase and protein carboxymethylase activities. Proc. Natl. Acad. Sci. USA, 78 10331036. 564. Gagnon, C., Sherins, R. J., Phillips, D. M., and Bardin, C. W. (1982): Deficiency of protein carboxymethylase in immotile sper¬ matozoa of infertile men. N. Engl. J. Med., 306:821-825. 565. Acott, T. S., and Carr, D. W. (1984): Inhibition of bovine sper¬ matozoa by cauda epididymal fluid: II. Interaction of pH and a qui¬ escence factor. Biol. Reprod., 30:926-935.

The Physiology of Reproduction, edited by E. Knobil and J. Neill el al. Raven Press, Ltd., New York © 1988.

CHAPTER

3

The Mammalian Ovum Paul M. Wassarman

The Mammalian Ovum: A Chronological Review, 70 Development of the Ovum, 70 Oogenesis: From Primordial Germ Cells to Eggs, 70 • Origin and Behavior of Primordial Germ Cells, 71 • From Primordial Germ Cells to Oogonia, 72 • From Oogonia to Nongrowing Oocytes, 72 • From Nongrow¬ ing to Fully Grown Oocytes, 72 • From Fully Grown Oocytes to Unfertilized Eggs, 73 • Summary, 73 Growth of the Oocyte: Structural Aspects, 73 Oocyte Growth: General Features, 73 • Nucleus (Ger¬ minal Vesicle) 73 • Nucleolus, 73 • Mitochondria, 74 • Golgi Complex, 75 • Cortical Granules, 76 • Zona Pellucida, 77 • Ribosomes, 77 • Cytoplasmic Lattices,

Oocyte Maturation: Regulatory Aspects, 82 Meiotic Maturation: General Considerations, 82 • Inhibitors of Meiotic Maturation, 82 • Role of cAMP, 84 • Role of Calcium, 85 • Role of Intercellular Com¬ munication, 85 • Role of Steroids, 86 • Role of Oocyte Maturation Inhibitor, 86 • Summary, 87 Growth of the Oocyte: Biochemical Aspects, 87 Oocyte Growth In Vitro, 87 • Overall Ribonucleic Acid Synthesis, 89 • Specific RNA Synthesis, 90 • RNA Synthesis: Summary, 92 • Overall Protein Synthesis, 92 • Specific Protein Synthesis, 93 • Protein Synthesis: Summary, 94 Oocyte Maturation: Biochemical Aspects, 94 Meiotic Maturation In Vitro, 94 • RNA Synthesis, 95 • Protein Synthesis Summary, 95 The Unfertilized Egg: Macromolecular Stores, 95 A Final Word, 96 Acknowledgments, 96 References, 96

78

Oocyte Maturation: Structural Aspects, 78 Meiotic Maturation: General Features, 78 • Acquisition of Meiotic Competence, 79 • Nuclear (Germinal Vesi¬ cle) Breakdown, 80 • Chromosome Condensation, 81 • Spindle Formation, 81 * Polar Body Emission, 82

The mammalian ovum, or egg, is the link between one generation and the next. In 1880, Nussbaum (187) recog¬ nized that “The fertilized egg, accordingly, divides into cells that constitute the individual and cells for maintenance of the species.” This concept was expanded insightfully by Wilson (285) in 1925 while explaining that the differences in form and function between sperm and egg are attributable to “a physiological division of labor between the gametes of the two sexes.” Continuing this theme, Wilson explains: “The ovum has to supply most of the material for the body of the embryo, and often also to provide for its protection and maintenance during development. For this service it prepares by extensive growth, accumulating a large amount of protoplasm, commonly laden with reserve food-matter (yolk or deutoplasm), and in many cases becoming sur¬ rounded by membranes or other protective envelopes. Dur¬ ing its early history, therefore, the ovum is characterized by predominance of the constructive or anabolic process of metabolism.” Herein lies the origin of the concept that there is not only a genetic but also a biochemical basis for the phrase embryogenesis begins during oogenesis. Today, there

is overwhelming experimental evidence that the zygote in¬ herits from the egg an extensive reserve of macromolecules and organelles that, to varying degrees, supports the nutri¬ tional, synthetic, energetic, and regulatory requirements of the early embryo (10,55,89). This is as true for mammals as it is for lower vertebrates and invertebrates, despite the obvious enormous differences in both reproductive and de¬ velopmental behavior of mammals and nonmammals. Here, many of the features of mammalian egg develop¬ ment are reviewed. Although emphasis is placed on mouse egg development in order to increase the clarity of the pre¬ sentation, most of the principles discussed can be applied to other mammals. Furthermore, in the interest of clarity and brevity, some important contributions are not cited and specific points of view are adopted on issues that may be the subject of some controversy among workers in the field. It is hoped that because of these and other shortcomings, the reader will be stimulated to refer to other detailed ac¬ counts of research on mammalian egg development (9,11,35,55,144,149,197,269,291,293), as well as tech¬ nical aspects of this research (87,88,144,217).

69

70

/ Chapter 3

THE MAMMALIAN OVUM: A CHRONOLOGICAL PERSPECTIVE Although the ovary was recognized as an anatomical en¬ tity by Herophilus of Alexandria in ca. 300 B.C. and de¬ scribed in some detail by Soranus of Ephesus in ca. 50 A.D., the mammalian ovum or egg was not identified until early in the 19th century A.D. by Karl Ernst von Baer. Earlier, De Graaf (ca. 1670) had recognized that eggs came from the ovary, but concluded incorrectly that the entire follicle (Graafian follicle) was an egg. This misconception was rectified somewhat by Cruickshank (ca. 1795) and oth¬ ers; however, it remained for von Baer (ca. 1825) to elu¬ cidate the exact anatomical relationship between egg and follicle in mammals (Fig. 1). Therefore, nearly 150 years separated identification of spermatozoa by Leeuwenhoek (ca. 1675) and eggs by von Baer. Waldeyer (ca. 1870) is credited with championing the concept that, in mammals, the sexually mature female pos¬ sesses a finite stock of oocytes that is drawn upon throughout her reproductive life. However, the alternative view, that generation of oocytes is a continual process throughout a female’s reproductive life (i.e., similar to spermatogenesis), although based on subjective interpretation of relevant ob¬ servations, prevailed during the first half of this century. Not until the early 1950s was this controversial issue finally put to rest and Waldeyer’s tenet accepted, due essentially to the work of Zuckerman and colleagues. Goette (ca. 1875) and Nussbaum (ca. 1880) were among the first to recognize that primordial germ cells, destined to give rise to oocytes and eggs, arise from undifferentiated cells located some distance away from, and appearing before the formation of, the genital ridges. Appreciation for the subsequent organogenesis of the mammalian ovary came about primarily as a result of the work of de Winiwarter and Sainmont in ca. 1910, and by the 1920s, a great deal

of cytological information about oocyte development was already available. In the 1920s it was recognized that the ovary is under functional control of the anterior hypophysis and, by the late 1920s and early 1930s, the relationship between oocyte and follicle development began to be ap¬ preciated as a result of work by Brambell, Parkes, Zuck¬ erman, Pincus, and colleagues. Particularly important work during this period, primarily from Pincus’s laboratory, in¬ volved comparison of mammalian egg behavior in vivo and in vitro. In the late 1930s the principal ovarian steroids were isolated and characterized, and by the early 1950s much of the endocrinological basis of pituitary-ovarian interaction affecting oogenesis and ovulation was appreciated. By the 1960s, in vitro culture of mammalian eggs and biochemical investigation of their metabolism became a reality as a result of work by Mintz, Biggers, McLaren, Epstein, Brinster, Graham, Piko, and others. * This brief history has been drawn from discussions found in refs. 8,187,219,241,285,292, and 294.

DEVELOPMENT OF THE OVUM Oogenesis: From Primordial Germ Cells to Eggs In mammals, oogenesis begins relatively early in fetal development and ends, months to years later, in the sexually mature adult (23,24,50-52,119,130,144,149,197,202, 224,242,292,294). Oogenesis begins with primordial germ cell formation and encompasses a series of cellular trans¬ formations, from primordial germ cells to oogonia (fetus), from oogonia to oocytes (fetus), and from oocytes to eggs (adult) (Fig. 2). This exquisitely orchestrated process results in a cell uniquely able to give rise to a new individual that expresses and maintains characteristics of the species. A description of oogenesis in mammals follows.

FIG. 1. Drawing of the mammalian egg within the Graafian follicle by von Baer, De Ovi Mammalium et Hominis Genesi (1827). Vesicula Graafiana (mediae magnitudinis) scrofae, decies aucta ad axin dissecta (IX). (1) Epithe¬ lium peritoneale; (2) tela formativa (stroma); (3) stratum externum (thecae); (4) stratum internum (thecae); (5) membrana granulosa (nuclei); (6) fluidum contentum; (7) discus proligerus (nuclei); (8) ovulum (nuclei); (x) stigma. (From ref. 241.)

The Mammalian Ovum

/

71

PRIMORDIAL GERM CELLS

o o

OOGONIA Mitosis

OOCYTES INITIATION OF MEIOSIS

0

leptotene zygotene

Recombination

pachytene diplotene (dictyate)

Fetus

MEIOTIC ARREST

Post partum

®

NON- GROWING Growth FULLY-GROWN

FIG. 2. Landmarks of oogenesis in the mouse. Progression from primor¬ dial germ cells to nongrowing oocytes during fetal development, as well as from nongrowing oocytes to fertilized eggs in sexually mature adults.

RESUMPTION OF MEIOSIS Ovulation Met a phase II

O

MEIOTIC ARREST

UNFERTILISED EGGS

-

RESUMPTION OF MEIOSIS

Activation

e

MEIOSIS COMPLETE

EMBRYOS

Origin and Behavior of Primordial Germ Cells Eggs originate from a small number of stem cells, the primordial germ cells, that have an extragonadal origin

(62,72,78,103,104,119,130,137,144,149,174,224). For¬ mation of these cells in presomite embryos signals the be¬ ginning of oogenesis. In 8-day mouse embryos (4 pairs of somites), as few as 15 and as many as 100 primordial germ cells are recognizable because of their size, distinctive mor¬ phology, and characteristic cytochemical staining proper¬ ties. These large cells (—12 |xm in diameter) are found in the yolk-sac endoderm and in that region of the allantois arising from the primitive streak. Several lines of evidence suggest that the embryonic rudiment of the allantois and caudal end of the primitive streak may be considered regions of primordial germ cell formation. In this context, it has been found that primitive ectoderm, taken from the caudal end of 7-to 7.5-day egg cylinders and cultured in vitro, differentiates into cells having properties characteristic of primordial germ cells. Subsequently, primordial germ cells migrate, first by passive transfer into the endodermal epi¬ thelium of the hindgut (170-350 primordial germ cells are found in or near the hindgut epithelium in 9-day embryos) and then by ameboid movement along the dorsal mesentery

of the genital ridges (present in 10- to 11-day embryos) found in the roof of the coelom; the site of gonad devel¬ opment. It appears likely that chemotactic mechanisms op¬ erate during migration of primordial germ cells from extraembryonic sites to the presumptive gonad. As a result of continuous mitotic activity, the number of primordial germ cells increases to approximately 5,000 in 11- to 12-day embryos and to more than 20,000 by the time genital ridges of 13- to 14-day embryos are fully colonized (Note: This represents a doubling of the number of primordial germ cells every 18 hr or so, between day 8 and day 14 of embryogenesis; it is estimated that they divide seven to eight times during the 4-day migration period). Primordial germ cells proliferate for only 2 to 3 days after migrating to the genital ridges; less than 1% of the germ cells exhibit an S phase in 16- to 17-day embryos. These primordial germ cells are the sole source of adult germ cells. It is noteworthy that the origin and migration of primordial germ cells to the genital ridges is the same in males and females; gonadal sex differentiation, to either testis or ovary, occurs in the 12- to 13-day embryo. In summary, the germ cell line originates before or during primitive streak formation (—7-day embryo), but precisely where, when, and how is still not clear. Mouse primordial

72

/

Chapter 3

germ cells are found sequentially in four different sites as embryogenesis proceeds: (i) First, they appear in the extraembryonic tissues of the yolk sac and allantois of 8- to 9-day embryos, (ii) Then they appear in the hindgut epi¬ thelium of 9- to 10-day embryos, (iii) Then they are found in the dorsal mesentery of the gut of 10- to 12-day embryos, (iv) Finally they are found in the developing gonads of 10.5to 11-day embryos. These primordial germ cells are the sole source of adult germ cells.

From Primordial Germ Cells to Oogonia Upon reaching the surface epithelium of the gonad, pri¬ mordial germ cells move into the cortex and, together with supporting epithelial cells, give rise to the cortical sex cords (119,129,149,293). The somatic components of the ovary arise from coelomic epithelium and mesenchyme of the dorsal body wall, and the mesonephros probably makes a contribution. In 13-day mouse embryos (52-60 pairs of somites), containing a differentiated ovary, migration of primordial germ cells is complete, with virtually all of the cells converted to actively dividing oogonia in the sex cords. Oogonia exhibit a characteristic morphology (including the presence of intercellular bridges connecting adjacent germ cells) and a high frequency of mitotic division.

From Oogonia to Nongrowing Oocytes As early as day 12 of embryogenesis, a few oogonia (~5%) enter the preleptotene, and then leptotene, stage of the first meiotic prophase (23,24,62,119,129,149,202,293). Once meiotic prophase commences, apparently there is no endocrine requirement for continued meiotic progression. Furthermore, since germ cells, either cultured in vitro or located at ectopic sites, enter and progress through meiosis, a gonadal environment is not required for the progress of meiosis (90,264). It is during preleptotene (interphase fol¬ lowing the last mitotic division of oogonia) that the final DNA replication takes place in preparation for meiosis. This synthetic activity signals transformation of oogonia into oo¬ cytes. It is possible that a factor originating from the rete ovarii, or simply contact with the rete ovarii, induces oog¬ onia to enter meiosis. In 14-day mouse embryos (61-62 pairs of somites), the germ cell population is about equally divided between oogonia and oocytes, and by day 17 (full quota of 65 pairs of somites) the ovary contains only oocytes at various stages of the first meiotic prophase (17,23,24, 26-30,118,135,249). Oocytes progress through leptotene in 3 to 6 hr and then take 12 to 40 hr to complete zygotene. During zygotene, homologous chromosomes pair and syn¬ apse to form what appear to be single chromosomes but are actually bivalents composed of four chromatids. In 16-day embryos nearly all oocytes are in pachytene of the first meiotic prophase, a stage that lasts about 60 hr and involves genetic crossing-over and recombination. Therefore, it takes

approximately 4 days to complete nuclear progression from leptotene through pachytene. By day 18 of embryogenesis, the first oocytes are seen in diplotene of the first meiotic prophase, with their chromosomes exhibiting chiasmata that result from crossing-over. By parturition, a majority of oo¬ cytes have entered late diplotene (“diffuse diplotene”), or the so-called dictyate stage, and by day 5 postpartum, nearly all oocytes have reached the dictyate stage, where they will remain until stimulated to resume meiosis at the time of ovulation. This pool of small (—12-15 pm in diameter), nongrowing oocytes is the sole source of unfertilized eggs in the sexually mature adult.

From Nongrowing to Fully Grown Oocytes Shortly after birth, the mouse ovary is populated with approximately 8,000 nongrowing oocytes arrested in meiosis and enclosed within several squamous follicular cells (23,24,52,149,202,293). Approximately 50% of these oo¬ cytes are lost during the first 2 weeks following birth; this is attributable, in large measure, to oocytes leaving the ovary through the surface epithelium. However, during this same period, more oocytes begin to grow (—5%) than at any other period in the lifetime of the mouse. Commencement of oocyte growth is apparently regulated within the ovary, with the number of oocytes entering the growth phase being a function of the size of the pool of nongrowing oocytes (153). The oocyte and its surrounding follicle grow coordinately, progressing through a series of definable morphological stages (198-203). In sexually mature mice, oocytes complete growth before formation of a follicular antrum; consequently, the vast majority of follicle growth occurs after the oocyte has stopped growing. Growth is continuous, ending in either ovulation of a matured oocyte (unfertilized egg) or degen¬ eration (atresia) of the oocyte and its follicle. Completion of oocyte growth in the mouse takes 2 to 3 weeks, a relatively short period of time in comparison to the months or years required for completion of oocyte growth in many nonmammalian species (9,11,24,89,149,200, 202,208,273,292,294). The oocyte grows from a diameter of about 12 pm (volume — 0.9 pi) to a final diameter of about 80 pm (volume —270 pi), not including the zona pellucida (discussed in the sections on structural and bio¬ chemical aspects of oocyte growth). Therefore, during its growth phase, while continually arrested in dictyate of the first meiotic prophase, the mouse oocyte undergoes a 300fold increase in volume and becomes one of the largest cells of the body. Each oocyte is contained within a cellular follicle (—17 pm in diameter) that grows concomitantly with the oocyte, from a single layer of a few flattened cells to three layers of cuboidal granulosa cells (~ 900 cells; ~125-pm-diameter follicle) by the time the oocyte has completed its growth (7,24,38,52,61,119,142,143,145,149,191,198203,273,292-294). The theca is first distinguishable, out-

The Mammalian Ovum side of and separated by a basement membrane from the granulosa cells, when the granulosa region is two cell-layers thick (—400 cells; — 100-p.m-diameter follicle). During a period of several days, while the oocyte remains a constant size, the follicular cells undergo rapid division, increasing to more than 50,000 cells and resulting in a Graafian follicle greater than 600 |xm in diameter. The follicle exhibits an incipient antrum when it is several layers thick (—6,000 cells; —250-|xm-diameter follicle) and, as the antrum ex¬ pands, the oocyte takes up an acentric position surrounded by two or more layers of granulosa cells. The innermost layer of granulosa cells becomes columnar in shape and constitutes the corona radiata; these cells form specialized intercellular junctions, called gap junctions, with the ool¬ emma.

From Fully Grown Oocytes to Unfertilized Eggs In sexually mature mice, fully grown oocytes in Graafian follicles resume meiosis and complete the first meiotic re¬ ductive division just prior to ovulation (discussed in the sections on structural, regulatory, and biochemical aspects of oocyte maturation). Resumption of meiosis can be me¬ diated by a hormonal stimulus in vivo or simply by the release of oocytes from their ovarian follicles into a suitable culture medium in vitro (23,24,110,256,257,281). Oocytes undergo nuclear progression from dictyate of the first meiotic prophase (four times the haploid DNA complement) to me¬ taphase II (two times the haploid DNA complement). They remain at this stage of meiosis in the oviduct, or in culture, until stimulated to complete meiosis when either fertilization or parthenogenetic activation occurs. Progression from the dictyate stage (oocyte) to metaphase II (egg) of meiosis is called meiotic maturation. Meiotic maturation is character¬ ized by dissolution of the oocyte’s nuclear (germinal vesicle) membrane, condensation of chromatin into distinct biva¬ lents, separation of homologous chromosomes, emission of the first polar body, and arrest of meiosis with chromosomes aligned on the metaphase II spindle (discussed in the section on structural aspects of oocyte maturation). These ovulated eggs complete meiosis, with separation of chromatids and emission of a second polar body (second reductive division), upon fertilization.

Summary It should be apparent that the process of oogenesis in mammals includes several noteworthy features, including: (a) extraembryonic and extragonadal origin of germ cells; (b) migration of germ cells to presumptive gonads; (c) sexual differentiation of germ cells into oogonia or spermatogonia; (d) cessation of mitosis (oogonia) and initiation of meiosis (oocytes); (e) prolonged cessation of meiosis in the dictyate stage; (f) oocyte growth; (g) reinitiation of meiosis (meiotic maturation; first meiotic reduction) and ovulation (eggs);

/

73

maturation; first meiotic reduction) and ovulation (eggs); and (h) completion of meiosis (second meiotic reduction) in response to fertilization. From the appearance of pri¬ mordial germ cells during fetal development until ovulation of unfertilized eggs in sexually mature adults, oogenesis represents one of the most highly specialized and regulated biological processes in mammals.

GROWTH OF THE OOCYTE: STRUCTURAL ASPECTS Oocyte Growth: General Features Throughout the reproductive life of mammals, their ova¬ ries contain pools of nongrowing and growing oocytes ar¬ rested in dictyate of the first meiotic prophase. Only fully grown oocytes resume meiosis and are ovulated during each reproductive (estrous) cycle. Recruitment of oocytes into the growing pool is apparently under control of pituitary gonadotropins, although oocyte growth, without follicle de¬ velopment, does occur in hypophysectomized animals and during culture in vitro in the absence of hormones. The volume of a mouse oocyte increases nearly 300-fold during its 2-3 week growth phase. This tremendous enlarge¬ ment of the cell is indicative of a period of intense metabolic activity (discussed in the section on biochemical aspects of oocyte growth) which, in turn, is reflected in marked changes in oocyte ultrastructure, including the appearance (bioge¬ nesis) of some novel organelles (2,12,37,149,190,248,252, 270,276,282,289,290). For example, cortical granules and the zona pellucida, both involved in regulating fertilization, first appear in oocytes during their growth phase. A de¬ scription of certain of the ultrastructural changes accom¬ panying oocyte growth follows. Nucleus (Germinal Vesicle) The nucleus, or germinal vesicle (GV), of growing mouse oocytes increases in diameter, from 9 to 10 |xm in small (—20 (im) oocytes to 20 to 22 |xm in fully grown (—80 |xm) oocytes (77,276). Consequently, growth of mouse oo¬ cytes results in a marked change in the ratio of cytoplasmic to nucleoplasmic volume, increasing from about 8 to 1 in small oocytes to about 64 to 1 in fully grown oocytes. Concomitant with nuclear enlargement, the nucleolus and extranucleolar bodies undergo progressive, characteristic ul¬ trastructural changes, while chromosomes remain as highly diffuse bivalents. Nucleolus The nucleus (GV) of growing mouse oocytes contains a single large nucleolus and, frequently, one or two smaller nucleoli. Throughout oocyte growth the nucleolus enlarges, increasing in diameter from 2 to 3 |xm in small (—20 |xm) oocytes to 9 to 10 (xm in fully grown (—80 |xm) oocytes

74 /

Chapter 3

(75,76,276) (Fig. 3). This enlargement is accompanied by progressive changes in nucleolar fine structure, indicative of a period of intense ribosomal-RNA synthesis (25,75,76,138,176,177,189,196). The nucleolus undergoes a transition during oocyte growth, from a diffuse, reticulated type of structure, composed primarily of a fibrillogranular network (small oocytes), to a dense, uniform mass, exclu¬ sively fibrillar in nature (fully grown oocytes).

Mitochondria Oocyte growth is accompanied not only by a substantial increase in the number of mitochondria present, but also by marked changes in mitochondrial ultrastructure (250,252,276,286,290) (Fig. 4). Small (—20 pm) oocytes contain elongated (—1.5 pm long) mitochondria with nu¬

FIG. 3. Transmission electron micrographs comparing nucleolar ultrastructure during growth of the mouse oocyte. Oocytes are isolated from 3- (A), 5- (B), 10- (C), and 14-day-old (D) animals and range in diameter from about 20 pm (3 days) to

merous transversely oriented cristae in the so-called ortho¬ dox configuration and, in most cases, contain a single vac¬ uole. Continued oocyte growth is accompanied by accumulation of round- and oval-shaped mitochondria, which are vacuolated and beginning to display columnar-shaped arched cristae. Throughout this growth period, mitochondria are closely associated with smooth endoplasmic reticulum and are present in increasing numbers. Many dumbbell¬ shaped mitochondria are found, indicative of extensive mi¬ tochondrial growth and division. Fully grown oocytes (—80 pm) contain round or oval (—0.5 pm diameter), highly vacuolated mitochondria (—105 per oocyte) (175,207) that have arched and concentrically arranged cristae. Conse¬ quently, the morphology of mitochondria in fully grown oocytes is radically different than that of mitochondria in nongrowing and small oocytes.

60 pm (14 days). Note the marked change from a sparse fibrillogranular (A) to a dense (D) nucleolus during oocyte qrowth’ (From ref. 276.)

The Mammalian Ovum

FIG. 4. Transmission electron micrographs comparing mito¬ chondrial ultrastructure during growth of the mouse oocyte. Oocytes are isolated from 5- (A), 3- (B), 5- (C), 10- (D-F), and 21 -day-old (G-l) animals and range in diameter from about 20 /urn (3 days) to 85 /u,m (21 days). Note the marked change

Golgi Complex As in the case of mitochondria, the Golgi complex under¬ goes dramatic ultrastructural changes during oocyte growth; these changes are indicative of increasing Golgi activity (252,276,290) (Fig. 5). In small (—20 pun) oocytes, Golgi membranes appear as flattened stacks of arched lamellae and are associated with few, if any, vacuoles or granules. During early stages of oocyte growth, Golgi membranes become more active, as evidenced by lamellae that are spaced further apart (swelling at termini of lamellae), by the ap¬ pearance of vacuoles, and by the proximity of numerous

/ 75

from elongated mitochondria with transverse cristae (“orthodox configuration”) (A-C) to round or oval mitochondria with con¬ centric cristae (“unorthodox configuration”) (D-l) during oocyte growth. (From ref. 276.)

lipid vesicles. In the middle to late stages of oocyte growth, Golgi membranes exhibit increased numbers of very swol¬ len, stacked lamellae that are associated with numerous vacuoles, granules, coated vesicles, and lipid vesicles. These changes are consistent with increased participation of the Golgi in processing and concentration of secretory products (e.g., zona pellucida glycoproteins) and cortical granule formation during oocyte growth. It is noteworthy that, con¬ comitant with the conversion of fully grown oocytes to fertilized eggs, there is a dramatic decrease in the amount of recognizable Golgi membrane and an increase in the number of small membrane vesicles.

76 /

Chapter 3

FIG. 5. Transmission electron micrographs comparing Golgi complex ultrastructure during growth of the mouse oocyte. Oocytes isolated from 3- (A), 5- (B), 8- (C), 10- (D), 14(E,F), and 21-day-old (G-l) animals and ranging in diameter

Cortical Granules Cortical granules are small, spherical, membrane-bound organelles that are found in the cortical region of unfertilized eggs and thought to resemble lysosomes (133,230). These granules fuse with the oolema at fertilization and, by re¬ leasing their contents (including proteinases) into the perivitelline space, alter functional properties of the zona pellucida (secondary block to polyspermy). Mouse eggs contain approximately 4,500 cortical granules (ranging in diameter from 200 to 600 |xm) within about 2 |xm of the plasma membrane (188). Cortical granules first appear during oo¬

from about 20 jam (3 days) to 85 ju.m (21 days). Note the marked change from flattened stacks of parallel lamellae (A-D) to swollen, highly vacuolated, granular lamellae (E-l) during oocyte growth. (From ref. 276.)

cyte growth, associated with an expanding Golgi that has moved to the subcortical region of growing oocytes. Al¬ though it is clear that cortical granules are derived from Golgi, certain evidence suggests that there is a contribution from multivesicular bodies as well as from granular endo¬ plasmic reticulum. Morphological studies suggest that the cortical-granule population is heterogeneous with respect to contents, with some granules even containing ordered crys¬ talline arrays. It is unclear whether this heterogeneity of cortical-granule contents reflects functional differences or simply different extents of granule maturation. In any case, it is clear that oocyte growth is accompanied by formation

The Mammalian Ovum and accumulation of increasing numbers of cortical granules in anticipation of ovulation and fertilization.

Zona Pellucida The zona pellucida, a relatively thick extracellular coat that surrounds all mammalian eggs, appears during oocyte growth, increasing in width as oocytes increase in diameter (40,55,73,102,136,150,273,276) (Fig. 6). In mice, the zona pellucida of fully grown oocytes is about 7 p,m thick, con¬ tains about 3 ng of protein, and is permeable to both large macromolecules and small viruses. The protein components of zonae pellucidae are synthesized and secreted by growing oocytes, representing a major metabolic activity of the oo¬ cyte during this period (discussed in the section on bio¬ chemical aspects of oocyte growth). Appearance of zona pellucida material in the perivitelline space correlates with initiation of oocyte growth; nongrowing oocytes do not have a zona pellucida. In early stages of oocyte growth, zona pellucida material appears as patches of fine filaments be¬ tween the oocyte and follicle cells (Fig. 7). These filaments are of uniform width, can be several microns in length, and exhibit a structural periodicity (132). As growth continues, the zona pellucida becomes a denser and thicker meshwork of interconnected filaments completely surrounding the oo¬ cyte and largely separating it from follicle cells. However, contact between the oocyte and the innermost layer of fol¬ licle cells continues via junctional complexes formed be¬ tween oocyte microvilli and follicle cell extensions that pen¬

/

77

etrate the zona pellucida (discussed in the section on the role of intercellular communication). As in the case of cor¬ tical granules, the zona pellucida is laid down during oocyte growth in anticipation of its roles during and after fertiliz¬ ation (11). The zona pellucida contains sperm receptors that mediate sperm-egg interaction as a prelude to fertilization; it also participates in a secondary block to polyspermy fol¬ lowing fertilization of eggs (41,42,115,116).

Ribosomes Ribosomal RNA accumulates through much of mouse oocyte growth (discussed in the section on biochemical as¬ pects of oocyte growth), consistent with changes in nu¬ cleolar morphology during this period. It is estimated that the number of ribosomes present in the cytoplasm increases three- to fourfold as the oocyte diameter increases from 20 to 65 pm (122). In view of the enormous change in oocyte volume during this period of growth, this suggests that ri¬ bosome density (i.e., number of ribosomes per square mi¬ cron of cytoplasm) decreases as much as 10-fold. The num¬ ber of ribosomes present in polysomes also increases severalfold during oocyte growth, with the number of single polysomes, which are abundant in small oocytes (>70% of total), decreasing dramatically ( 1A Rb 1 Aid

4

4

4

A

9

8

10

11

12

13

Rb6Bnr

4

4

1 1

14

15

16

17

18

4

Ctf

1

i i

!

4

uu T6Ca

19 Rb163H

Rb6Bnr Rb3Bnr

_

4

f

1 1 T13H

1 1

;

T70H

T26H

1 1 1 1 1 ir> Rtf

T11H

;

pa

T9H

P

T18H

.

m

1 |

; •

T9H

hr

‘

T50H

*

T18H

6

tK

T70H T264Ca

ht

Si '

T50H e

•

pe

Col

1 1 1

JT30H

,t26h

T138Ca

9r

1 i

ITISn

sa

1 1

' T13H bf

;

d

Jt 17Hse

mi l b

T30H

T11H

t | T26UCa ,

1

bg

T138Ca

ija-l *

In

‘

1 1

1 TISn

1 1

FIG. 8. Diagram of mouse autosomes showing regions re¬ sulting in abnormal development (broken lines) or normal de¬ velopment (solid lines)', in some regions, evidence is incon¬

clusive (dotted lines): (A) maternal duplication with paternal deficiency; (B, p. 206) paternal duplication with maternal de¬ ficiency. (From ref. 308.)

206

/ Chapter 6

1

2

3

4

5

__ Rb2Bnr

RbIBnr

4

4

6

7

D. Rb2Bnr Rb3Bnr RblAld

4

4

4

8

9

Rb5Bnr

Rb6Bnr

10

11

12

13

14

RbRBnr

Rb6Bnr

aw

15 uw T6Ca

1 fz

i i

:

i i

T26H

T13H

17

18

19 Rbl63H

.

t J

; •

i i

• ;

i i

4

*

i

1 1

T70H

16

Rb3B^r

.

jT138Ca

bg

vt

fi

1 hr

1

T9H

T30H

.

T18H Rw

T264Ca ua-1

In

T50H

T17H

T11H

d se

T18H 8

9r

T6Ca

tk mi

pa

T70H bt Si

T13H bf

•

T50H

1

Ca

a T30H bp T26H T ISn

TISr.

B FIG. 8. See legend on previous page.

and Markert and Petters (290), who demonstrated the fea¬ sibility of pronuclear removal and subsequent diploidization. Nuclei of inner-cell-mass cells, but not of trophectoderm cells, reportedly supported development to term when transferred to enucleated zygotes (281). Subsequent work has indeed shown that transplantation of trophectoderm nu¬ clei to zygotes in which the maternal and paternal pronuclei remain intact interferes with preimplantation development, but that inner-cell-mass nuclei do not exert this effect (280). However, a carefully controlled attempt to confirm that inner-cell-mass nuclei by themselves support preimplantation development has failed (312), thus casting doubt on the previous claim by Illmensee and Hoppe (281). In the study by McGrath and Solter (312), only nuclei of 2-cell embryos supported development into morphologically normal blas¬ tocysts when transferred to enucleated zygotes; nuclei from the 8-cell stage and beyond led to abortive cleavage (313).

Robl et al. (314) obtained better development of transferred 8-cell nuclei of mouse embryos by combining them with 2cell cytoplasm. In this case, half of the embryos formed blastocysts (a small fraction of which developed to early postimplantation stages), but none lived beyond day 12 of gestation (314). Even this limited success of 8-cell-stage mouse nuclei in supporting development of 2-cell cytoplasm indicates the importance of nuclear and cytoplasmic syn¬ chrony. Recent experiments with sheep embryos indicate that this species is more amenable to development with donor nuclei from 8-cell embryos. When 8-cell sheep blastomeres were fused with anucleate zygote halves, several developed into full-term, healthy individuals, whereas em¬ bryos derived from donor 16-cell nuclei developed to normal fetuses (315). Although the data for a species comparison are sparse, there appears to be a correlation between the cell numbers at which nuclear totipotency is lost and bias-

Early Embryogenesis

tocyst formation occurs (vide supra). That is, transfer of late-cleavage-stage nuclei was most successful in the species with a high cell number at the time of blastocyst formation (i.e., the sheep). Whereas the previously described studies establish certain time limits for nuclear totipotency (at least in the mouse), they do not formally address the issue of whether or not genomic imprinting is stable throughout cleavage, because the inability of more advanced nuclei to support early de¬ velopment may be due to asynchrony between donor nucleus and host cytoplasm (or other more trivial reasons). A recent study by Surani et al. (316), however, addresses this point. Experimentally produced haploid gynogenetic or androgenetic embryos were used as a source of donor haploid nuclei between the 2- and 16-cell stages. These were then trans¬ ferred to haploid gynogenetic or androgenetic zygotes. Re constituted diploids developed normally to term if they con¬ tained both maternal and paternal genetic contributions, but not if they were digynic or diandric (successful androgenetic donors were at the 2-4-cell stages; gynogenetic donors were at the 2-8-cell stages). Although several questions still must be resolved in this approach, the results indicate that ge¬ nomic imprinting acquired during gametogenesis is stable at least through early cleavage. A final implication of imprinting during gametogenesis is that there must be a mechanism for stable differentiation of the maternal and paternal genetic contributions. X-chromosome inactivation during embryogenesis provides a model for such a process.

X-Chromosome Activity During Development One X chromosome of female placental mammals is in¬ activated during development, leading to dosage compen¬ sation of X-linked genes (317-320). The process of Xinactivation is a paradigm for gene regulation, because changes in X-chromosome chromatin are correlated with changes in the expression of genes on this chromosome. Furthermore, the activity of the second X chromosome in females changes in a regular, well-documented sequence throughout the life cycle. Finally, the pattern of X-chromosome inactivation in extraembryonic tissues reinforces the idea of imprinting dur¬ ing gametogenesis. The status of X-chromosome activity has been studied by cytogenetic techniques, ratios of autosomal and X-linked enzymes, and isozymic patterns of polymorphic X-linked genes. Cytogenetic approaches rely on the differential stain¬ ing of the inactive X chromosome in interphase nuclei or metaphase chromosomes or on its asynchronous replication. The dosage of X-linked gene products is generally deduced from the absolute amounts of one or more gene products [including glucose-6-phosphate dehydrogenase (G6PD), hypoxanthine-guanine phosphoribosyl transferase (HPRT), and phosphoglycerate kinase (PGK)] or from the ratio of these activities to autosomal gene products. Isozymic variants in

/

207

human and mouse G6PD and mouse HPRT and PGK have been used as qualitative indications of X-chromosome ac¬ tivity. By combining the information from these approaches, we now have a detailed picture of the temporal and spatial aspects of X-chromosome activity, although the mechanistic basis of these patterns remains obscure.

X-Chromosome Activity During Gametogenesis and Embryogenesis The activity ratios of X-linked enzymes in XX and XO mice indicate that both X chromosomes of normal females are active in the oocyte (321). This is confirmed by the isozyme patterns of G6PD in human ovaries early in ges¬ tation (322) and by the patterns of G6PD and HPRT in mice (323). Similar studies of germ cells isolated from various stages of M. caroli embryos indicate that onset of activity in the previously inactive X chromosome coincides with meiosis (323). By contrast, the X chromosome of spermatogenic cells appears to be expressed at a low level or not at all (324,325). Thus, X-chromosome reactivation oc¬ curs during early stages of meiosis in the female, and this activity persists throughout oocyte growth until the time of ovulation, when transcription ceases (175). Because of the relative inactivity of the X chromosome during spermatogenesis, it was important to determine whether both X chromosomes are expressed in the early embryo or only that inherited from the egg. Cytogenetic observations made at various stages of embryogenesis first revealed asyn¬ chronous X-chromosome replication at the blastocyst stage; before this stage, both chromosomes have the appearance of active chromatin (326,327). Quantitative assays of Xlinked gene products demonstrated a bimodal distribution when single embryos were assayed (328-331), presumably indicating the presence of embryos with one active X (males) and two active Xs (females). Epstein and associates accom¬ plished a direct comparison of male and female embryos by separating blastomeres at the 2-cell stage, characterizing the sex of one half of the embryo cytogenetically, and assaying small groups of the other halves of embryos for HPRT levels (241,332). Female embryos had twofold higher levels of HPRT on the average than males. Together, these results demonstrate that the female preimplantation mouse embryo has two active X chromosomes. This has been confirmed in studies of feral mice heterozygous for HPRT isozymes, showing that both X chromosomes are expressed at least from the 8-cell stage (319; V. M. Chapman and G. Johnson, unpublished observations). Therefore, the preimplantation embryo, like the egg, is characterized by activation of both X chromosomes. When does X-inactivation occur? Cytogenetic observa¬ tions suggest that inactivation begins at the late morula to early blastocyst stage (327) or the late blastocyst stage (326). However, evidence from injection chimera experiments us¬ ing single inner-cell-mass cells from genetically marked

208

/ Chapter 6

mouse blastocysts (3.5-4.5 d.g.) reveals that both X chro¬ mosomes are active in the inner cell mass at these donor stages (333). This implies that the cytogenetic results on blastocysts are derived only from the trophectoderm cells. In addition, biochemical evidence indicates that there are two active X chromosomes in the embryonic ectoderm until day 6 of gestation (330,331,334). However, not all inner¬ cell-mass derivatives remain active until this time. Cyto¬ genetic observations by Takagi et al. (335) showed that there is an early-replicating X chromosome in the visceral endoderm as early as 5.3 d.g. and that this pattern changes to late replication in the visceral endoderm between 6.0 and 6.5 d.g., and also in the extraembryonic ectoderm and ectoplacental cone. Although the significance of the shift from early to late replication is unknown, both states reflect in¬ activity of the allocyclic X (336). Thus, X-chromosome inactivation accompanies cellular differentiation, occurring first in trophectoderm and its derivatives, then in primitive endoderm and its derivatives, and finally in the primitive ectoderm and its derivatives, including the germ cells (337). Only in early embryos and oocytes are both X chromosomes of a female individual active (Fig.9). Perhaps the most intriguing observation about the pattern of X-inactivation in early mouse embryos is that inactivation in the extraembryonic tissue lineages is nonrandom, the paternal X chromosome (Xp) being preferentially inacti¬ vated. This was first seen at the cytogenetic level (335) and was confirmed by analyzing PGK isozyme patterns in het¬ erozygous female embryos (335,338-342). These studies demonstrate conclusively that the Xp is preferentially in¬ activated, whereas the Xm is active in the visceral and pari¬ etal endoderm, the ectoplacental cone, and the extraem¬ bryonic ectoderm. Of the extraembryonic tissues examined, only visceral yolk sac mesoderm had a mosaic pattern of random X-inactivation. This is another manifestation of her¬ itable imprinting of maternal and paternal chromosomes, presumably acquired during gametogenesis (319). In mar¬ supials, the paternal X chromosome is preferentially inac¬ tivated in somatic tissues (320). The similarity between the spatial pattern of paternal X-inactivation and the specific extraembryonic developmental retardation in digynics and parthenotes implies a mechanistic relationship at the level of imprinting.

Trophectoderm Xm(+) Xp(-)

Oogenesis Xm(+) Xp(+)

Cleavage

ICM Xm(+) Xp(+)

Spermatogenesis Xm(-)

Mechanisms of X-Inactivation The alternation of periods of X-chromosome activity and inactivity throughout the life cycle (Fig. 9) implies that there is a mechanism for controlling X-chromosome function. This process can be viewed either as inhibitory inactivation (343) or as stimulatory activation (319,344). Although these models differ in their implications, both involve stage-spe¬ cific synthesis of chromatin-modifying gene products and their subsequent dilution or degradation. It has not yet been possible to define these products and thus to discriminate between the two models. Cattanach (345) has described a region (Xce) modifying X-chromosome expression, but there is no evidence that it directly encodes the putative regulatory factors. Secondary modification of DNA structure, particularly by methylation, has also been considered likely to be in¬ volved in X-chromosome regulation. The methylation of cytosine in CpG sequences often correlates with decreased expression in the affected or adjacent genes (346,347). Studies of DNA methylation in X-linked genes, however, have suggested a secondary rather than a primary role in X-inactivation (348). This doubt about the role of methyl¬ ation in the initial events of X-inactivation is heightened by the observation that the first tissues to show X-inactivation are extensively hypomethylated. Initial work on rabbit blas¬ tocysts revealed that trophoblast was significantly hypomethylated in comparison with inner-cell-mass derivatives, but not with adult tissues (349). Further work has focused on the extraembryonic tissues of the mouse conceptus and has revealed extensive undermethylation in satellite and dis¬ persed repetitive DNA sequences in all the derivatives of trophectoderm and primitive endoderm, as compared with early inner-cell-mass derivatives or adult tissues (350-352). Recent work shows that unique DNA sequences are also undermethylated in these same extraembryonic tissues (353). The causes and consequences of hypomethylation in the trophectoderm and primitive endoderm lineages are still un¬ clear. However, the hypomethylation pattern provides in¬ formation about its role in X-chromosome inactivation. DNA from the inactive X chromosome of adult tissues functions poorly in DNA-mediated transformation experiments with cultured HPRT-deficient cells (354). By contrast, DNA from

Primitive

Definitive

Endoderm

Endoderm, Mesoderm

Xm(+) Xp(-)

Xm(+) Xp(-) or Xm(-) Xp(+)

Primitive Ectoderm Xm(+) Xp(+)

Ectoderm,

Germ Cells Xm(+) Xp(-) or Xm(-) Xp(+

FIG. 9. Hypothesis linking X-chromo¬ some differentiation to cellular differ¬ entiation. X-chromosome differentiation is shown as occurring in different cell lineages as they differentiate (m, ma¬ ternal; p, paternal; +, active; inac¬ tive). Tissues differentiating early, that is, trophectoderm and primary endo¬ derm, show preferential inactivation of the paternally inherited X chromosome, suggesting some imprinting mechanism for the paternal origin of the two X chro¬ mosomes. This mechanism is presum¬ ably nullified by the time of gastrulation, when inactivation is random. (Adapted from ref. 337.)

Early Embryogenesis / the inactive X chromosome of extraembryonic tissues func¬ tions as well as DNA from the active X chromosome (355). Similar results have recently been obtained with a trans¬ genic mouse line containing an a-fetoprotein minigene in¬ tegrated into the X chromosome. Unlike the endogenous Xlinked genes, the exogenous a-fetoprotein gene was ex¬ pressed in visceral endoderm even when it was paternally inherited, but it was not expressed on the inactive X chro¬ mosome in the fetal liver (356). These results may indicate a role for DNA methylation in stabilizing X-inactivation, because the inactivation is less profound in cells exhibiting low levels of methylation. As yet, there is no information about the nature of Xchromosome imprinting leading to nonrandom inactivation in the extraembryonic tissues, or about how this terminates before X-inactivation in the primitive ectoderm lineage. The change from early to late replication of the inactive X in the extraembryonic tissues is intriguing because it coincides with the beginning of random X-inactivation between 6.0 and 6.5 d.g. (335,336). This may imply that growth pro¬ gressively dilutes either the imprint itself or the embryonic gene products that recognize it, with extinction occurring at the time of intense postimplantation growth. What information serves as the basis for nonrandom in¬ activation? The extraembryonic tissue lineages are notably all derived from cells that occupy external positions in the late preimplantation embryo, when inactivation of their X chromosome occurs. The pattern of preferential paternal Xinactivation in somatic tissues of the marsupial (320) rein¬ forces this correlation because the unilaminar structure of its blastocyst results in all cells having an external position (65,66). X-inactivation in primitive ectoderm also coincides with epithelial differentiation and formation of the proamniotic cavity. The correlation between embryonic position, epithelial differentiation, DNA hypomethylation, and pre¬ cocious inactivation of the paternal X chromosome indicates there is an epigenetic source of critical information common to these processes, but it does not provide any hint about its content. The evidence for a contribution from inner cell mass to the polar trophectoderm during mouse blastocyst growth implies that this epigenetic information is present during a period of several cell cycles, rather than occurring as a singular event at a specific early time. The net result of these early events is a restriction of random X-inactivation to the innermost cells of the periimplantation embryo. These cells are also characterized by levels of DNA methylation similar to those in adult tissues. In light of recent evidence that preimplantation mouse em¬ bryos are also hypomethylated in repetitive DNA sequences (J. Sanford, J. Rossant, and V. M. Chapman, unpublished observations), it appears that de novo methylation may be¬ gin shortly after the divergence of the first two cell types that form in the preimplantation embryo. When random Xinactivation occurs in early postimplantation development, it results in stable, permanently heritable patterns of Xlinked gene activity, perhaps a secondary consequence of

209

DNA methylation. Another consequence of this capacity for de novo methylation is the modification of exogenous DNA sequences introduced into the preimplantation embryo (357). In summary, X-chromosome inactivation during early de¬ velopment first occurs in extraembryonic tissue lineages as they differentiate (337). The pattern of inactivation in these tissues is nonrandom, resulting in preferential Xp-inactivation, in contrast to the random inactivation occurring slightly later in the primitive ectoderm lineage. Finally, the tissuespecific pattern of DNA methylation suggests a secondary rather than a primary role for these modifications in the Xinactivation process.

Introduction of Exogenous Genes Probably no single genetic technique has generated more excitement for developmental biologists than the introduc¬ tion of exogenous genes into the early embryo. This ap¬ proach has flourished with the availability of recombinant DNA techniques for producing and analyzing DNA se¬ quences. Furthermore, the use of cloned DNA sequences to transform cultured cells (358) was a compelling incentive for studying the fate of these genes throughout the whole of development, when they could potentially be regulated during the growth and differentiation of the intact organism. However, the first attempt at genetically altering mammalian embryos by the introduction of foreign DNA used virus as the DNA source. Jaenisch microinjected SV40 DNA into the blastocoele cavity in mouse embryos and detected the sequences in the adult mice (359,360). Subsequent studies performed with cloned DNA focused on the pronuclearstage mouse embryo (40) and on the early Drosophila em¬ bryo (42). When the injected DNA integrates into the host embryo genome, it is subsequently transmitted as a Mendelian trait, and the offspring of these novel individuals become unique strains. The first such mammals were termed transgenic mice (40,361-364). Subsequent similar studies have resulted in transgenic rabbits, sheep, and pigs (365). This approach has been strikingly successful in generating informative integration events (366-372). Although the earliest transgenic mice produced by injec¬ tion of pronuclear DNA did not express the exogenous genes, Wagner and associates eventually obtained activity using the herpes simplex virus (HSV) thymidine kinase gene (363), and Brinster and associates also obtained expression with a mouse metallothionein-HSV thymidine kinase fusion gene (361). These results stimulated numerous subsequent stud¬ ies. The primary objectives of such studies have been to determine the requirements for efficient, tissue-specific expression and to localize the genetic determinants of this specificity. Successful studies of exogenous gene expression have provided numerous insights into the biochemical and physiological aspects of hematopoiesis, immune system de¬ velopment, tumorigenesis, and other subjects. As a con¬ sequence of recent results that show a high incidence of

210

/

Chapter 6

efficient expression, gene therapies involving hematopoietic stem cells in humans and gene augmentation in livestock appear to be realistic possibilities. There are other potential uses for transgenic animals, such as insertional mutagenesis, which provides access to developmentally significant genes {vide supra). The remainder of this section focuses on the requirements and consequences of exogenous gene expression in transgenic mice.

Transgenics Produced by Pronuclear Injection To date, more than 40 cloned gene sequences have been introduced into pronuclear-stage mouse embryos, with at least some degree of expression (Table 3). The major cat¬ egories of sequences used are intact genomic clones (with their native controlling elements) and cDNA or genomic clones with heterologous controlling elements. The control sequences have most often been the immediate, 5' (up-

TABLE 3. Transgenic mice: function of exogenous DNAs Exogenous gene

Expression

a-Fetoprotein (mouse) a-Globin (mouse) (3-Globin (human) * 3-Globin (human-mouse fusion) p-Globin (rabbit)

Collagen a2(l)-chloramphenicol acetyltransferase (fusion) aA-crystallin-chloramphenicol acetyltransferase £a (mouse) Elastase 1 (rat) Elastase-growth hormone (rathuman fusion) Growth hormone (human) Hepatitis B surface antigen Herpes simplex virus Immunoglobulin

k

(mouse)

Immunoglobulin k, p. Immunoglobulin p, lnsulin-SV40 T antigen (fusion) Metallothionein-growth hormone (mouse-human fusion) Metallothionein-growth hormone (mouse-rat fusion) Metallothionein-growth hormonereleasing factor (mouse-human fusion) Metallothionein-HPRT (mousehuman fusion) Metallothionein-thymidine kinase Metallothionein-somatostatin (mouse-rat fusion) Mouse mammary tumor virus-thymidine kinase (fusion) Mouse mammary tumor virus-myc Moloney murine leukemia virus Myosin light chain 2 (rat) Rous sarcoma virus-chloramphenicol acetyltransferase SV40 SLA (pig class 1 major histocompatibility gene) SV40 T antigen

SV40 and v-myc Transferrin (chicken)

References

Tissue-specific Not specific None Tissue-specific Tissue-specific Not determined Not specific Questionable Tissue-specific

376 401 402 382 403,404 362 405 364 379

Tissue-specific

378

Tissue-specific Tissue-specific Tissue-specific

391-393 374 380

Not expressed Not specific Not determined Not specific Not specific Tissue-specific Tissue-specific Tissue-specific Tissue-specific Not specific

406 400 40 363 386 387 389 388 398 384

Not specific

383

Not specific

407

Semispecific

408

Not specific Not specific Semispecific Semispecific Not specific Tissue-specific Not specific Not determined Tissue-specific Not specific Semispecific Tissue-specific Semispecific Semispecific

361 409 410 397 411,412 375 379 413 390 396 394 395 414 373

Abbreviations: HPRT, hypoxanthine guanine phosphoribosyl transferase; SV40, simian virus 40.

Early Embryogenesis / stream) regions adjacent to structural genes. This region includes the promoter and enhancer sequences that are thought to be necessary for tissue-specific expression, based on stud¬ ies in cultured cells (358). Thus, the major hypothesis being tested in studies with transgenic mice is whether or not the controlling elements used are both necessary and sufficient for normal expression. Several studies have demonstrated that specific expression can be obtained using genomic DNA. The earliest indication of tissue-specific expression was a study by McKnight et al. (373) demonstrating synthesis of chicken transferrin in the liver of transgenic mice. A subsequent study using the rat elastase I gene showed high levels of tissue-specific synthesis in the mouse pancreas (374). The rat myosin light chain 2 was expressed specifically in the muscles of two transgenic mice (375). Interestingly, the mice varied widely in their levels of specific synthesis, despite similar copy numbers of the exogenous genes. This theme was repeated in mice containing a modified a-fetoprotein minigene; expression was specific to the visceral yolk sac and the fetal liver and gut, but at highly varying levels (376). In studies by Brinster, Palmiter, and associates, the metallothionein promoter was used to obtain gene expression predominantly in the liver of transgenic mice. However, the correlation between levels of expression and number of integrated cop¬ ies was poor (361,377). The major conclusions arising from these and other re¬ lated studies (Table 3) are that genomic sequences located adjacent (generally 5') to the structural genes studied are indeed necessary and sufficient in many cases for tissuespecific gene expression. Other factors are clearly involved, however, in the specific level of transcription obtained in each case. One possible cause for variable expression be¬ tween mice is the effect of the local chromatin environment at the site of integration. Another is the number of gene copies integrated, which varies from 1 to about 100, though there is generally little correlation between number of copies and total transcriptional activity. This creates a problem in the quantitative analysis of gene expression in transgenic mice, because it is therefore impossible to determine if transgenic individuals are synthesizing normal amounts of product per active exogenous gene, as compared with the native gene. Recent studies have demonstrated the effectiveness of using reporter or indicator genes encoding distinctive prod¬ ucts to study tissue-specific expression. Fusion genes con¬ taining the bacterial structural gene for chloramphenicol acetyltransferase and either the mouse aA-crystallin pro¬ moter or the a2(I) collagen promoter have led to tissue- and temporal-specific expression (378,379). A detailed analysis of control by rat elastase promoter using elastase-human growth hormone fusion genes indicated that sequences be¬ tween 80 and 200 base pairs “upstream” of the 5' end of the structural gene were responsible for the transcriptional regulation conferred by this enhancer (380). This was the first demonstration of a mammalian gene regulatory se¬

211

quence by the criteria of tissue specificity in transgenic animals. Reporter sequences are also valuable in the analysis of gene expression during embryogenesis. They are particu¬ larly useful when the expression of the structural gene is detectable in the intact embryo by virtue of a histochemical product. The use of histochemically detectable gene prod¬ ucts has been a powerful asset in studies of mouse chimeras (26). The expression of metallothionein promoter-regulated Escherichia coli (3-galactosidase, can be detected in single embryos after microinjection (M. Stevens, J. J. Meneses, and R. A. Pedersen, unpublished observations). Similarly, tissue-specific expression of a y-crystallin/(3-galactosidase fusion gene has recently been observed (381). These ob¬ servations indicate that structural genes, including some derived from prokaryotes, can be fused with mammalian controlling elements and yield tissue-specific expression. Although these indicator genes with bacterial structural components were expressed in an apparently normal manner in the transgenic mice described earlier, inclusion of bac¬ terial cloning vector sequences (e.g., pBR322) in the DNA injected into zygote pronuclei resulted in poor or erratic expression. Conversely, expression of human (3-globin genes occurred in a tissue- and temporal-specific way if vector sequences were reduced or eliminated entirely (382). Sim¬ ilar observations have been made with the a-fetoprotein gene (376) and with growth hormone genes (368). It is not yet clear what sequences of the bacterial plasmid confer inactivity or what mechanisms are involved in the silencing process. Although a comprehensive review of the information gained through injection of pronuclear DNA is beyond the scope of this chapter (371,372), several accomplishments are not¬ able because they illustrate the power of this approach in generating novel basic information and possible future prac¬ tical applications. The dramatic growth of mice carrying metallothionein-growth hormone fusion genes demonstrates the high levels of gene product that can be produced from exogenous genes and their impact on the individual’s phys¬ iology (383,384). The induction of growth in dwarf mice homozygous for the little mutation (deficient for growth hormone) demonstrates the potential for correcting genetic deficiencies with exogenous DNA (385). Several recent studies have demonstrated the enormous utility of transgenic mice for understanding immune system development. When a functional k light chain was intro¬ duced, the rearrangement of endogenous k genes was blocked (386,387). Similar results were obtained by injecting rear¬ ranged immunoglobulin p heavy-chain gene (388,389). Other studies have demonstrated the expression of pig major his¬ tocompatibility antigen genes in transgenic mice (390) and of exogenous Ea genes. In the latter case, a new immune response resulted from expression of the exogenous gene (391-393). These results should have profound implications for future work in immunology. A third example is the analysis of oncogene function in

212

/ Chapter 6

transgenic mice. Expression of the SV40 early-region genes led to tumors of the choroid plexus (394). Tumorigenesis could be attributed to the large-T antigen, although the tissue specificity resided in the SV40 enhancer (395,396). Trans¬ genic mice with the mouse mammary tumor virus promoter driving the c-myc proto-oncogene developed mammary ad¬ enocarcinomas (397), and mice with the SV40 large-T an¬ tigen under the control of the rat insulin promoter developed pancreatic tumors in the islets of Langerhans (398). Finally, transgenic mice expressing genes from human hepatitis B virus have been produced by two groups (399,400) and may be useful as an animal model for this disease. These examples, though by no means exhaustive (Table 3), confirm the tremendous advantages of generating novel individuals and strains that express defined genetic elements in a tissue-specific manner.

Trans genics Produced by Virus Infection Although virus DNA was used in the first studies dem¬ onstrating that exogenous genes could be integrated into the germ line of mice at embryonic stages (359), only recently have viruses become efficient vectors for producing trans¬ genic animals (371). The primary DNA viruses used for this purpose are the closely related simian virus 40 (SV40) and polyoma virus. These differ in that mouse cells are not permissive for productive infection by SV40, but are for polyoma infection. Recent work shows that preimplantation mouse embryos can support the replication of both viruses (415) and that plasmids containing the polyoma origin of replication can persist without integration into the host chro¬ mosomes of transgenic mice (416). The other virus now used widely in transgenic studies is the RNA tumor virus, Moloney murine leukemia virus (MuLV) (417). During their life cycle, the RNA viruses are transformed into a circular DNA that is integrated into the host cell chromosome. The integrated retroviral genome is not expressed in the preim¬ plantation embryo (418), and, moreover, it becomes irre¬ versibly inactivated in most cases, presumably by de novo DNA methylation (357,419). An equally persistent block to expression occurs for SV40 virus after infection of early embryonic cells (420). A fraction of mice derived from MuLV-infected embryos become viremic as adults, presum¬ ably because the position of integration is close to developmentally regulated genes (421). In recent work (422,423) it has been shown that genes regulated by internal promoters, rather than by the promoters of the MuLV long terminal repeats, can be expressed efficiently in embryonal car¬ cinoma cells, which are closely analogous to embryos (371, 424-426). This, together with the demonstration that MuLV can be used to efficiently transform preimplantation mouse embryos, offers an alternative to pronuclear-DNA injec¬ tion for introducing exogenous genes in a functional state (427).

Transgenics Produced by Transforming Embryonic Stem Cells Another means for introducing exogenous genes into the mouse germ line combines viral technology with cell culture and experimental chimerism. This approach is based on the recent ability to culture euploid pluripotent cells (embryonic stem cells) directly from embryos, thereby circumventing the long growth period previously required to adapt em¬ bryonal carcinoma cells from tumors to culture (426). The main advantage in using the embryonic stem cells is their euploidy and the attendant high frequency with which they colonize the germ line in chimeras produced by blastocyst injection (428). Using this approach, Stewart et al. (424) have recently introduced genes conferring neomycin resis¬ tance into mice. Moreover, it has recently proved to be possible to obtain germ line integration by means of em¬ bryonic stem cell transformation, selection, and transfer to host blastocysts (429,430). These efforts have benefited markedly from the availability of dominant selectable mark¬ ers, such as the bacterial neomycin-resistance gene and guanine phosphoribosyltransferase gene (431,432). These attempts to transform mouse embryos share technologies and goals with current efforts at gene therapy. The benefits of transgenic animals include elucidating the mechanisms of tissue-specific gene regulation, optimizing procedures for introducing exogenous genes into cells, and determining which promoters are effective in a wide variety of target tissues. This information is the necessary background for clinically effective gene therapy to correct biochemical de¬ fects in humans.

BIOCHEMICAL ASPECTS This section emphasizes the lineage-specific biochemical differentiation of the cell types that form in the early mam¬ malian embryo. The reason for this emphasis is that celllineage-related products are likely to provide clues to mech¬ anisms of differentiation. In addition, the general aspects of metabolic activities in mammalian embryos have been known for some time or have been adequately reviewed elsewhere (8,433). Because of their potential importance as probes for cell differentiation, many of the genes for prod¬ ucts described here have been cloned from one or more mammalian species. Molecular cloning goes hand in hand with analysis of early mammalian embryos, because the use of molecular probes confers such sensitivity and specificity.

Secreted Products The serum glycoprotein, a-fetoprotein (AFP), is probably the most thoroughly studied lineage-specific product in mammalian embryos. AFP is produced from early postim¬ plantation stages specifically by the visceral endoderm of

Early Embryogenesis / rat and mouse embryos (434-436). AFP accumulates to high levels in the amniotic fluid, where it appears to function as an embryonic counterpart to albumin, to which it is evolutionarily related and genetically linked (437). The halflife of AFP mRNA is relatively long (30 hr) in the mouse visceral yolk sac (438). There is evidence for independent regulation of AFP and albumin genes in mouse and rat development (439,440). Evidence from studies of AFPtransgenic mice shows that the DNA sequences necessary for temporal- and tissue-specific expression reside in the immediately adjacent 5' sequences (441,442). Several other serum proteins are also synthesized and secreted specifically by the visceral yolk sac. These include transferrin, metallothionein, a]-antitrypsin, and apolipoproteins (443-448). The similarity of products of the rodent visceral yolk sac and fetal liver has prompted the observation that these two organs, albeit derived from independent cell lineages, have homologous functions at early and late stages of intrauterine development, respectively (448).

Extracellular Matrix The substrates surrounding cells in an intact embryo have a profound impact on their morphology, behavior, and gene expression (449). Mouse embryos begin to synthesize fibronectin at the blastocyst stage, when it accumulates on the basal surfaces of primitive endoderm cells and near the junction of parietal endoderm and trophectoderm (450,451). At later stages, fibronectin is present in the basement mem¬ brane of visceral endoderm, trophoblast, and mesoderm. In addition to fibronectin, laminin and type IV collagen are major components of basement membranes in the early mouse embryo (452,453). Although one component of laminin is synthesized in unfertilized eggs, intercellular deposits do not accumulate until the 8- to 16-cell stage (454,455). Type III collagen deposits begin to accumulate at the 16-cell stage (456), but type I collagen does not begin to be synthesized until day 8 of gestation, when it accumulates in several fetal and extraembryonic tissues (453). An essential role for type I collagen in development is evident from the death at 12 d.g. of transgenic mice homozygous for a retrovirus inser¬ tion into this gene (457). The most prominent basement membrane in the early rodent embryo is Reichert’s mem¬ brane, consisting mainly of laminin, type IV collagen, and chondroitin sulfate proteoglycans; it is apparently synthe¬ sized by parietal endoderm cells (458,459). The availability of DNA probes to the mRNAs of most of these basement membrane components should facilitate the analysis of the precise location and time of their synthesis (448).

Cell Surface Molecules The cell surface differentiation of mammalian embryos has been the subject of extensive work (460-466). The

213

purpose of this section is to provide an overview of recent work.

Cell Adhesion Molecules The compacting morula in placental mammals is unique in its precocious cell adhesiveness and junctional differ¬ entiation (125,464). The fact that cleavage-stage embryos of marsupials and embryos in other vertebrate classes do not undergo compaction implies that this process may be integrally involved in the development of trophectoderm. Indeed, removing calcium from the culture medium prevents or reverses compaction of morula-stage mouse embryos and thwarts trophectoderm differentiation (467). Similar results can be obtained by treating early-cleavage-stage embryos with cytochalasin, which dissociates microfilaments, or with certain antibodies to embryonal carcinoma cells (125,465). The common denominator of this disruption of compaction appears to be cell adhesion molecules that are necessary for compaction. Described variously as uvomorulin (468), cadherin (469), and cell adhesion molecule 120/80 (470,471), cell adhesion molecules present on the preimplantation mouse embryo are calcium-dependent surface molecules resem¬ bling those present on liver cells (472). Because these cell adhesion molecules are present on mouse embryos even before the beginning of compaction, they cannot, however, be considered sufficient for compaction (473).

Lectins and Their Receptors The effects of some lectins on preimplantation develop¬ ment suggests a role for lectin-saccharide interactions in compaction and blastocyst formation. Agarose beads coated with concanavalin A (con-A), peanut agglutinin, or wheat germ agglutinin (WGA) induce spreading of 8-cell blastomeres of mouse embryos (474). Moreover, WGA induces compaction and cavitation in 2-cell mouse embryos (474). On the other hand, culturing 4-cell mouse embryos in either WGA or con-A prevents compaction and blastocyst for¬ mation and leads to vacuole formation in the blastomeres (476). As would be expected from these results, these three lectins bind to preimplantation mouse embryos (477). There are also receptors for the lectins Dolichos biflorus agglutinin and Ricinus communis agglutinin in early mouse embryos (478). Although the distribution of con-A on blastomeres is uniform before compaction, it becomes polarized in late8-cell embryos, and this persists at subsequent stages; this distribution is one component of the polarized phenotype (vide supra) that precedes the differentiation of trophecto¬ derm (125). Despite the presence of these lectin receptors, there is no compelling evidence that they have a role in development, because physiological concentrations of var¬ ious saccharides did not inhibit compaction of 8-cell mouse embryos (474).

214

/ Chapter 6

Oligosaccharide Antigens With the advent of monoclonal antibodies, several car¬ bohydrate antigens have been defined on the surfaces of embryonic cells. The best-studied of these, SSEA-1, is a lacto-series carbohydrate that appears on the 8-cell-stage mouse embryo, then segregates to the inner cell mass at the blastocyst stage (479). This antigen is expressed on the surfaces of visceral endoderm and embryonic ectoderm cells in postimplantation mouse embryos (480,481). Other lactoseries carbohydrates expressed on early mouse embryos are the human blood group precursor antigens I and i; I antigens are present at all preimplantation stages, and both antigens are present on parietal and visceral endoderm after day 6 of gestation (482-484). The globo-series carbohydrates rec¬ ognized by the monoclonal antibodies SSEA-3 and SSEA4 are present on mouse oocytes, preimplantation embryos, inner cell mass, and first on primitive and then on visceral endoderm at postimplantation stages (485-487); globoside is present on 2-cell- to morula-stage mouse embryos (488); Forssman antigen is present on late-morula- and blastocyststage embryos (489). The chemical structures of these car¬ bohydrate antigens have been well defined, so their bio¬ synthetic pathways can be inferred (465). Consistent with these findings, glycosyl transferases are present on the sur¬ faces of early embryo cells and embryonal carcinoma cells (490-492) and appear to have roles in compaction (493) and in the interconversion of the carbohydrate antigens (465).

Other Surface Molecules Alkaline phosphatase, Na+,K + -ATPase, and histocom¬ patibility antigens deserve special mention. Alkaline phos¬ phatase activity appears on the surfaces of 2-cell blastomeres and increases in intensity at the 8- to 16-cell stages, when it is concentrated on apposing cell surfaces; activity in blas¬ tocysts is restricted to the inner cell mass and subsequently to its derivatives (494,495). Activity on embryonic ecto¬ derm decreases gradually after gastrulation, but remains associated with the primordial germ cells, which are local¬ ized at the base of the allantois in 8-day mouse embryos (159). The activity can be used to localize the primordial germ cells during their migration through hindgut endoderm and mesenteries to the genital ridges during early stages of gonad formation (496,497). The accumulation of blastocoel fluid in the rabbit and mouse has been shown to depend on Na+ ,K + -ATPase ac¬ tivity, which becomes localized on apposed surfaces of blas¬ tomeres, beginning at the morula stage (498). Analysis of the blastocoel fluid composition in rabbit embryos cultured in normal and hyperosmotic medium indicated that Na+ and Cl" are transported, with presumed passive movement of water (499). The ionic movements have been attributed to Na + ,K + -ATPase, which can be identified on the juxtacoelic

surfaces of the cells of the rabbit blastocyst by using labeled ouabain as a specific marker (500,501). Mouse blastocysts exposed to ouabain in the culture medium (exposing only outer surfaces of trophectoderm cells) develop normally; however, embryos treated with cytochalasin to cause a col¬ lapse of the blastocoel failed to reexpand when they were treated with ouabain, presumably because the drug inhibited the juxtacoelic Na + ,K + -ATPase (502). An integral role for this enzyme in the polarization of mouse blastomeres at the morula stage has been hypothesized by Wiley (503), who has suggested that it is responsible for osmotic flows leading to a shift in the positions of cytoplasmic organelles and other processes leading to the formation of the blastocoel. Except for a transient appearance on the outer surfaces of trophectoderm cells at the blastocyst stage (504,505), major histocompatibility antigens are absent from mouse embryos until midgestation (506). However, p2-microglobulin synthesis begins at the 2-cell stage (507) and continues through preimplantation development (508). Because major histocompatibility antigen synthesis does not occur at early cleavage stages, the function of (32-microglobulin at these stages may be to anchor the minor histocompatibility an¬ tigens, which appear on cleavage-stage embryos and blas¬ tocysts (463,509).

Cytoskeletal Elements The cytoskeleton in early mammalian embryos has been studied using inhibitors of specific cytoskeletal elements and using antibodies or other probes to localize and quantitate the components of the cytoskeleton at various stages. Col¬ chicine, colcemid, and nocodazole have been used to study the role of tubulin in the mouse embryo. These inhibitors cause cleavage arrest and prevent blastocoel formation (510— 512). There have been conflicting reports that microtubule inhibitors induce decompaction of 8-cell embryos (513), but that nocodazole, also a microtubule-depolymerizing drug, accelerates compaction at the same stage (514). Cytochalasins, which disrupt microfilaments, cause cleavage arrest and inhibit compaction (510,513,515-517). Mouse em¬ bryos treated with cytochalasins during cleavage form ag¬ gregates of vacuolated cells at the time the blastocoel nor¬ mally forms; if the inhibitor is removed before vacuolation occurs, the embryos form trophectoderm vesicles that have polypeptide synthetic profiles similar to that of intact blas¬ tocysts (515). The compound effects of inhibitors that dissociate mi¬ crotubules or microfilaments on cell division and other mor¬ phogenetic events make it difficult to distinguish between effects induced because of cytoskeletal disruption and those resulting from cell cycle arrest. Thus, the inhibitor studies must be interpreted with caution. When the effects of such inhibitors were studied in interphase cells, surface (microvillar) polarization occurred in the absence of microtubules

Early Embryogenesis

or microfilaments, even though underlying cytoplasmic po¬ larization of the cytoskeleton and of endocytotic organelles was prevented (125,518,519). These results imply that or¬ ganization of the major cytoskeletal elements derives from the cell surface and cell-cell interactions. In addition to the microtubules and microfilaments that are evident in cleavage-stage mouse embryos (520-523), several cytoskeletal elements implicated in cytoskeletal-cell surface interaction are present. Their roles in the embryo are inferred from the functions of similar or identical pro¬ teins in erythrocytes, muscle cells, and nonmuscle cells (465). These include a-actinin (522) and spectrin (524), which are present at cleavage stages and could anchor actin filaments to the membrane, and myosin (525-527), which would be involved with actin in cell shape changes, mo¬ bility, and contractility. Another category of cytoskeletal elements contained in embryos is intermediate-filament proteins, which are ex¬ pressed in lineage-specific patterns (448,528). For example, desmins are found in muscle, glial fibrillar acidic protein is found in glial cells and astrocytes, and cytokeratins are found in epithelial cells. Cytokeratins designated as Endo A (TROMA-1) and Endo B first accumulate in the trophectoderm of mouse embryos at the blastocyst stage (529532). At later stages, both cytokeratins are found also in the primitive-endoderm lineage of mouse embryos (533). The mRNA encoding Endo A can be detected in trophectoderm, but not inner cell mass, at the blastocyst stage; at postimplantation stages, it appears in visceral endoderm and amnion (534). This pair of cytokeratins corresponds to the major pair of cytokeratins in adult mouse liver (Endo A = type II; Endo B = type I), as indicated by DNA cloning and sequencing (535). Although the role of cytokeratins in early mammalian development is not yet understood, their lineage specificity makes them useful as markers for study¬ ing differentiation and commitment at late preimplantation and early postimplantation stages. The potential for interactions between the cytoskeleton and the cell membrane makes the analysis of cytoskeletal elements in mammalian embryos an important area for fur¬ ther work. This is particularly evident from the effects of cell shape and extracellular matrix on proliferation and gene expression in other systems (536-539). Despite the advan¬ tages of a reductionist approach to development, the highly regulative nature of the mammalian embryo is a compelling argument for studying cell interactions in intact embryos whenever possible.

Regulatory Elements Several categories of lineage-related products may be im¬ plicated in the regulation of growth and differentiation. These include growth factors and their receptors, heat-shock pro¬ teins, and homeobox-containing genes.

/

215

Growth Factors and Their Receptors In addition to the secreted products previously described, early mammalian embryos produce and secrete growth fac¬ tors; in some cases, they also possess the receptors for these same factors (448,540). Transforming growth factor (TGF) a and (3 are epidermal-growth-factor-like polypeptides that stimulate anchorage-independent growth of untransformed TGF-a binds to the epidermal growth factor (EGF) receptor, but not to anti-EGF antibodies; using these criteria to assess the quantity of TGF-a reveals a high specific activity in the 7.5-d.g. mouse embryo (541) and in the human placenta (542). Somatomedins, or insulin-like growth factors (IGF), are produced by postimplantation mouse embryo tissues, specifically allantois and amnion (J. Heath, unpublished observations), and are present in human amniotic fluid (543). Trophoblast giant cells produce a prolactin-like protein (544,545), and prolactin is present in high concentrations in human amniotic fluid. Human chorionic gonadotropin is produced by the syncytiotrophoblast cells of the placenta and can be detected in the maternal serum as soon as 2 weeks after fertilization (546,547). The presence of growth factor receptors may be indicative of a developmental function of the relevant factor. Not re¬ stricted to a specific lineage, the EGF receptor appears at 11 to 12 d.g. on all cell types except parietal endoderm and is present at high levels on cells of the amnion and various fetal tissues (548,549). Because TGF-a can bind to the EGF receptor, it is plausible that the midgestation embryo can respond to TGF-a in an autocrine fashion. Alternatively, receptors may bind EGF produced in maternal tissues; no embryonic EGF source has been detected (550). The inner cell mass and primitive endoderm of the mouse embryo react with antibodies to the transferrin receptor, as do lab¬ yrinthine cells of the placenta, whereas trophoblast giant cells derived by in vitro outgrowth do not (E. D. Adamson, unpublished observation). The function of proto-oncogenes during development is of considerable interest because viral oncogenes promote the growth of cells. Moreover, a number of proto-oncogenes are either identical with or homologous with known growth factors or their receptors (551). The class I proto-oncogenes c-abl and c-src are expressed in a wide variety of tissues in mouse fetuses (448,552). This class is associated with tyrosine protein kinase activity. The class-I-related proto¬ oncogene c-erb-A is maximally expressed at day 14 of mouse gestation, but c-erb-B, which is homologous with EGF re¬ ceptor, is not present in high amounts in any tissue, contrary to expectation (552). Other class-I-related proto-oncogenes, c-fms [homologous with the CSF-1 receptor (553)] and cmos, are active in mouse embryos, the former predominantly in the placenta (555,556). Class II proto-oncogenes (me¬ diating GTP binding) are active in fetal tissues throughout development (557). The class III secreted proto-oncogene v-sis, homologous with platelet-derived growth factor, reaches

216

/ Chapter 6

its highest levels at 8 days in the mouse (552). Class IV nuclear proto-oncogenes c-fos and c-myc are actively ex¬ pressed during development, the former in the amnion and chorion of the mouse, and the latter in the cytotrophoblast of the human placenta (558,559). Although it is not yet possible to identify the role of each of these proto-oncogenes in mammalian development, their functions clearly deserve further analysis, because they may be among the multiple factors regulating embryonic and fetal growth.

Heat-Shock Proteins Exposure to heat and a variety of other environmental stresses induces the synthesis of a set of proteins known as heat-shock proteins (HSP) (560). Embryos of a wide variety of species, both vertebrate and invertebrate, synthesize HSP constitutively at early developmental stages (175). Mam¬ malian embryos are not exceptional, mouse embryos con¬ stitutively synthesizing HSP 68-70 at the 2-cell stage (197). Indeed, this is perhaps the first product of the mouse em¬ bryonic genome after division to the 2-cell stage (195,561). The capacity to respond to environmental stress by increased HSP synthesis does not appear until the blastocyst stage in mouse and rabbit embryos (562-564). At postimplantation stages, mouse embryos continue to synthesize HSP 68 con¬ stitutively in several extraembryonic tissues, but not in em¬ bryonic ectoderm (R. Kothary and J. Rossant, unpublished observations). The preimplantation period is the most heatsensitive time in mammalian development (565), with the most sensitive event in the mouse being activation of gene expression at the 2-cell stage (566). Even though the func¬ tion of HSP remains obscure, and the acquisition of ther¬ motolerance in mammalian embryos by means of HSP syn¬ thesis is still conjectural, the pattern of HSP synthesis suggests that these proteins may have an important role during cleav¬ age and early postimplantation development. Tissue-specific levels of HSP synthesis appear to support this view (567).

Homeobox Genes as Possible Regulatory Elements Morphogenesis of Drosophila is known to require the function of genes that control segment number and polarity and genes that specify segment identity (568-570). The latter group, known as homeotic genes, often possess a conserved sequence of nucleotides known as the “homeo¬ box.” Moreover, this family of sequences has remained highly conserved throughout evolution (571). In mammals (primarily the mouse and human have been studied) there is a steadily increasing number of known genes containing sequences homologous with the Drosophila homeobox (572— 579). Of these, two have been mapped to mouse chromo¬ some 1, one to chromosome 6, and five to chromosome 11. Two of the genes located on mouse chromosome 11 were mapped first to human chromosome 17, implying synteny between regions of these chromosomes (576,581). Although

several known morphogenetic mutations also map to these chromosomes in the mouse (215,577), none of them has been identified as a homeobox-containing gene. The horneobox gene on mouse chromosome 11 is expressed begin¬ ning at 7.5 d.g. (574), and several others are expressed at later stages, some specifically in the spinal cord and brain and others during spermatogenesis as well (574,578-583). Homeobox genes described in Xenopus are also expressed during early development (584,585). The extreme degree of evolutionary conservation of the homeobox sequences and the regulated developmental expression of these genes raise the possibility that they have an integral role in switch¬ ing differentiative pathways, as they do in Drosophila. The relatively small number of homeobox sequences and their restricted tissue expression in spinal cord make it seem unlikely that they regulate metamerism or developmental decision making in any general way. Nonetheless, the high arginine and lysine contents of polypeptides encoded by the conserved homeobox domains and the nuclear location of the product in Drosophila suggest that the homeobox se¬ quences themselves may have a DNA-binding function, whatever the role of the remainder of the protein may be (586). It is this intriguing idea that sets homeobox-contain¬ ing genes apart from those involved in strictly physiological activities for consideration as possible regulatory genes.

CONCLUSION The Mammalian Embryo In Vitro: Advantages and Limitations The success of experimental approaches to early mam¬ malian embryogenesis is largely attributable to work with cultured embryos. Since the 1950s, most investigators have used in vitro methods for part or all of their analyses. The enormous advantage conferred by this approach is the ac¬ cessibility of the embryo itself for treatment and interven¬ tion. During preimplantation stages, the embryo is normally free in the oviduct or uterine lumen anyway, so the task of maintaining the embryo in a normal state is one of approx¬ imating the fluid and gas conditions of the native environ¬ ment. Maintaining normal development in vitro throughout the preimplantation period has been accomplished with only a few species, including the mouse, rabbit, and human. In these cases, the successful culture was largely a result of empirically altering the growth conditions to optimize de¬ velopment, rather than mimicking reproductive tract con¬ ditions in vivo; these latter parameters were largely un¬ known. Nonetheless, the adequacy of preimplantation culture conditions for any species could be determined by trans¬ ferring preimplantation-stage embryos to the oviduct or uterus of the natural or a surrogate mother, with development to term as the endpoint (1). In the case of postimplantation culture, however, return¬ ing embryos to the uterus has been marginally successful,

Early Embryogenesis / without development to term after transfer (587). Thus, despite the advantages of accessibility provided by culture, postimplantation embryos cannot yet be restored to their normal intrauterine potential for growth. The extent of nor¬ mal growth and morphology in vitro for postimplantation embryos is limited to 2 to 3 days for rodents, which have been the subject of most experimental work (588). The alternative approach of culturing preimplantation embryos to postimplantation stages (589,590) yields a low incidence of successful development, and certain features of these embryos do not closely resemble those of the equivalent stages in vivo. The main differences are (a) the two-dimen¬ sional growth of trophoblast derivatives as a monolayer on the substrate used for culture and (b) the retardation of the embryo proper. These phenomena may be related, in the sense that failure to maintain three-dimensionality in vitro may prevent normal cell-cell interactions necessary for pla¬ cental morphogenesis and therefore embryonic nutrition; alternatively, autocrine factors produced by either placenta or embryo may become diluted by culture medium, thus depriving the embryo of essential growth-promoting sub¬ stances. Clearly, defining the nature of interactions between mammalian embryonic and maternal cells and tissues through membrane contacts, extracellular matrix, and humoral fac¬ tors remains one of the major tasks in early mammalian embryology. Mammalian Embryos as Models for Vertebrate Development Are laboratory animals justifiable models for vertebrate development? Whatever insights are gained in such model systems will have to be examined closely to verify their relevance to the human or to other vertebrates. This com¬ parison will have to take place at all levels of organization (e.g., cellular as well as molecular). It should be apparent from this review that current experimental approaches to mammalian embryos make them superb systems for many aspects of developmental biology. The body of knowledge obtained from transgenic mice alone attests to the power of these novel approaches. Among higher eukaryotes, mice rival Drosophila in their diversity of genetic markers. In situ hybridization and other amplification techniques now make it possible to study gene expression in the small tissue samples available from laboratory mammals. Finally, clonal analysis of cell lineages has provided insight into the ar¬ chitectural rules for construction of the mammalian embryo. Another compelling reason for studying mammals as models for vertebrate development is that development during and after gastrulation appears to be extensively conserved. This conclusion is based on recent work showing the extensive similarity of the gastrula fate map of the mouse to those of the chick and urodele embryo (127,138,169; K. A. Lawson, J. J. Meneses, and R. A. Pedersen, unpublished observa¬ tions). Consequently, it appears that the basic way of es¬ tablishing the vertebrate body plan has remained essentially

217

unchanged for more than 250 million years. Naturally, this could be an argument for using chicks, frogs, and fish to study mammals, but mammals have some features that jus¬ tify their use instead. These pertain mainly, but not exclu¬ sively, to extraembryonic tissues and structures. They in¬ clude the precocious differentiation and allocation of the progenitors of the extraembryonic lineages, random X-chromosome inactivation in the fetus, preferential paternal Xchromosome inactivation in placental and yolk sac tissues, and genetic imprinting of genes in the maternal and paternal genomes such that all individuals require both a mother and a father. Thus, mammals are deeply generalized in some aspects of their organogenesis, yet uniquely specialized in other aspects of their early gene expression. This combi¬ nation makes a compelling case for studying mechanisms of mammalian embryogenesis and organogenesis. Analysis of these mechanisms in mammals should be informative of developmental phenomena among vertebrates in general and should provide numerous opportunities for future beneficial applications to problems of human health.

ACKNOWLEDGMENTS Work in my laboratory was supported by the Office of Health and Environmental Research, U.S. Department of Energy, contract DE-AC03-76-SF01012. This review was partially written while I was a guest of the Laboratory of Developmental and Reproductive Toxicology of the Na¬ tional Institute of Environmental Health Sciences, Research Triangle Park, North Carolina, under the terms of the Inter¬ agency Personnel Act. I thank Drs. John Me Lachlan and E. Mitchell Eddy for their encouragement and support. I thank Drs. Greg Barsh, Julia Emerson, Kirstie Lawson, and Akiko Spindle for their comments, and Drs. Eileen Ad¬ amson, Verne Chapman, John Heath, Kirstie Lawson, and Janet Rossant for allowing me to cite their unpublished work. I am grateful to Michelle Bloom and Leslie Roberts for their assistance with the manuscript.

REFERENCES 1. Adams, C. E. (1982): Mammalian Egg Transfer. CRC Press, Boca Raton, Fla. 2. Brackett, B. G., Seidel, G. E., and Seidel, S. M. (editors) (1981): New Technologies in Animal Breeding. Academic Press, New York. 3. Hafez, E. S. E., and Semm, K. (editors) (1982): In Vitro Fertilization and Embryo Transfer. Alan R. Liss, New York. 4. Beier, H. M., and Lindner, H. R. (editors) (1983): Fertilization of the Human Egg In Vitro. Biological Basis and Clinical Application. Springer-Verlag, Berlin. 5. Jones, H. W., Jr., Jones, G. S., Hodgen, G. D., and Rosenwaks, Z. (editors) (1986): In Vitro Fertilization. Norfolk. Williams and Wilkins, Baltimore. 6. Seppala, M., and Edwards, R. G. (editors) (1985): In Vitro Fertil¬ ization and Embryo Transfer. New York Academy of Sciences, New York. 7. Trounson, A., and Woods, C. (editors) (1984): In Vitro Fertilization and Embryo Transfer. Churchill Livingstone, Edinburgh.

218

/ Chapter 6

8. Kaye, P. L. (1986): Metabolic aspects of the physiology of the preimplantation embryo. In: Experimental Approaches to Mammal¬ ian Embryonic Development, edited by J. Rossant and R. A. Ped¬ ersen, pp. 267-292. Cambridge University Press, Cambridge. 9. Bodemer, C. W. (1971): The biology of the blastocyst in historical perspective. In: Biology of the Blastocyst, edited by R. J. Blandau, pp. 1-25. University of Chicago Press, Chicago. 10. Horder, T. J., Witkowski, J. A., and Wilie, C. C. (editors) (1986): A History of Embryology. Cambridge University Press, London. 11. Heape, W. (1891): Preliminary note on the transplantation and growth of mammalian ova within a uterine foster mother. Proc. R. Soc. bond. [Biol.], 48:457. 12. Rossant, J., and Papaioannou, V. E. (1977): The biology of embryogenesis. In: Concepts in Mammalian Embryogenesis, edited by M. I. Sherman, pp. 1-36. MIT Press, Cambridge, Mass. 13. Whitten, W. K. (1956): Culture of tubal mouse ova. Nature, 177:96. 14. Brinster, R. L. (1965): Studies on the development of mouse embryos in vitro. II. The effect of energy source. J. Exp. Zool., 158:59-68. 15. Biggers, J. D., and Brinster, R. L. (1965): Biometrical problems in the study of early mammalian embryos in vitro. J. Exp. Zool., 158:39— 48. 16. Biggers, J. D., Whitten, W. K., and Whittingham, D. G. (1971): The culture of mouse embryos in vitro. In: Methods in Mammalian Embryology, edited by J. C. Daniel, Jr., pp. 86-116. W. H. Free¬ man, San Francisco. 17. Hogan, B., Costantini, F., and Lacy, E. (1986): Manipulating the Mouse Embryo. A Laboratory Manual, pp. 12-15. Cold Spring Har¬ bor Laboratory, Cold Spring Harbor, N.Y. 18. Edwards, R. G., and Gates, A. H. (1959): Timing of the stages of the maturation, divisions, ovulation, fertilization and the first cleav¬ age of eggs of adult mice treated with gonadotrophins. J. Endocri¬ nol., 18:292-304. 19. Tarkowski, A. K. (1961): Mouse chimaeras developed from fused eggs. Nature, 190:857-860. 20. Mintz, B. (1964): Formation of genetically mosaic mouse embryos, and early development of “lethal (tl2/f12)-normal” mosaics. J. Exp. Zool., 157:273-292. 21. McLaren, A. (1976): Mammalian Chimaeras. Cambridge University Press, London. 22. Chapman, V. M., Ansell, J. D., and McLaren, A. (1972): Trophoblast giant cell differentiation in the mouse: expression of glucose phosphate isomerase (GPI-1) electrophoretic variants in transferred and chimeric embryos. Dev. Biol., 29:48-54. 23. West, J. D. (1978): Analysis of clonal growth using chimaeras and mosaics. In: Development in Mammals, Vol. 3, edited by M. H. Johnson, pp. 413-460. North-Holland, Amsterdam. 24. LeDouarin, N., and McLaren, A. (editors) (1984): Chimeras in De¬ velopmental Biology. Academic Press, New York. 25. Gardner, R. L. (1968): Mouse chimaeras obtained by the injection of cells into the blastocyst. Nature, 220:596-597. 26. Gardner, R. L. (1984): An in situ cell marker for clonal analysis of development of the extraembryonic endoderm in the mouse. J. Embryol. Exp. Morphol., 80:251-288. 27. Rossant, J., Vijh, M., Siracusa, L. D., and Chapman, V. M. (1983): Identification of embryonic cell lineages in histological sections of M. musculus M. caroli chimaeras. J. Embryol. Exp. Morphol., 73:179-191. 28. Solter, D., and Knowles, B. (1975): Immunosurgery of mouse blas¬ tocyst. Proc. Natl. Acad. Sci. USA, 72:5099-5102. 29. Lin, T. P. (1971): Egg micromanipulation. In: Methods in Mam¬ malian Embryology, edited by J. C. Daniels, Jr., pp. 157-185. W. H. Freeman, San Francisco. 30. Wilson, I. B., Bolton, E., and Cutler, R. H. (1972): Preimplantation differentiation in the mouse egg as revealed by microinjection of vital markers. J. Embryol. Exp. Morphol., 27:467^179. 31. Graham, C. F., and Deussen, Z. A. (1978): Features of cell lineage in preimplantation mouse development. J. Embryol. Exp. Morphol., 48:53-72. 32. Jacobson, M., and Hirose, G. (1978): Origin of the retina from both sides of the embryonic brain: a contribution to the problem of crossing at the optic chiasma. Science, 202:637-639. 33. Weisblat, D. A., Sawyer, R. T., and Stent, G. S. (1978): Cell lineage analysis by intracellular injection of a tracer enzyme. Science, 202:1295-1298.

34. Gimlich, R. L., and Braun, J. (1985): New fluorescent cell lineage tracers. Dev. Biol., 115:340-352. 35. Balakier, H., and Pedersen, R. A. (1982): Allocation of cells to inner cell mass and trophectoderm lineages in preimplantation mouse em¬ bryos. Dev. Biol., 90:352-362. 36. Pedersen, R. A. (1987): Analysis of cell lineage during early mouse embryogenesis. In: Developmental Toxicology: Mechanisms and Risk, edited by J. A. McLachlan, R. M. Pratt, and C. L. Markert. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y (in press). 37. Briggs, R., and King, T. J. (1952): Transplantation of living nuclei from blastula cells into enucleated frogs’ eggs. Proc. Natl. Acad. Sci. USA, 38:455^*63. 38. Gurdon, J. B. (1962): The developmental capacity of nuclei taken from intestinal epithelium cells of feeding tadpoles. J. Embryol. Exp. Morphol., 10:622. 39. McGrath, J., and Solter, D. (1983): Nuclear transplantation in the mouse embryo by microsurgery and cell fusion. Science, 220:13001303. 40. Gordon, J. W., Scangos, G. A., Plotkin, D. J., Barbosa, J. A., and Ruddle, F. H. (1980): Genetic transformation of mouse embryos by microinjection of purified DNA. Proc. Natl. Acad. Sci. USA, 77: 7380-7384. 41. Brinster, R. L., and Palmiter, R. D. (1986): Introduction of genes into the germ line of animals. Harvey Lect., 80:1-38. 42. Rubin, G. M., and Spradling, A. C. (1982): Transposition of cloned P elements into Drosophila germ line chromosomes. Science, 218:348353. 43. Gaddum-Rosse, P., Blandau, R. J., Langley, L. B., and Sato, K. (1982): Sperm tail entry into the mouse egg in vitro. Gamete Res., 6:215-223. 44. Howlett, S. K., and Bolton, V. N. (1985): Sequence and regulation of morphological and molecular events during the first cell cycle of mouse embryogenesis. J. Embryol. Exp. Morphol., 87:175-206. 45. Yanagimachi, R. (1966): Time and process of sperm penetration into hamster and ova in vivo and in vitro. J. Reprod. Fertil., 11:359370. 46. Molls, M., Zamboglou, N., and Streffer, C. (1983): A comparison of the cell kinetics of preimplantation mouse embryos from two different mouse strains. Cell Tissue Kinet., 16:277-283. 47. Naish, S. J., Perreault, S. D., Foehner, A. L., and Zirkin, B. R. (1987): DNA synthesis in the fertilizing hamster sperm nucleus: sperm template availability and egg cytoplasmic control. Biol. Reprod., (in press). 48. Shire, J. G. M., and Whitten, W. K. (1980): Genetic variation in the timing of first cleavage in mice: effect of paternal genotype. Biol. Reprod., 23:363-368. 49. Shire, J. G. M., and Whitten, W. K. (1980): Genetic variation in the timing of first cleavage in mice: effect of maternal genotype. Biol. Reprod., 23:369-376. 50. Goldbard, S. B., and Warner, C. M. (1982): Genes affect the timing of early mouse embryo development. Biol. Reprod., 27:419^*24. 51. Harlow, G. M., and Quinn, P. (1982): Development of pre-implan¬ tation mouse embryos in vivo and in vitro. Aust. J. Biol. Sci., 35:187193. 52. Streffer, C., van Beuningen, D., Molls, M., Zamboglou, N., and Schultz, S. (1980): Kinetics of cell proliferation in the preim¬ planted mouse embryo in vivo and in vitro. Cell Tissue Kinet., 13: 135-143. 53. Sawicki, W., Abramczuk, J., and Blaton, O. (1978): DNA cycles in the second and third cell cycles of mouse preimplantation devel¬ opment. Exp. Cell Res., 112:199-205. 54. Pedersen, R. A. (1986): Potency, lineage and allocation in preim¬ plantation mouse embryos. In: Experimental Approaches to Mam¬ malian Embryonic Development, edited by J. Rossant and R. A. Pedersen, pp. 3-33. Cambridge University Press, Cambridge. 55. Smith, R. K. W., and Johnson, M. H. (1986): Analysis of the third and fourth cell cycles of mouse early development. J. Reprod. Fertil. 76:393-399. 56. Sundstrom, P., Nilsson, O., and Liedholm, P. (1981): Cleavage rate and morphology of early human embryos obtained after artificial fertilization and culture. Acta Obstet. Gynecol. Scand., 60:109-120. 57. Trouson, A. O., Mohr, L. R., Wood, C., and Leeton, J. F. (1982): Effect of delayed insemination on in-vitro fertilization, culture and transfer of human embryos. J. Reprod. Fertil., 64:285-294.

Early Embryogenesis /

58. Kelly, S. J., Mulnard, J. G., and Graham, C. F. (1978): Cell division and cell allocation in early mouse development. J. Embryol. Exp. Morphol., 48:37-51. 59. Lewis, W. H., and Wright, E. S. (1935): On the early development of the mouse egg. Carnegie Instit. Contrib. Embryol., 25:113-143. 60. Ducibella, T., Albertini, D. F., Anderson, E., and Biggers, J. (1975): The preimplantation mammalian embryo: characterization of intra¬ cellular junctions and their appearance during development. Dev. Biol., 45:231-250. 61. Magnuson, T., Dempsey, A., and Stackpole, C. W. (1977): Char¬ acterization of intercellular junctions in the preimplantation mouse embryo by freeze-fracture and thin-section electron microscopy. Dev. Biol., 61:252-261. 62. Magnuson, T., Jacobson, J. B., and Stackpole, C. W. (1978): Re¬ lationship between intercellular permeability and junctional organi¬ zation in the preimplantation mouse embryo. Dev. Biol., 67:214224. 63. McLachlin, J. R., Caveney, S., and Kidder, G. M. (1983): Control of gap junction formation in early mouse embryos. Dev. Biol., 98:155164. 64. Goodall, H., and Johnson, M. H. (1984): The nature of intercellular coupling within the preimplantation mouse embryo. J. Embryol. Exp. Morphol., 79:53-76. 65. Selwood, L., and Young, G. J. (1983): Cleavage in vivo and in culture in the dasyurid marsupial Antechinus stuartii (Macleay). J. Morphol., 176:43-60. 66. Wimsatt, W. A. (1975): Some comparative aspects of implantation. Biol. Reprod., 12:1 —40. 67. Papaioannou, V. E., and Ebert, K. M. (1986): Comparative aspects of embryo manipulation in mammals. In: Experimental Approaches to Mammalian Embryonic Development, edited by J. Rossant and R. A. Pedersen, pp. 67-96. Cambridge University Press, Cambridge. 68. Barlow, P.,Owen, D. A. J., and Graham, C. (1972): DNA synthesis in the preimplantation mouse embryo. J. Embryol. Exp. Morphol., 27:431—455. 69. Chisholm, J. C., Johnson, M. H., Warren, P. D., Fleming, T. P., and Pickering, S. J. (1985): Developmental variability within and between mouse expanding blastocysts and their ICMs. J. Embryol. Exp. Morphol., 86:311-336. 70. Smith, R., and McLaren, A. (1977): Factors affecting the time of formation of the mouse blastocoel. J. Embryol. Exp. Morphol., 41:7992. 71. Sawicki, W., and Mystkowska, E. T. (1981): Phorbol ester-mediated modulation of cell proliferation and primary differentiation of mouse preimplantation embryos. Exp. Cell Res., 136:455—458. 72. Alexandre, H. (1979): The utilization of an inhibitor of spermidine and spermine synthesis as a tool for the study of determination of cavitation in the preimplantation mouse embryo. J. Embryol. Exp. Morphol., 53:145-162. 73. Alexandre, H. (1982): Effet de Tinhibition specifique de la replication de l’ADN par l’aphidicoline sur la differentiation primarie de l’oeuf de souris en preimplantation. C. R. Acad. Sci. (Paris), 294:1001 1006. 74. Dean, W. L., and Rossant, J. (1984): Effect of delaying DNA rep¬ lication on blastocyst formation in the mouse. Differentiation, 26:134137. 75. Spindle, A. I., Nagano, H., and Pedersen, R. A. (1985): Inhibition of DNA replication in preimplantation mouse embryos by aphidicolin. J. Exp. Zool., 235:289-295. 76. Newport, J., and Kirschner, M. (1982): A major developmental transition in early Xenopus embryos: II. Control of the onset of transcription. Cell, 30:687-696. 77. Mita-Miyazawa, I., Ikegami, S., and Satoh, N. (1985). Histospecific acetylcholinesterase development in the presumptive muscle cells isolated from 16-cell-stage ascidian embryos with respect to the number of DNA replications. J. Embryol. Exp. Morphol., 87:

1-12. 78. Waksmundzka, M., Krysiak, E., Karasiewicz, J., Czolowska, R., and Tarkowski, A. K. (1984): Autonomous cortical activity in mouse eggs controlled by a cytoplasmic clock. J. Embryol. Exp. Morphol., 79:77-96. r , 79. Masui, Y., and Markert, C. L. (1971): Cytoplasmic control of nuclear behavior during meiotic maturation of frog oocytes. J. Exp. Zool., 117:129-146.

219

80. Newport, J., and Kirschner, M. (1984): Regulation of the cell cycle during early Xenopus development. Cell, 37:731-742. 81. Kishimoto, T., Yamazaki, K., Kato, Y., Koide, S. S., and Kanatani, H. (1984): Induction of starfish oocyte maturation by maturationpromoting factor of mouse and surf clam oocytes. J. Exp. Zool., 231:293-295. 82. Sorensen, R. A., Cyert, M. S., and Pedersen, R. A. (1985): Active maturation promoting factor is present in mature mouse oocytes. J. Cell Biol., 100:1637-1640. 83. Balakier, H., and Czolowska, R. (1977): Cytoplasmic control of nuclear maturation in mouse oocytes. Exp. Cell Res., 110:466-469. 84. Balakier, H. (1978): Induction of maturation in small oocytes from sexually immature mice by fusion with meiotic or mitotic cells. Exp. Cell Res., 112:137-141. 85. Balakier, H., and Masui, Y. (1986): Chromosome condensation ac¬ tivity in the cytoplasm of anucleate and nucleate fragments of mouse oocytes. Dev. Biol., 113:155-159. 86. Masui, Y., and Clarke, J. H. (1979): Oocyte maturation. Int. Rev. Cytol., 57:185-283. 87. Berridge, M. J. (1984): Inositol triphosphate and diacylglycerol as second messengers. Biochem. J., 220:345-360. 88. Macara, I. G. (1985): Oncogenes, ions, and phospholipids. Am. J. Physiol. (Cell Physiol., 17), 248:C3-C11. 89. Busa, W. B., and Nuccitelli, R. (1984): Metabolic regulation via intracellular pH. Am. J. Physiol., 246:R409-R438. 90. Zwierzchowski, L., Czlonkowska, M., and Guszkiewicz, A. (1986): Effect of polyamine limitation on DNA synthesis and development of mouse preimplantation embryos in vitro. J. Reprod. Fertil., 76:115121. 91. Heby, O. (1981): Role of polyamines in the control of cell prolif¬ eration and differentiation. Differentiation, 19:1-20. 92. Weiss, P. (1939): Principles of Development. Holt, New York. 93. Tarkowski, A. K., and Wroblewska, J. (1967): Development of blastomeres of mouse eggs isolated at the 4- and 8-cell stage. J. Embryol. Exp. Morphol., 18:155-180. 94. Tarkowski, A. K. (1959): Experiments on the development of iso¬ lated blastomeres of mouse eggs. Nature, 184:1286-1287. 95. Kelly, S. J. (1977): Studies of the developmental potential of 4- and 8-cell stage mouse blastomeres. J. Exp. Zool., 200:365-376. 96. Hillman, N., Sherman, M. I., and Graham, C. F. (1972): The effect of spatial arrangement on cell determination during mouse devel¬ opment. J. Embryol. Exp. Morphol., 28:263-278. 97. Rossant, J., and Vijh, K. M. (1980): Ability of outside cells from preimplantation mouse embryos to form inner cell mass derivatives. Dev. Biol., 76:475^182. 98. Ziomek, C. A., Johnson, M. H., and Handyside, A. H. (1982): The developmental potential of mouse 16-cell blastomeres. J. Exp. Zool., 221:345-355. 99. Stem, M. S. (1972): Experimental studies on the organization of the preimplantation mouse embryo. II. Reaggregation of disaggregated embryos. J. Embryol. Exp. Morphol., 28:255-261. 100. Handyside, A. H. (1978): Time of commitment of inside cells iso¬ lated from preimplantation mouse embryos. J. Embryol. Exp. Mor¬ phol., 45:37-53. 101. Hogan, B., and Tilly, R. (1978): In vitro development of inner cell masses isolated immunosurgically from mouse blastocysts. II. Inner cell masses from 3.5- to 4.0 day p.c. blastocysts. J. Embryol. Exp. Morphol., 45:107-121. 102. Spindle, A. I. (1978): Trophoblast regeneration by inner cell masses isolated from cultured mouse embryos. J. Exp. Zool., 203:483—489. 103. Rossant, J., and Lis, W. T. (1979): Potential of isolated mouse inner cell masses to form trophectoderm derivatives in vivo. Dev. Biol., 70:255-261. 104. Rossant, J. (1975): Investigation of the determinative state of the mouse inner cell mass. II. The fate of isolated inner cell masses transferred to the oviduct. J. Embryol. Exp. Morphol., 33: 991-1001. 105. Nichols, J., and Gardner, R. L. (1984): Heterogeneous differentiation of external cells in individual isolated early mouse inner cell masses in culture. J. Embryol. Exp. Morphol., 80:225-240. 106. Izquierdo, L., and Ortiz, M. E. (1975): Differentiation in the mouse morulae. Wilhelm Roux’s Archiv, 177:67—74. 107. Daniel, J. C., Jr. (1976): The first potential I.C.M. cell during cleav¬ age of the rabbit ovum. Wilhelm Roux’s Archiv, 179:249-250.

220

/ Chapter 6

108. Handyside, A. H. (1981): Immunofluorescence techniques for de¬ termining the numbers of inner and outer blastomeres in mouse morulae. J. Reprod. Immunol., 2:339-350. 109. Copp, A. J. (978): Interaction between inner cell mass and trophectoderm of the mouse blastocyst. I. A study of cellular proliferation. J. Embryol. Exp. Morphol., 48:109-125. 110. Gamer, W., and McLaren, A. (1974): Cell distribution in chimaeric mouse embryos before implantation. J. Embryol. Exp. Morphol., 32:495-503. 111. Kelly, S. J. (1979): Investigations into the degree of cell mixing that occurs between the 8-cell and the blastocyst stage of mouse devel¬ opment. J. Exp. Zool., 207:121-130. 112. Surani, M. A. H., and Barton, S. C. (1984): Spatial distribution of blastomeres is dependent on cell division order and interactions in mouse morulae. Dev. Biol., 102:335-343. 113. Spindle, A. I. (1982): Cell allocation in preimplantation mouse chi¬ meras. J. Exp. Zool., 219:361-367. 114. Meinecke-Tillmann, S., and Meinecke, B. (1984): Experimental chimaeras—removal of reproductive barrier between sheep and goat. Nature, 307:637-638. 115. Surani, M. A. H., Kimbers, S. J., and Barton, S. C. (1981): Dif¬ ferential adhesiveness as a mechanism of cell allocation to inner cell mass and trophectoderm in the mouse blastocyst. In: Culture Tech¬ niques . Applicability for Studies on Prenatal Differentiation and Tox¬ icity, edited by D. Neubert and H.-J. Merker, pp. 397^412. W. de Gruyter, Berlin. 116. Kimber, S. J., Surani, M. A. H., and Barton, S. C. (1982): Inter¬ actions of blastomeres suggest changes in cell surface adhesiveness during the formation of inner cell mass and trophectoderm in the preimplantation mouse embryo. J. Embryol. Exp. Morphol., 58: 231-249. 117. Handyside, A. H. (1980): Distribution of antibody- and lectin-bind¬ ing sites on dissociated blastomeres from mouse morulae: evidence of polarization at compaction. J. Embryol. Exp. Morphol., 60:99116. 118. Ziomek, C. A., and Johnson, M. H. (1980): Cell surface interaction induces polarization of mouse 8-cell blastomeres at compaction. Cell, 21:935-942. 119. Ziomek, C. A., and Johnson, M. H. (1982): The roles of phenotype and position in guiding the fate of 16-cell mouse blastomeres. Dev. Biol., 91:440^147. 120. Johnson, M. H., and Ziomek, C. A. (1983): Cell interactions influ¬ ence the fate of mouse blastomeres undergoing the transition from the 16- to the 32-cell stage. Dev. Biol., 95:211-218. 121. Pedersen, R. A., Wu, K., and Balakier, H. (1986): Origin of the inner cell mass in mouse embryos: cell lineage analysis by microin¬ jection. Dev. Biol., 117:581-595. 122. Cruz, Y. P., and Pedersen, R. A. (1985): Cell fate in the polar trophectoderm of mouse blastocysts as studied by microinjection of cell lineage tracers. Dev. Biol., 112:73-83. 123. Pedersen, R. A., and Spindle, A. I. (1980): Role of the blastocoel microenvironment in early mouse embryo differentiation. Nature 284:550-552. 124. Johnson, M. H., Pratt, H. M. P., and Handyside, A. H. (1981): The generation and recognition of positional information in the preim¬ plantation mouse embryo. In: Cellular and Molecular Aspects of Implantation, edited by S. R. Glasser and D. W. Bullock, pp. 5574. Plenum, New York. 125. Johnson, M. H., and Maro, B. (1986): Time and space in the mouse early embryo: a cell biological approach to cell diversification. In: Experimental Approaches to Mammalian Embryonic Development, edited by J. Rossant and R. A. Pedersen, pp. 35-65. Cambridge University Press, Cambridge. 126. Rossant, J. (1977): Cell commitment in early rodent development. In: Development in Mammals, Vol. 2, edited by M. H. Johnson, pp. 119-150. North-Holland, Amsterdam. 127. Beddington, R. (1986): Analysis of tissue fate and prospective po¬ tency in the egg cylinder. In: Experimental Approaches to Mam¬ malian Embryonic Development, edited by J. Rossant and R. A. Pedersen, pp. 121-147. Cambridge University Press, Cambridge. 128. Gardner, R. L., Papaioannou, V. E., and Barton, S. C. (1973): Origin of the ectoplacental cone and secondary giant cells in mouse blastocysts reconstituted from isolated trophoblast and inner cell mass. J. Embryol. Exp. Morphol., 30:561-572.

129. Papaioannou, V. E. (1982): Lineage analysis of inner cell mass and trophectoderm using microsurgically reconstituted mouse blasto¬ cysts. J. Embryol. Exp. Morphol., 68:199-209. 130. Rossant, J., and Croy, B. A. (1985): Genetic identification of tissue of origin of cellular populations within the mouse placenta. J. Em¬ bryol. Exp. Morphol., 86:177-189. 131. Johnson, M. H. (1979): Molecular differentiation of inside cells and inner cell masses isolated from the preimplantation mouse embryo. J. Embryol. Exp. Morphol., 53:335-344. 132. Rossant, J., Gardner, R. L., and Alexandre, H. L. (1978): Inves¬ tigation of the potency of cells from the postimplantation mouse embryo by blastocyst injections. A preliminary report. J. Embryol. Exp. Morphol., 48:239-247. 133. Copp, A. J. (1979): Interaction between inner cell mass and tro¬ phectoderm of the mouse blastocyst. II. Fate of the polar trophec¬ toderm. J. Embryol. Exp. Morphol., 51:109-120. 134. Rossant, J. (1986): Development of extraembryonic cell lineages in the mouse embryo. In: Experimental Approaches to Mammalian Em¬ bryonic Development, edited by J. Rossant and R. A. Pedersen, pp. 97-120. Cambridge University Press, Cambridge. 135. Gardner, R. L., and Rossant, J. (1979): Investigation of the fate of 4.5 day post-coitum mouse inner cell mass cells by blastocyst in¬ jection. J. Embryol. Exp. Morphol., 52:141-152. 136. Gardner, R. L. (1982): Investigation of cell lineage and differentia¬ tion in the extraembryonic endoderm of the mouse embryo. J. Em¬ bryol. Exp. Morphol., 68:175-198. 137. Hogan, B. L. M., and Tilly, R. (1981): Cell interactions and en¬ doderm differentiation in cultured mouse embryos. J. Embryol. Exp. Morphol., 62:379-394. 138. Lawson, K. A., Meneses, J. J., and Pedersen, R. A. (1986): Cell fate and cell lineage in the endoderm of the presomite mouse embryo, studied with an intracellular tracer. Dev. Biol., 115 325329. 139. Monk, M., and Harper, M. I. (1979): Sequential X chromosome inactivation coupled with cellular differentiation in early mouse embros. Nature, 281:311-313. 140. Rossant, J. (1984): Somatic cell lineages in mammalian chimeras. In: Chimeras in Developmental Biology, edited by N. LeDouarin and A. McLaren, pp. 89-109. Academic Press, Orlando, FI. 141. Gardner, R. L., Lyon, M. F., Evans, E. P., and Burtenshaw, M. D. (1985): Clonal analysis of X-chromosome inactivation and the origin of the germ line in the mouse embryo. J. Embryol. Exp. Morphol., 88:349-363. 142. Gardner, R. L. (1985): Regeneration of endoderm from primitive ectoderm in the mouse embryo: fact or artifact? J. Embryol. Exp. Morphol., 88:303-326. 143. Pedersen, R. A., Spindle, A. I., and Wiley, L. M. (1977): Regen¬ eration of endoderm by ectoderm isolated from mouse blastocysts Nature, 270:435-437. 144. Dziadek, M. (1979): Cell differentiation in isolated inner cell masses of mouse blastocysts in vitro: onset of specific gene expression. J. Embryol. Exp. Morphol., 53:367-379. 145. Atienza-Samols, S. B., and Sherman, M. I. (1979): In vitro devel¬ opment of core cells of the inner cell mass of the mouse blastocyst: effects of conditioned medium. J. Exp. Zool., 208:67-71. 146. Svajger, A., Levak-Svajger, B., and Skreb, N. (1986): Rat embry¬ onic ectoderm as renal isograft. J. Embryol. Exp. Morphol., 94:1147. Grobstein, C. (1951): Intraocular growth and differentiation of the mouse embryonic shield implanted directly and following in vitro cultivation. J. Exp. Zool., 116:501-525. 148. Grobstein, C. (1952): Intraocular growth and differentiation of clus¬ ters of mouse embryonic shields cultured with and without primitive endoderm and in the presence of possible inductors. J Exp Zool 119:355-380. 149. Levak-Svajger, B., and Svajger, A. (1971): Differentiation of endodermal tissues in homografts of primitive ectoderm from two¬ layered rat embryonic shields. Experientia, 27:683-684. 150. Skreb, N., and Svajger, A. (1975): Experimental teratomas in rats. In: Teratomas and Differentiation, edited by D. Solter, pp. 83-97. Academic Press, London. 151. Diwan, S. B., and Stevens, L. C. (1976): Development of teratomas from ectoderm of mouse egg cylinders. J. Natl. Cancer Inst 57 937942.

Early Embryogenesis /

152. Beddington, R. S. P. (1983): Histogenic and neoplastic potential of different regions of the mouse embryonic egg cylinder. J. Embryol. Exp. Morphol., 75:189-204. 153. Levak-Svajger, B., and Svajger, A. (1974): Investigation of the origin of definitive endoderm in the rat embryo. J. Embryol. Exp. Morphol., 32:445-459. 154. Solter, D., and Damjanov, I. (1973): Explantation of cxtraembryonic parts of 7d mouse egg cylinders. Experientia, 29:701-703. 155. Skreb, N., Svajger, A., and Levak-Svajger, B. (1976): Develop¬ mental potentialities of the germ layers. In: Embryogenesis in Mam¬ mals, CIBA Foundation Symposium 40 (new series), pp. 27-39. Elsevier, Amsterdam. 156. Svajger, A., Levak-Svajger, B., Kostovic-Knezevic, L., and Bradamante, Z. (1981): Morphogenetic behaviour of the rat embryonic ectoderm as a renal homograft. J. Embryol. Exp. Morphol. [Suppl.], 65:243-267. 157. Tam, P. P. L. (1984): The histogenetic capacity of tissues in the caudal end of the embryonic axis of the mouse. J. Embryol. Exp. Morphol., 82:253-266. 158. Snow, M. H. L. (1981): Autonomous development of parts isolated from primitive-streak-stage mouse embryos. Is development clonal? J. Embryol. Exp. Morphol. [Suppl.], 65:269-287. 159. Ozdzenski, W. (1967): Observations on the origin of the primordial germ cells in the mouse. Zool. Pol., 17:367-379. 160. Rosenquist, G. C. (1966): A radioautographic study of labeled grafts in the chick blastoderm. Development from primitive-streak stages to stage 12. Carnegie Inst. Wash. Contrib. Embryol., 38:71-110. 161. Beddington, R. S. P. (1981): An autoradiographic analysis of the potency of embryonic ectoderm in the 8th day postimplantation mouse embryo. J. Embryol. Exp. Morphol., 64:87-104. 162. Beddington, R. S. P. (1982): An autoradiographic analysis of tissue potency in different regions of the embryonic ectoderm during gastrulation in the mouse. J. Embryol. Exp. Morphol., 69:265-285. 163. Copp, A. J., Roberts, H. M., and Polani, P. E. (1986): Chimaerism of primordial germ cells in the early postimplantation mouse embryo following microsurgical grafting of posterior primitive streak cells in vitro. J. Embryol. Exp. Morphol., 15:95-115. 164. Soriano, P., and Jaenisch, R. (1986): Retroviruses as probes for mammalian development: allocation of cells to the somatic and germ cell lineages. Cell, 46:19-29. 165. Wilkie, T. M., Brinster, R. L., and Palmiter, R. D. (1986): Germline and somatic mosaicism in transgenic mice. Dev. Biol., 118:9-18. 166. Russell, L. B., and Russell, W. L. (1954): Analysis of the changing radiation response of the developing mouse embryo. J. Cell. Comp. Physiol. [Suppl. 1], 43:103-147. 167. Pedersen, R. A. (1987): Analysis of cell lineage during early mouse embryogenesis. In: Developmental Toxicology: Mechanisms and Risk, edited by J. A. McLachlan, R. M. Pratt, and C. L. Markert. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. (in press). 168. Gardner, R. L. (1978): The relation between cell lineage and dif¬ ferentiation in the early mouse embryo. In: Genetic Mosaics and Cell Differentiation, edited by W. J. Gehring, pp. 205-241. SpringerVerlag, Berlin. 169. Beddington, R. S. P. (1983): The origin of the foetal tissues during gastrulation in the rodent. In: Development in Mammals, Vol. 5, edited by M. H. Johnson, pp. 1-32. Elsevier, Amsterdam. 170. Sanes, J. R., Rubenstein, J. L. R., and Nicolas, J.-F. (1986): Use of a recombinant retrovirus to study post-implantation cell lineage in mouse embryos. EMBO J., 5:3133—3142. 171. Rossant, J. (1987): Cell lineage analysis in mammalian embryo¬ genesis. Curr. Top. Dev. Biol., (in press). 172. Johnson, M. H., Handyside, A. H., and Braude, P. R. (1977): Control mechanisms in early mammalian development. In: Devel¬ opment in Mammals, Vol. 2, edited by M. H. Johnson, pp. 67-97. North-Holland, Amsterdam. 173. Johnson, M. H. (1981): The molecular and cellular basis of preim¬ plantation mouse development. Biol. Rev., 56:463-498. 174. Schultz, G. A., Clough, J. R., Braude, P. R., Pelham, H. R. B., and Johnson, M. H. (1981): A reexamination of messenger RNA populations in the preimplantation mouse embryo. In: Cellular and Molecular Aspects of Implantation, edited by S. R. Glasser and D. W. Bullock, pp. 137-154. Plenum, New York. 175. Schultz, G. A. (1986): Utilization of genetic information in the preim¬ plantation mouse embryo. In: Experimental Approaches to Mam¬

176.

177.

178.

179.

180.

181.

182.

183.

184.

185.

186.

187.

188.

189. 190.

191. 192.

193.

194.

195.

196.

197.

198.

221

malian Embryonic Development, edited by J. Rossant and R. A. Pedersen, pp. 239-265. Cambridge University Press, Cambridge. Piko, L., and Clegg, K. B. (1982): Quantitative changes in total RNA, total poly(A), and ribosomes in early mouse embryos. Dev. Biol., 89:362-378. Clegg, K. B., and Piko, L. (1983): Poly(A) length, cytoplasmic adenylation, and synthesis of poly A + RNA in early mouse embryos. Dev. Biol., 95:331-341. Giebelhaus, D. H., Weitlauf, H. M., and Schultz, G. A. (1985): Actin mRNA content in normal and delayed implanting mouse em¬ bryos. Dev. Biol., 107:407-413. Graves, R. A., Marzluff, W. F., Giebelhaus, D. H., and Schultz, G. A. (1985): Quantitative and qualitative changes in histone gene expression during early mouse development. Proc. Natl. Acad. Sci. USA, 82:5685-5689. Braude, P. R., Pelham, H., Flach, G., and Lobatto, R. (1979): Posttranscriptional control in the early mouse embryo. Nature, 282:102105. van Blerkom, J. (1981): The structural relation and post-translational modification of stage-specific proteins synthesized during early preimplantation development in the mouse. Proc. Natl. Acad. Sci. USA, 78:7629-7633. van Blerkom, J. (1985): Post-translational regulation of early de¬ velopment in the mammal. In: Differentiation and Proliferations, edited by C. Venizale, pp. 67-86. Van Nostrand Reinhold, New York. van Blerkom, J., and Runner, M. N. (1984): Mitochondrial reor¬ ganization during resumption of arrested meiosis in the mouse oocyte. Am. J. Anat., 171:335-355. Howlett, S. K., and Bolton, V. N. (1985): Sequence and regulation of morphological and molecular events during the first cell cycle of mouse embryogenesis. J. Embryol. Exp. Morphol., 87:175-206. Kaplan, G., Jelinek, W. R., and Bachvarova, R. (1985): Repetitive sequence transcripts and U1 RNA in mouse oocytes and eggs. Dev. Biol., 109:15-24. Forbes, D. J., Komberg, T. B., and Kirschner, M. W. (1983): Small nuclear RNA transcription and ribonucleoprotein assembly in early Xenopus development. J. Cell Biol., 97:62-72. Zeller, R., Nyffenegger, T., and DeRobertis, E. M. (1983): Nucleocytoplasmic distribution of snRNPs and stockpiled snRNA-binding proteins during oogenesis and early development in Xenopus laevis. Cell, 32:425^134. Ebert, K. M., and Brinster, R. L. (1983): Rabbit a-globin messenger RNA translation by the mouse ovum. J. Embryol. Exp. Morphol., 74:159-168. Brinster, R. L., Wiebold, J. L., and Brunner, S. (1976): Protein metabolism in preimplanted mouse ova. Dev. Biol., 51:215-224. Merz, E. A., Brinster, R. L., Brunner, S., and Chen, H. Y. (1981): Protein degradation during preimplantation development of the mouse. J. Reprod. Fertil., 61:415-418. Bachvarova, R., and De Leon, V. (1977): Stored and polysomal ribosomes of mouse ova. Dev. Biol., 58:248-254. Kidder, G. M., and Conlon, R. A. (1985): Utilization of cytoplasmic poly(A) + RNA for protein synthesis in preimplantation mouse em¬ bryos. J. Embryol. Exp. Morphol., 89:223-234. Brinster, R. L., Chen, H. Y., Trumbauer, M. E., and Avarbock, M. R. (1980): Translation of rabbit globin messenger RNA by the mouse ovum. Nature, 283:499-501. Petzoldt, U., Hoppe, P. C., and Illmensee, K. (1980): Protein syn¬ thesis in enucleated fertilized and unfertilized mouse eggs. Wilhelm Roux’s Arch. Dev. Biol., 189:215-219. Flach, G., Johnson, M. H., Braude, P. R., Taylor, R. A. S., and Bolton, V. N. (1982): The transition from maternal to embryonic control in the 2-cell mouse embryo. EMBO J., 1:681-686. Bolton, V. N., Oades, P. J., and Johnson, M. H. (1984): The re¬ lationship between cleavage, DNA replication, and gene expression in the mouse 2-cell embryo. J. Embryol. Exp. Morphol., 79:139163. Bensaude, O Babinet, C., Morange, M., and Jacob, F. (1983): Heat shock proteins, first major products of zygotic gene activity in the mouse embryo. Nature, 305:331-333. Clegg, K. B., and Piko, L. (1983): Quantitive aspects of RNA synthesis and polyadenylation in 1-cell and 2-cell mouse embryos. J. Embryol. Exp. Morphol., 74:169-182.

222

/ Chapter 6

199. Wudl, L., and Chapman, V. (1976): The expression of 3-glucuron¬ idase during preimplantation development of mouse embryos. Dev. Biol., 48:104-109. 200. Sawicki, J. A., Magnuson, T., and Epstein, C. J. (1982): Evidence for expression of the paternal genome in the two-cell mouse embryo. Nature, 294:450-451. 201. Young, R. J., and Sweeny, K. (1979): Adenylation and ADP-ribosylation in the mouse 1-cell embryo. J. Embryol. Exp. Morphol., 49:139-152. 202. Young, R. J. (1977): Appearance of 7-methylguanosine-5'-phosphate in the RNA of 1-cell embryos three hours after fertilization. Biochem. Biophys. Res. Commun., 76:32-39. 203. Schultz, G. A., Clough, J. R., and Johnson, M. H. (1980): Presence of cap structures in messenger RNA of mouse eggs. J. Embryol. Exp. Morphol., 56:139-156. 204. Young, R. J., Sweeny, K., and Bedford, J. M. (1978): Uridine and guanosine incorporation by the mouse one-cell embryo. J. Embryol. Exp. Morphol., 44:133-148. 205. Braude, P. R. (1979): Control of protein synthesis during blastocyst formation in the mouse. Dev. Biol., 68:440^452. 206. Braude, P. R. (1979): Time-dependent effects of a-amanitin on blas¬ tocyst formation in the mouse. J. Embryol. Exp. Morphol., 52:193202. 207. Kidder, G. M., and McLachlin, J. R. (1985): Timing of transcription and protein synthesis underlying morphogenesis in preimplantation mouse embryos. Dev. Biol., 112:265-275. 208. van Blerkom, J., Barton, S. C., and Johnson, M. H. (1976): Mo¬ lecular differentiation in the preimplantation mouse embryo. Nature, 259:319-321. 209. Handyside, A. H., and Johnson, M. H. (1978): Temporal and spatial patterns of the synthesis of tissue-specific polypeptides in the preim¬ plantation mouse embryo. J. Embryol. Exp. Morphol., 44:191-199. 210. Howe, C. C., Gmur, R., and Solter, D. (1980): Cytoplasmic and nuclear protein synthesis during in vitro differentiation of murine ICM and embryonal carcinoma cells. Dev. Biol., 74:351-363. 211. Green, M. C. (1981): In: Genetic Variants and Strains of the Lab¬ oratory Mouse, edited by M. C. Green. Gustav Fischer, Stuttgart. 212. McLaren, A. (1976): Genetics of the early mouse embryo. Annu. Rev. Genet., 10:361-388. 213. Magnuson, T., and Epstein, C. J. (1981): Genetic control of very early mammalian development. Biol. Rev., 56:369-408. 214. Magnuson, T. (1983): Genetic abnormalities and early mammalian development. In: Development in Mammals, Vol. 5, edited by M. H. Johnson, pp. 209-249. Elsevier, Amsterdam. 215. Magnuson, T. (1986): Mutations and chromosomal abnormalities: How are they useful for studying genetic control of early mammalian development? In: Experimental Approaches to Mammalian Embry¬ onic Development, edited by J. Rossant and R. A. Pedersen, pp. 437-474. Cambridge University Press, Cambridge. 216. Pedersen, R. A., and Spindle, A. I. (1981): Cellular and genetic analysis of mouse blastocyst development. In: Cellular and Molec¬ ular Aspects of Implantation, edited by S. R. Glasser and D. W. Bullock, pp 91-108. Plenum, New York. 217. Gluecksohn-Waelsch, S. (1979): Genetic control of morphogenetic and biochemical differentiation: lethal albino deletions in the mouse. Cell, 16:225-237. 218. Bennett, D. (1975): The T locus of the mouse. Cell, 6:441-454. 219. Bennett, D. (1980): The T-complex in the mouse: an assessment after 50 years of study. Harvey Led., 74:1-21. 220. Silvers, L. M. (1985): Mouse t haplotypes. Annu. Rev. Genet., 19:179-208. 221. Lewis, S. E. (1978): Developmental analysis of lethal effects of homozygosity for the c25" deletion in the mouse. Dev. Biol., 65:553557. 222. Nadijcka, M. D., Hillman, N., and Gluecksohn-Waelsch, S. (1979): Ultrastructural studies of lethal c^/c25" mouse embryos. J. Embryol. Exp. Morphol., 52:1-11. 223. Hillman, N., and Hillman, R. (1975): Ultrastructural studies of tw32/tw32 mouse embryos. J. Embryol. Exp. Morphol., 33:685-695. 224. Wakasugi, N. (1973): Studies on fertility of DDK mice: reciprocal crosses between DDK and C57BL/6J strains and experimental trans¬ plantation of the ovary. J. Reprod. Fertil., 33:283-291. 225. Paterson, H. F. (1980): In vivo and in vitro studies on the early embryonic lethal tail-short (7s) in the mouse. J. Exp. Zool., 211 247— 256.

226. Babiarz, B. (1983): Deletion mapping of the Tit complex: evidence for a second region of critical embryonic genes. Dev. Biol., 95:342351. 227. Papaioannou, V. E., and Mardon, H. (1983): Lethal nonagouti (cT): description of a second embryonic lethal at the agouti locus. Dev. Genet., 4:21-29. 228. Papaioannou, V. E., and Gardner, R. L. (1979): Investigation of the lethal yellow Ay/Ay embryo using mouse chimaeras. J. Emryol. Exp. Morphol., 52:153-163. 229. Paterson, H. F. (1979): In vivo and in vitro studies on the early embryonic lethal oligosyndactylism (Os) in the mouse. J. Embryol. Exp. Morphol., 52:115-125. 230. Magnuson, T., and Epstein, C. J. (1984): Oligosyndactyly: a lethal mutation in the mouse that results in mitotic arrest very early in development. Cell, 38:823-833. 231. Spiegelman, M., Artzt, K., and Bennett, D. (1976): Embryological study of a Tit locus mutation (tw73) affecting trophectoderm devel¬ opment. J. Embryol. Exp. Morphol., 36:373-381. 232. Babiarz, B., Garrisi, G. J., and Bennett, D. (1982): Genetic analysis of the tw73 haplotype of the mouse using deletion mutations: evidence for a parasitic lethal mutation. Genet. Res., 39:111-120. 233. Guenet, J., Condamine, H., Gaillard, J., and Jacob, F. (1980): twPa~', twPa "2, twPa~3: three new /-haplotypes in the mouse. Genet. Res., 36:211-217. 234. Lewis, 8. E., Turchin, H. A., and Gluecksohn-Waelsch, S. (1976): The developmental analysis of an embryonic lethal (c6H) in the mouse. J. Embryol. Exp. Morphol., 36:363-371. 235. Nadijcka, M. D., and Hillman, N. (1975): Autoradiographic studies of Fit” mouse embryo. J. Embryol. Exp. Morphol., 33:725-730. 236. Gluecksohn-Schoenheimer, S. (1940): The effect of an early lethal (/°) in the house mouse. Genetics, 25:391-400. 237. Nusslein-Volhard, C., Wieschaus, E., and Kluding, H. (1984): Mu¬ tations affecting the pattern of the larval cuticle of Drosophila melanogaster. I. Zygotic loci on the second chromosome. Wilhelm Roux’s Arch. Dev. Biol., 193:267-282. 238. Sternberg, P. W., and Horvitz, H. R. (1984): The genetic control of cell lineage during nematode development. Annu. Rev. Genet., 18:489-524. 239. Silvers, L. M., Uman, J., Danska, J., and Garrels, J. I. (1983): A diversified set of testicular cell proteins specified by genes within the mouse / complex. Cell, 35:35-45. 240. Copeland, N. G., Jenkins, N. A., and Lee, B. K. (1983): Association of the lethal yellow (Av) coat-color mutation with an ecotropic murine leukemia virus genome. Proc. Natl. Acad. Sci. USA, 80:247-249. 241. Epstein, C. J., Smith, S., Travis, B., and Tucker, G. (1978): Both X-chromosomes function before visible X-chromosome inactivation in female mouse embryos. Nature, 274:500-502. 242. Kaufman, M. H., and Gardner, R. L. (1974): Diploid and haploid mouse parthenogenetic development following in vitro activation and embryo transfer. J. Embryol. Exp. Morphol., 31:635-642. 243. Kaufman, M. H., and Sachs, L. (1975): The early development of haploid and aneuploid parthenogenetic embryos. J. Embryol. Exp. Morphol., 34:645-655. 244. Modlinski, J. A. (1975): Haploid mouse embryos obtained by microsurgical removal of one pronucleus. J. Embryol. Exp Morphol 33:897-905. " 245. Tarkowski, A. K. (1977): In vitro development of haploid mouse embryos produced by bisection of one-cell fertilized eggs. J. Em¬ bryol. Exp. Morphol., 38:187-202. 246. Tarkowski, A. K., and Rossant, J. (1976): Haploid mouse blastocysts developed from bisected zygotes. Nature, 259:663-665. 247. Luthardt, F. W. (1976): Cytogenetic analysis of oocytes and early preimplantation embryos from XO mice. Dev. Biol., 54:73-81. 248. Burgoyne, P. S., and Biggers, J. D. (1976): The consequences of X-dosage deficiency in the germ line: impaired development in vitro of preimplantation embryos from XO mice. Dev. Biol., 51:109-117. 249. Epstein, C. J. (1986): The Consequences of Chromosome Imbalance: Principles, Mechanisms, Models. Cambridge University Press, Lon¬ don. 250. Baranov, V. S. (1983): Chromosomal control of early embryonic development in mice. I. Experiments on embryos with autosomal monosomy. Genetica, 61:165-177. 251. Magnuson, T., Debrot, S., Dimpfl, J., Zweig, A., Zamora, T., and Epstein, C. J. (1985): The early lethality of autosomal monosomy in the mouse. J. Exp. Zool., 236:353-360.

Early Embryogenesis

252. Epstein, C. J., Smith, S. A., Zamora, T., Sawicki, J. A., Magnuson, T. R., and Cox, D. R. (1982): Production of viable adult trisomy 17 diploid mouse chimeras. Proc. Natl. Acad. Sci. USA, 79:43764380. 253. Cox, D. R., Smith, S. A., Epstein, L. B., and Epstein, C. J. (1984): Mouse trisomy 16 as an animal model of human trisomy 21 (Down syndrome): production of viable trisomy 16 diploid mouse chi¬ meras. Dev. Biol., 101:416-424. 254. Epstein, C. J., Smith, S. A., and Cox, D. R. (1984): Production and properties of mouse trisomy 15 «-» diploid chimeras. Dev. Ge¬ net., 4:159-165. 255. Boue, J., Boue, A., and Lazar, P. (1975): Retrospective and pro¬ spective epidemiological studies of 1500 karyotyped spontaneous human abortions. Teratology, 12:11-26. 256. Hassold, T. J., Matsuyama, A., Newlands, J. M., Matsuura, J. S., Jacobs, P. A., Manuel, B., and Tsuei, J. (1978): A cytogenetic study of spontaneous abortions in Hawaii. Ann. Hum. Genet., 41:443^454. 257. Markert, C. L. (1982): Parthenogenesis, homozygosity, and cloning in mammals. J. Hered., 73:390-397. 258. Bode, V. C. (1984): Ethylnitrosourea mutagenesis and the isolation of mutant alleles for specific genes located in the f-region of mouse chromosome 17. Genetics, 108:457. 259. Jenkins, N. A., Copeland, N. G., Taylor, B. A., and Lee, B. K. (1981): Dilute (d) coat colour mutation of DBA/2J mice is associated with the site of integration of an ecotropic MuLV genome. Nature, 293:370-374. 260. Copeland, N. G., Hutchison, K. W., and Jenkins, N. A. (1983): Excision of the DBA ecotropic provirus in dilute coat-color revertants of mice occurs by homologous recombination involving the viral LTRs. Cell, 33:379-387. 261. Jaenisch, R., Harbers, K., Schnieke, A., Lohler, J., Chumakov, I., Jahner, D., Grotkopp, D., and Hoffman, E. (1983): Germline in¬ tegration of Moloney murine leukemia virus at the Mov 13 locus leads to recessive lethal mutation and early embryonic death. Cell, 32:209-216. 262. Schnieke, A., Harbers, K., and Jaenisch, R. (1983): Embryonic lethal mutation in mice induced by retrovirus insertion into the a 1(1) collagen gene. Nature, 304:315-320. 263. Harbers, K., Kuehn, M., Delius, H., and Jaenisch, R. (1984): In¬ sertion of retrovirus into the first intron of a 1 (I) collagen gene leads to embryonic lethal mutation in mice. Proc. Natl. Acad. Sci. USA, 81:1504-1508. 264. Breindle, M., Harbers, K., and Jaenisch, R. (1984): Retrovirusinduced lethal mutation in collagen 1 gene is associated with altered chromatin structure. Cell, 38:9-16. 265. Lohler, J., Timpl, R., and Jaenisch, R. (1984): Embryonic lethal mutation in mouse collagen 1 gene causes rupture of blood vessels and is associated with erythropoietic and mesenchymal cell death. Cell, 38:597-607. 266. Kratochwil, K., Dziadek, M., Lohler, J., Harbers, K., and Jaenisch, R. (1986): Normal epithelial branching morphogenesis in the absence of collagen 1. Dev. Biol., 117:596-606. 267. Reik, W., Weiher, H., and Jaenisch, R. (1985): Replication com¬ petent Moloney leukemia virus carrying a bacterial suppressor tRNA gene: selective cloning of proviral and flanking host sequences. Proc. Natl. Acad. Sci. USA, 82:1141-1145. 268. Wagner, E. F., Covarrubias, L., Stewart, T. A., and Mintz, B. (1983): Prenatal lethalities in mice homozygous for human growth hormone gene sequences integrated in the germ line. Cell, 35:647655. 269. Palmiter, R. D., Wilkie, T. M., Chen, H. Y., and Brinster, R. L. (1984): Transmission distortion and mosaicism in an unusual trans¬ genic mouse pedigree. Cell, 36:869-877. 270. Mahon, K. A., Overbeek, P. A., and Westphal, H. (1986): Dominant pre-natal lethality in a transgenic mouse line is associated with a chromosomal translocation. J. Cell Biol., 103:146a. 271. Woychik, R. P., Stewart, T. A., Davis, L. G., D’Eustachio, P., and Leder, P. (1985): An inherited limb deformity created by insertional mutagenesis in a transgenic mouse. Nature, 318:36—40. 272. Izant, J. G., and Weintraub, H. (1984): Inhibition of thymidine kinase gene expression by anti-sense RNA: a molecular approach to genetic analysis. Cell, 36:1007-1015. 273. Melton, D. (1985): Injected anti-sense RNAs specifically block mes¬ senger RNA translation in vivo. Proc. Natl. Acad. Sci. USA, 82:144-

MS.

/

223

274. Melton, D. A., and Rebogliati, M. R. (1986): Anti-sense RNA injections in fertilized eggs as a test for the function of localized mRNAs. J. Embryol. Exp. Morphol. [Suppl.], 97:211-221. 275. Rosenberg, U. B., Preiss, A., Seifert, E., Jackie, H., and Knipple, D. C. (1985): Production of phenocopies by Kruppel antisense RNA injection into Drosophila embryos. Nature, 313:703-706. 276. Kaufman, M. H. (1981): Parthenogenesis: A system facilitating understanding of factors that influence early mammalian develop¬ ment. In: Progress in Anatomy, Vol. 1, edited by R. J. Harri¬ son and R. L. Holmes, pp. 1-34. Cambridge University Press, London. 277. Mittwoch, U. (1978): Parthenogenesis. J. Med. Genet., 15:165-181. 278. Kaufman, M. H. (1983): Early Mammalian Development: Parthenogenetic Studies. Cambridge University Press, London. 279. Markert, C. L., and Seidel, G. E., Jr. (1981): Parthenogenesis, identical twins and cloning in mammals. In: New Technologies in Animal Breeding, edited by B. G. Brackett, G. E. Seidel, Jr., and S. M. Seidel, pp. 181-200. Academic Press, New York. 280. Modlinski, J. A. (1981): The fate of inner cell mass and trophectoderm nuclei transplanted to fertilized mouse eggs. Nature, 292:342343. 281. Illmensee, K., and Hoppe, P. C. (1981): Nuclear transplantation in Mus musculus: developmental potential of nuclei from preimplan¬ tation embryos. Cell, 23:9-18. 282. Hoppe, P. C., and Illmensee, K. (1982): Full term development after transplantation of parthenogenetic embryonic nuclei into fertilized mouse eggs. Proc. Natl. Acad. Sci. USA, 79:1912-1916. 283. Surani, M. A. H. (1986): Evidences and consequences of differences between maternal and paternal genomes during embryogenesis in the mouse. In: Experimental Approaches to Mammalian Embryonic De¬ velopment, edited by J. Rossant and R. A. Pedersen, pp. 401-435. Cambridge University Press, Cambridge. 284. Surani, M. A. H., Reik, W., Norris, M. L., and Barton, S. C. (1986): Influence of germline modifications of homologous chro¬ mosomes on mouse development. J. Embryol. Exp. Morphol., [Suppl.], 97:123-136. 285. Stevens, L. C. (1975): Teratocarcinogenesis and spontaneous par¬ thenogenesis in mice. In: Developmental Biology of Reproduction, edited by C. L. Markert and J. Papaconstantinou, pp. 13-106. Ac¬ ademic Press, New York. 286. Graham, C. F. (1974): The production of parthenogenetic mam¬ malian embryos and their use in biological research. Biol. Rev., 49:399-422. 287. Kaufman, M. H., Barton, S. C., and Surani, M. A. H. (1977): Normal post-implantation development of mouse parthenogenetic embryos to the forelimb bud stage. Nature (Lond.), 265:53-55. 288. Stevens, L. C., Vamum, D. S., and Eicher, E. M. (1977): Viable chimaeras produced from normal and parthenogenetic mouse em¬ bryos. Nature (Lond.), 269:515. 289. Surani, M. A. H., Barton, S. C., and Kaufman, M. H. (1977): Development to term of chimaeras between diploid parthenogenetic and fertilized embryos. Nature (Lond.), 270:601-602. 290. Markert, C. L. and Petters, R. M. (1977): Homozygous mouse em¬ bryos produced by microsurgery. J. Exp. Zool., 201:295-302. 291. Hoppe, P. C., and Illmensee, K. (1977): Microsurgically produced homozygous-diploid uniparental mice. Proc. Natl. Acad. Sci. USA, 74:5657-5661. 292. Modlinski, J. A. (1980): Preimplantation development of micro¬ surgically obtained haploid and homozygous diploid mouse embryos and effects of pretreatment with cytochalasin B on enucleated eggs. J. Embryol. Exp. Morphol., 60:153-161. 293. McGrath, J., and Solter, D. (1983): Nuclear transplantation in the mouse embryo by microsurgery and cell fusion. Science, 220: 13001303. 294. Mann, J. R., and Lovell-Badge, R. H. (1984): Inviability of parthenogenones determined by pronuclei, not egg cytoplasm. Nature (Lond.), 310:66-67. 295. McGrath, J., and Solter, D. (1984): Completion of mouse embry¬ ogenesis requires both maternal and paternal genomes. Cell, 37:179183. 296. Surani, M. A. H., Barton, S. C., and Norris, M. L. (1984): De¬ velopment of reconstituted mouse eggs suggests imprinting of the genome during gametogenesis. Nature (Lond.), 308:548-550. 297. Anderegg, C., and Markert, C. L. (1986): Successful rescue of microsurgically produced homozygous, uniparental mouse embryos

224

/ Chapter 6

via production of aggregation chimeras. Proc. Natl. Acad. ci. USA, 83:6509-6513. 298. Barton, S. C., Surani, M. A. H., and Norris, M. L. (1984): Role of paternal and maternal genomes in mouse development. Nature (bond.), 311:374-376. 299. Barton, S. C., Adams, C. A., Norris, M. L., and Surani, M. A. H. (1985): Development of gynogenetic and parthenogenetic inner cell mass and trophectoderm tissues in reconstituted blastocysts in the mouse. J Embryol. Exp. Morphol., 90:267-285. 300. McGrath, J., and Solter, D. (1984): Maternal (Thp) lethality in the mouse is a nuclear, noncytoplasmic defect. Nature (Load.), 308:550551. 301. Johnson, D. R. (1974): Hairpin-tail: a case of post-reductional gene action in the mouse egg. Genetics, 76:795-805. 302. Johnson, D. R. (1975): Further observations on the hairpin-tail (Thp) mutation in the mouse. Genet. Res., 24:207-213. 303. McLaren, A. (1979): The impact of pre-fertilization events on post¬ fertilization development in mammals. In: Maternal Effects in De¬ velopment, edited by D. R. Newth, and M. Balls, pp. 287-320. Cambridge University Press, London. 304. Winking, H., and Silver, L. M. (1984): Characterization of a re¬ combinant mouse t haplotype that expresses a dominant lethal ma¬ ternal effect. Genetics, 108:1013-1020. 305. Cattanach, B. M., and Kirk, M. (1985): Differential activity of maternally and paternally derived chromosome regions in mice. Na¬ ture, 315:496-498. 306. Cattanach, B. M. (1986): Parental origin effects in mice. J. Embryol. Exp. Morphol. [Suppl.], 97:137-150. 307. Searle, A. G., and Beechey, C. V. (1978): Complementation studies with mouse translocations. Cytogenet. Cell Genet., 20:282-303. 308. Searle, A. G., and Beechey, C. V. (1985): Non-complementation phenomena and their bearing on non-disjunctional effects. In: Aneuploidy, Aetiology and Mechanisms, edited by V. L. Dellarco, P. E. Voytek, and A. Hollaender, pp. 363-376. Plenum, New York. 309. Jacobs, P. A., Wilson, C., Sprenkle, J. A., Rosenshein, N. B., and Migeon,B. R. (1980): Mechanism of origin of complete hydatidiform moles. Nature (bond.), 286:714-716. 310. Szulman, A. E., and Surti, V. (1984): Complete and partial hyda¬ tidiform moles: cytogenetic and morphological aspects. In: Human Trophoblast and Neoplasms, edited by R. A. Patillo and R. O. Hussa, pp. 135-146. Plenum, New York. 311. Modlinski, J. A. (1975): Haploid mouse embryos obtained by microsurgical removal of one pronucleus. J. Embryol. Exp. Morphol., 33:897-905. 312. McGrath, J., and Solter, D. (1984): Inability of mouse blastomere nuclei transferred to enucleated zygotes to support development in vitro. Science, 226:1317-1319. 313. McGrath, J., and Solter, D. (1986): Nucleocytoplasmic interactions in the mouse embryo. J. Embryol. Exp. Morphol. [Suppl.], 97:277289. 314. Robl, J. M., Gilligan, B., Critser, E. S., and First, N. L. (1986): Nuclear transplantation in mouse embryos: assessment of recipient cell stage. Biol. Reprod., 34:733-739. 315. Willadson, S. M. (1986): Nuclear transplantation in sheep embryos. Nature, 320:63-65. 316. Surani, M. A. H., Barton, S. C., and Norris, M. L. (1986): Nuclear transplantation in the mouse: heritable differences between parental genomes after activation of the embryonic genome. Cell, 45:127136. 317. Lyon, M. F. (1961): Gene action in the X-chromosome of the mouse (Mus musculus L.). Nature, 190:372-373. 318. West, J. D. (1982): X-chromosome expression during mouse embryogenesis. In: Genetic Control of Gamete Production and Func¬ tion, edited by P. G. Crosignani and B. W. Rubin, pp. 49-91. Academic Press, New York. 319. Chapman, V. M. (1986): X-chromosome regulation in oogenesis and early mammalian development. In: Experimental Approaches to Mammalian Embryonic Development, edited by J. Rossant and R. A. Pedersen, pp. 365-398. Cambridge University Press, Cambridge. 320. Van de Berg, J. L., Johnston, P. G., Cooper, D. W., and Robinson, E. S. (1983): X-chromosome inactivation and evolution in marsu¬ pials and other mammals. In: Isozymes: Current Topics in Biological and Medical Research, Vol. 9: Gene Expression and Development, edited by M. C. Ratazzi, J. G. Scandalios, and G. S. Whitt, pp. 201-218. Alan R. Liss, Inc., New York.

321. Epstein, C. J. (1972): Expression of the mammalian X-chromosome before and after fertilization. Science, 175:1467-1468. 322. Migeon, B. R., and Jelalian, K. (1977): Evidence for two active X chromosomes in germ cells of female before meiotic entry. Nature, 269:242-241 323. Kratzer, P. G., and Chapman, V. M. (1981): X chromosome reac¬ tivation in oocytes of Mus caroli. Proc. Natl. Acad. Sci. USA, 78:30933097. 324. Lifschytz, E., and Lindsley, D. L. (1974): Sex chromosome acti¬ vation during spermatogenesis. Genetics, 78:323-331. 325. Kramer, J. M., and Erickson, R. P. (1981): Developmental program of PGK-1 and PGK-2 isozymes in spermatogenic cells of the mouse: specific activities and rates of synthesis. Dev. Biol., 87:37^45. 326. Mukherjee, A. B. (1976): Cell cycle analysis and X-chromosome inactivation in the developing mouse. Proc. Natl. Acad. Sci. USA, 73:1608-1611. 327. Takagi, N. (1974): Differentiation of X-chromosomes in early female mouse embryos. Exp. Cell Res., 86:127-135. 328. Adler, D. A., West, J. D., and Chapman, V. M. (1977): Expression of a-galactosidase in preimplantation mouse embryos: implications for X-chromosome inactivation. Nature, 267:838-839. 329. Monk, M., and Kathuria, H. (1977): Dosage compensation for an X-linked gene in preimplantation mouse embryos. Nature, 270:599601. 330. Monk, M., and Harper, M. (1978): X-chromosome activity in preim¬ plantation mouse embryos from XX and XO mothers. J. Embryol. Exp. Morphol., 46:53-64. 331. Kratzer, P. G., and Gartler, S. M. (1978): HGPRT activity changes in preimplantation mouse embryos. Nature, 274:503-504. 332. Epstein, C. J., Travis, B., Tucker, G., and Smith, S. (1978): The direct demonstration of an X-chromosome dosage effect prior to inactivation. In: Genetic Mosaics and Chimeras in Mammals, edited by L. B. Russell, pp. 261-267. Plenum, New York. 333. Gardner, R. L., and Lyon, M. F. (1971): X-chromosome inactivation studies by injection of a single cell into the mouse blastocyst. Nature, 231:385-386. 334. Kratzer, P. G., and Gartler, S. M. (1978): Hypoxanthine guanine phosphoribosyl transferase expression in early mouse development. In: Genetic Mosaics and Chimeras in Mammals, edited by L. B. Russell, pp. 247-260. Plenum, New York. 335. Takagi, N., Sugawara, O., and Sasaki, M. (1982): Regional and temporal changes in the pattern of X-chromosome replication during the early post-implantation development of the female mouse. Chro¬ mosoma (Berl.), 85:275-286. 336. Sugawara, O., Takagi, N., and Sasaki, M. (1983): Allocyclic early replicating X chromosome in mice: genetic inactivity and shift into a later replication in early embryogenesis. Chromosoma (Berl.), 88:133-138. 337. Monk, M., and Harper, M. I. (1979): Sequential X-chromosome inactivation coupled with cellular differentiation in early mouse em¬ bryos. Nature, 281:311-313. 338. West, J. D., Frels, W. I., Chapman, V. M., and Papaioannou, V. E. (1977): Preferential expression of the maternally derived Xchromosome in the mouse yolk sac. Cell, 12:873-882. 339. Frels, W. I., Rossant, J., and Chapman, V. M. (1979): Maternal X chromosome expression in mouse chorionic ectoderm. Dev Genet 1:123-132. 340. Frels, W. I., and Chapman, V. (1980): Expression of the maternally derived X chromosome in the mural trophoblast of the mouse. J. Embryol. Exp. Morphol., 56:179-190. 341. Papaioannou, V. E., West, D. D., Bucher, T., and Linke, I. M. (1981): Non-random X-chromosome expression early in mouse de¬ velopment. Dev. Genet., 2:305-315. 342. McMahon, A., and Monk, M. (1983): X-chromosome activity in female mouse embryos heterozygous for Pgk-1 and Searle’s trans¬ location, T(X;16)16H. Genet. Res. Camb., 41:69-83. 343. Gartler, S. M., and Riggs, A. D. (1983): Mammalian X-chromosome inactivation. Annu. Rev. Genet., 17:155-190. 344. Lyon, M. F. (1974): Mechanisms and evolutionary origins of variable X-chromosome activity in mammals. Proc. R. Soc. Lond [Biol I 187:243-268. 345. Cattanach, B. M. (1975): Control of chromosome inactivation. Annu. Rev. Genet., 9:1-18. 346. Razin, A., Cedar, H., and Riggs, A. (editors) (1984): DNA Methylation. Springer-Verlag, Berlin.

Early Embryogenesis /

347. Doerfler, W. (1983): DNA methylation and gene activity. Annu. Rev. Biochem., 52:93-124. 348. Wolf, S. F., Jolly, D. J., Lunnen, K. D., Friedman, T., and Migeon, B. R. (1984): Methylation of the hypoxanthine phosphoribosyl trans¬ ferase locus on the human X-chromosome: implications for Xchromosome inactivation. Proc. Natl. Acad. Sci. USA, 81:28062810. 349. Manes, C., and Menzel, P. (1981): Demethylation of CpG sites in DNA of early rabbit trophoblast. Nature, 293:589-590. 350. Chapman, V., Forrester, L., Sanford, J., Hastie, N., and Rossant, J. (1984): Cell lineage-specific undermethylation of mouse repetitive DNA. Nature, 307:284-286. 351. Razin, A., Webb, C., Szyf, M., Ysraeli, J., Rosenthal, A., NavehMany, T., Sciaky-Gallili, N., and Cedar, H. (1984): Variations in DNA methylation during mouse cell differentiation in vivo and in vitro. Proc. Natl. Acad. Sci. USA, 81:2275-2279. 352. Young, P. R., and Tilghman, S. M. (1984): Induction of a-fetoprotein synthesis by differentiation of teratocarcinoma cells is ac¬ companied by a genome-wide loss of DNA methylation. Mol. Cell. Biol., 4:898-907. 353. Rossant, J., Sanford, J. P., Chapman, V. M., and Andrews, G. K. (1986): Undermethylation of structural gene sequences in extraembryonic lineages of the mouse. Dev. Biol., 117:567-573. 354. Chapman, V., Kartzer, P. G., Siracusa, L. D., Quarantillo, B. A., Evans, R., and Liskay, R. M. (1982): Evidence from DNA modi¬ fication in the maintenance of X-chromosome inactivation of adult mouse tissues. Proc. Natl. Acad. Sci. USA, 79:5357-5361. 355. Kratzer, P. G., Chapman, V. M., Lambert, H., Evans, R. E., and Liskay, R. M. (1983): Differences in the DNA of the inactive Xchromosomes of fetal and extraembryonic tissues of mice. Cell, 33:37-42. 356. Krumlauf, R., Hammer, R. E., Brinster, R., Chapman, V. M., and Tilghman, S. M. (1985): Regulated expression of a-fctoprotein genes in transgenic mice. Cold Spring Harbor Symp. Quant. Biol., 50:371378. 357. Jahner, D., Stuhlmann, H., Stewart, W. L., Harbers, K., Lohler, J., Simon, J., and Jaenisch, R. (1982): De novo methylation and expression of retroviral genomes during mouse embryogenesis. Na¬ ture, 298:623-628. 358. Pellicer, A., Robins, D., Wold, B., Sweet, R., Jackson, J., Lowy, I., Roberts, J. M., Sim, G. K., Silverstein, S., and Axel, R. (1980): Altering genotype and phenotype by DNA-mediated gene transfer. Science, 29:1414-1422. 359. Jaenisch, R. (1974): Infection of mouse blastocysts with SV40 DNA: Normal development of the infected embryos and persistence of SV40 specific DNA sequences in the adult animals. Cold Spring Harbor Symp. Quant. Biol., 39:375-380. 360. Jaenisch, R., and Mintz, B. (1974): Simian virus 40 DNA sequences in DNA of healthy adult mice derived from preimplantation blas¬ tocysts injected with viral DNA. Proc. Natl. Acad. Sci. USA, 71:12501254. 361. Brinster, R. L., Chen, H. Y., Trumbauer, M. E., Denear, A. W., Warren, R., and Palmiter, R. D. (1981): Somatic expression of herpes thymidine kinase in mice following injection of a fusion gene into eggs. Cell, 27:223-231. 362. Costantini, F., and Lacy, E. (1981): Introduction of a rabbit (3-globin gene into the mouse germ line. Nature, 294:92-94. 363. Wagner, E. F., Stewart, T. A., and Mintz, B. (1981): The human P-globin gene and a functional viral thymidine kinase gene in de¬ veloping mice. Proc. Natl. Acad. Sci. USA, 78:5016-5020. 364. Wagner, T. E., Hoppe, P. C., Jollick, J. D., Scholl, D. R., Hodinka, R. L., and Gault, J. B. (1981): Microinjection of a rabbit p-globin gene into zygotes and its subsequent expression in adult mice and their offspring. Proc. Natl. Acad. Sci. USA, 78:6376-6380. 365. Hammer, R., Pursel, V. G., Rexroad, C. E., Jr., Wall, R. J., Bolt, D. J., Ebert, K. M., Palmiter, R. D., and Brinster, R. L. (1985): Production of transgenic rabbits, sheep, and pigs by microinjection. Nature, 315:680-683. 366. Gordon, J. W., and Ruddle, F. H. (1985): DNA mediated genetic tranformation of mouse embryos and bone marrow—a review. Gene, 33:121-136. 367. Palmiter, R. D., and Brinster, R. L. (1985): Transgenic mice. Cell, 14:343-345. 368. Brinster, R. L., Chen, H. Y., Trumbauer, M., Yagle, M. K., and Palmiter, R. D. (1985): Factors affecting the efficiency of introducing

369.

370.

371.

372. 373.

374.

375. 376.

377.

378.

379.

380.

381.

382.

383.

384.

385.

386.

387.

388.

225

foreign DNA into mice by microinjecting eggs. Proc. Natl. Acad. Sci. USA, 82:4438^4442. Wagner, E. F., Ruther, U., and Stewart, C. L. (1984): Introducing genes into mice and into embryonal carcinoma stem cells. In: The Impact of Gene Transfer Techniques in Eucaryotic Cell Biology, edited by J. S. Schell and P. Starlinger, pp. 127-133. SpringerVerlag, Berlin. Wagner, E. F., Ruther, U., and Stewart, C. L. (1986): Gene transfer into mouse stem cells. In: Dahlem Workshop Report: Biotechnology: Potentials and Limitations. Life Sciences Report No. 35, edited by S. Silver, pp. 185-196. Springer-Verlag, Cambridge. Wagner, E. F., and Stewart, C. L. (1986): Integration and expression of genes introduced into mouse embryos. In: Experimental Ap¬ proaches to Mammalian Embryonic Development, edited by J. Ros¬ sant and R. A. Pedersen, pp. 509-549. Cambridge University Press, Cambridge. Brinster, R. L., and Palmiter, R. D. (1985): Introduction of genes into the germ line of animals. Harvey Led., 80:1-38. McKnight, G. S., Hammer, R. E., Kuenzel, E. A., and Brinster, R. L. (1983): Expression of the chicken transferrin gene in transgenic mice. Cell, 34:335-341. Swift, G. H., Hammer, R. E., McDonald, R. J., and Brinster, R. L. (1984): Tissue-specific expression of the rat pancreatic elastase I gene in transgenic mice. Cell, 38:639-646. Shani, M. (1985): Tissue-specific expression of rat myosin lightchain 2 gene in transgenic mice. Nature, 314:283-286. Krumlauf, R., Hammer, R., Tilghman, S., and Brinster, R. L. (1985): Developmental regulation of alphafoetoprotein genes in transgenic mice. Mol. Cell. Biol., 5:1639-1648. Palmiter, R. D., Chen, H. Y., and Brinster, R. L. (1982): Differential regulation of metallothionein-thymidine kinase fusion gene in trans¬ genic mice and their offspring. Cell, 29:701-710. Overbeek, P. A., Chepelinsky, A., Khillan, J. S., Piatigorsky, J., and Westphal, H. (1985): Lens-specific expression and develop¬ mental regulation of the bacterial chloramphenicol acetyltransferase gene driven by the murine aA-cystallin promoter in transgenic mice. Proc. Natl. Acad. Sci. USA. 82:7815-7819. Westphal, H., Overbeek, P. A., Khillan, J. S., Chepelinsky, A. B., Schmidt, A., Mahon, K. A., Bernstein, K. E., Piatigorsky, J., and de Crombrugghe, B. (1985): Promoter sequences of murine aA cystallin, murine a2(l) collagen or avian sarcoma virus genes linked to the bacterial CAT gene direct tissue specific patterns of CAT expres¬ sion in transgenic mice. Cold Spring Harbor Symp. Quant. Biol., 50:411-416. Omitz, D. M., Palmiter, R. D., Hammer, R. E., Brinster, R. L., Swift, G. H., and McDonald, J. R. (1985): Specific expression of an elastase-human growth hormone fusion gene in pancreatic acinar cells of transgenic mice. Nature, 313:600-602. Goring, D. R., Rossant, J., Clapoff, S., Breitman, M. L., andTsui, L. -C. (1986): In situ detection of p-galactosidase activity in lenses of transgenic mice harboring a y-crystallinlLac-z hybrid gene. Sci¬ ence 235:456^458. Townes, T. M., Lingrel, J. B., Chen, H. Y., Brinster, R. L., and Palmiter, R. D. (1985): Erythroid specific expression of human (3globin genes in transgenic mice. EMBO J., 4:1715-1723. Palmiter, R. D., Brinster, R. L., Hammer, R. E., Trumbauer, M. E., Rosenfeld, M. G., Bimberg, N. C., and Evans, R. M. (1982): Dramatic growth of mice that develop from eggs microinjected with metallothionein-growth hormone fusion genes. Nature, 300:611-615. Palmiter, R. D., Norstedt, G., Gelinas, R. E., Hammer, R. E., and Brinster, R. L.( 1983): Metallothionein-human GH fusion genes stim¬ ulate growth of mice. Science, 222:809-814. Hammer, R. E., Palmiter, R. D., and Brinster, R. L. (1984): Partial correction of murine hereditary growth disorder by germ line incor¬ poration of a new gene. Nature, 311:65-67. Brinster, R. L., Ritchie, K. A., Hammer, R. E., O’Brien, R. L., Arp, B., and Storb, U. (1983): Expression of a microinjected im¬ munoglobulin gene in the spleen of transgenic mice. Nature, 306:332336. Storb, U., O’Brien, R. L., McMullen, M. D., Gollahon, K. A., and Brinster, R. L. (1984): High expression of cloned immunoglobulin k gene in transgenic mice is restricted to B lymphocytes. Nature, 310:238-241. Grosschedl, R., Weaver, D., Baltimore, D., and Costantini, F. (1984): Introduction of a |x-immunoglobulin gene into the mouse germ line:

226

/ Chapter 6

specific expression in lymphoid cells and synthesis of functional antibody. Cell, 38:647—658. 389. Rusconi, S., and Kohler, G. (1985): Transmission and expression of a specific pair of rearranged immunoglobulin p, and k genes in a transgenic mouse line. Nature, 314:330-334. 390. Frels, W. 1., Bluestone, J. A., Hodes, R. J., Capecchi, M. R., and Singer, D. S. (1985): Expression of a microinjected porcine class I major histocompatibility complex gene in transgenic mice. Science, 228:577-580. 391. Le Meur, ML, Gerlinger, P., Benoist, C., and Mathis, D. (1985): Correcting an immune response deficiency by creating Ea gene trans¬ genic mice. Nature, 316:38^42. 392. Yamamura, K., Kikutani, H., Folson, V., Clayton, L. K., Kimoto, M., Akira, S., Kashiwamura, S., Tonegawa, S., and Kishimoto, R. (1985): Functional expression of a microinjected Eda gene in C57BL/6 transgenic mice. Nature, 316:67-69. 393. Pinkert, C. A., Widera, G., Cowing, C., Heber-Katz, E., Palmiter, R. D., Flavell, R. A., and Brinster, R. L. (1985): Tissue-specific, inducible and functional expression of the Eda MHC class II gene in transgenic mice. EMBO J., 4:2225-2230. 394. Brinster, R. L., Chen, H. Y., Messing, A., van Dyke, T., Levine, A. J., and Palmiter, R. D. (1984): Transgenic mice harboring SV40 T-antigen genes develop characteristic brain tumors. Cell, 37:367379. 395. Palmiter, R. D., Chen, H. Y., Messing, A., and Brinster, R. L. (1985): SV40 enhancer and large-T antigen are instrumental in de¬ velopment of choroid plexus tumours in transgenic mice. Nature, 36:457—460. 396. Messing, A., Chen, H. Y., Palmiter, R. D., and Brinster, R. L. (1985): Peripheral neuropathies, hepatocellular carcinomas, and islet cell adenomas in transgenic mice. Nature, 316:461-463. 397. Stewart, T. A., Pattengale, P. K., and Leder, P. (1984): Spontaneous mammary adenocarcinomas in transgenic mice that carry and express MTV/wyc fusion genes. Cell, 23:627-637. 398. Hanahan, D. (1985): Heritable formation of pancreatic [3-cell tumors in transgenic mice expressing recombinant insulin/SV40 oncogenes. Nature, 315:115-122. 399. Babinet, C., Farza, H., Morello, D., Hadchouel, M., and Pourcel, C. (1985): Specific expression of hepatitis B surface antigen (HBsAg) in transgenic mice. Science, 230:160-163. 400. Chisari, F. V., Pinkert, C. A., Milich, D. R., Filippi, P., McLachlan, A., Palmiter, R. D., and Brinster, R. L. (1985): A transgenic mouse model of the chronic hepatitis B surface antigen carrier state. Science 230:1157-1160. 401. Rusconi, S. (1984): Gene transfer in living organisms. In: The Impact of Gene Transfer Techniques in Eucaryotic Cell Biology, edited by J. S. Schell and P. Starlinger, pp. 134-152. Springer-Verlag, Berlin. 402. Stewart, T. A., Wagner, E. F., and Mintz, B. (1982): Human (3globin gene sequences injected into mouse eggs, retained in adults, and transmitted to progeny. Science, 217:1046-1048. 403. Chada, K., Magram, J., Raphael, K., Radice, G., Lacy, E., and Costantini, F. (1985): Specific expression of a foreign |3-globin gene in erythroid cells of transgenic mice. Nature, 314:377-380. 404. Magram, J., Chada, K., and Costantini, F. (1985): Developmental regulation of a cloned adult (3-globin gene in transgenic mice. Nature, 315:338-340. 405. Lacy, E., Roberts, S., Evans, E. P., Burtenshaw, M. D., and Cos¬ tantini, F. (1983): A foreign p-globin gene in transgenic mice: in¬ tegration at abnormal chromosomal positions and expresson in in¬ appropriate tissues. Cell, 34:343-358. 406. Wagner, E. F., Covarrubias, L., Stewart, T. A., and Mintz, B. (1983): Prenatal lethalities in mice homozygous for human growth hormone gene sequences integrated in the germ line. Cell, 35:647655. 407. Hammer, R. E., Brinster, R. L., Rosenfeld, M. G., Evans, R. M., and Mayo, K. E. (1985): Expression of human growth factor in transgenic mice results in increased somatic growth. Nature, 315:413416. 408. Stout, J. T., Chen, H. Y., Brennand, J., Caskey, C. T., and Brinster, R. L. (1985): Expression of human HPRT in the central nervous system of transgenic mice. Nature, 317:250-252. 409. Low, M. J., Hammer, R. E., Goodman, R. H., Habener, J. F., Palmiter, R. D., and Brinster, R. L. (1985): Tissue-specific posttranslational processing of pre-prosomatostatin encoded by a metal-

lothionein-somatostatin fusion gene in transgenic mice. Cell, 41:211219. 410. Ross, S. R., and Solter, D. (1985): Glucocorticoid regulation of mouse mammary tumor virus sequences in transgenic mice. Proc. Natl. Acad. Sci. USA, 82:5880-5884. 411. Harbers, K., Jahner, D., and Jaenisch, R. (1981): Microinjection of cloned retroviral genomes into mouse zygotes: integration and expres¬ sion in the animal. Nature, 293:540-542. 412. Stewart, C. L., Harbers, K., Jahner, D., and Jaenisch, R. (1983): X chromosome-linked transmission and expression of retroviral ge¬ nomes microinjected into mouse zygotes. Science, 221:760-762. 413. Jaenisch, R., and Mintz, B. (1974): Simian virus 40 DNA sequences in DNA of healthy adult mice derived from preimplantation blas¬ tocysts injected with viral DNA. Proc. Natl. Acad. Sci. USA, 71:1250— 1254. 414. Small, J. A., Blair, D. G., Showalter, S. D., and Scangos, G. A. (1985): Analysis of a transgenic mouse containing SV40 and \-myc sequences. Mol. Cell. Biol., 5:642-648. 415. Wirak, D. O., Chalifour, L. E., Wassarman, P. M., Muller, W. J., Hassell, J. A., and De Pamphilis, M. L. (1985): Sequence-dependent DNA replication in preimplantation mouse embryos. Mol. Cell. Biol., 5:2924-2935. 416. Rassoulzadegan, M., Leopold, P., Vailly, J., and Cuzin, F. (1986): Germ line transmission of autonomous genetic elements in transgenic mouse strains. Cell, 46:513-519. 417. Kelly, F., and Condamine, H. (1982): Tumor viruses and early mouse embryos. Biochim. Biophys. Acta, 651:105-141. 418. Jaenisch, R., Fan, H., and Croker, B. (1975): Infection of preim¬ plantation mouse embryos and of newborn mice with leukaemia virus: tissue distribution of viral DNA and RNA and leukemogenesis in the adult animal. Proc. Natl. Acad. Sci. USA, 72:4008-4012. 419. Jaenisch, R., and Jahner, D. (1984): Methylation, expression and chromosomal position of genes in mammals. Biochim. Biophys. Acta, 782:1-9. 420. Willison, K., Babinet, C., Boccara, M., and Kelly, F. (1983): In¬ fection of preimplantation mouse embryos with simian virus 40. In: Cold Spring Harbor Conference, Vol. 10, edited by L. M. Silver, G. R. Martin, and S. Strickland, pp. 307-317. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 421. Jaenisch, R., Jahner, D., Nobis, P., Simon, I., Lohler, J., Harbers, K., and Grotkopp, D. (1981): Chromosomal position and activation of retroviral genomes inserted into the germ line of mice. Cell, 24:519-529. 422. Rubinstein, J. L. R., Nicolas, J.-F., and Jacob, F. (1984): Construc¬ tion of a retrovirus capable of transducing and expressing genes in multipotential embryonic cells. Proc. Natl. Acad. Sci. USA, 81:7137— 7140. 423. Wagner, E. F., Vanek, M., and Vennstrom, B. (1985): Transfer of genes into embryonal carcinoma cells by retrovirus infection: efficient expression from an internal promoter. EMBO J., 4:663-669. 424. Stewart, C. L., Vanek, M., and Wagner, E. F. (1985): Expression of foreign genes from retroviral vectors in mouse teratocarcinoma chimaeras. EMBO J., 4:3701-3709. 425. Stewart, C. L., Ruther, U., Garber, C., Vanek, M., and Wagner, E. F. (1986): The expression of retroviral vectors in murine stem cells and transgenic mice. J. Embryol. Exp. Morphol. [Suppl / 97:263-275. 426. Robertson, E. J., and Bradley, A. (1986): Production of permanent cell lines from early embryos and their use in studying developmental problems. In: Experimental Approaches to Mammalian Embryonic Development, edited by J. Rossant and R. A. Pedersen, pp. 475508. Cambridge University Press, Cambridge. 427. Soriano, P., Cone, R. D., Mulligan, R. C., and Jaenisch, R. (1986): Tissue-specific and ectopic expression of genes introduced into trans¬ genic mice by retroviruses. Science, 234:1409-1413. 428. Bradley, A., Evans, M., Kaufman, M. H., and Robertson, E. (1984): Formation of germ line chimaeras from embryo derived teratocar¬ cinoma cell lines. Nature, 309:255-256. 429. Gossler, A., Doetschman, T., Korn, R., Serfling, E., and Kemler, R. (1986): Transgenesis via blastocyst-derived embryonic stem cell lines. Proc. Natl. Acad. Sci. USA, 83:9065-9069. 430. Robertson, E., Bradley, A., Kuehn, M., and Evans, M. (1986): Germ-line transmission of genes introduced into cultured pluripotential cells by retroviral vector. Nature, 323:445^148.

Early Embryogenesis /

431. Mulligan, R. C., and Berg, P. (1981): Selection for animal cells that express the Escherichia coli gene coding for xanthine-guanine phosphoribosyl transferase. Proc. Natl. Acad. Sci. USA, 78:2072-2076. 432. Southern, P. J., and Berg, P. (1982): Transformation of mammal¬ ian cells to antibiotic resistance with a bacterial gene under control of the SV40 early region promoter. J. Mol. Appl. Genet., 1:327341. 433. Johnson, M. H. (1981): The molecular and cellular basis of preim¬ plantation mouse development. Biol. Rev., 56:463-498. 434. Dziadek, M., and Adamson, E. D. (1978): Localisation and synthesis of alpha-foetoprotein in post-implantation mouse embryos. J. Embryol. Exp. Morphol., 43:289-313. 435. Dziadek, M. (1978): Modulation of alphafoetoprotein synthesis in the early postimplantation mouse embryo. J. Embryol. Exp. Mor¬ phol., 46:135-146. 436. Albrechtsen, R., and Norgaard-Pederson, B. (1978): Immunofluorescent localisation of alpha-fetoprotein synthesis in the endodermal sinus of rat placenta. Scand. J. Immunol., 8:193-199. 437. Ruoslahti, E., and Seppala, M. (1979): a-Fetoprotein in cancer and fetal development. Adv. Cancer Res., 29:275-346. 438. Andrews, G. K., Janzen, R. G., and Tamaski, T. (1982): Stability of a-fetoprotein messenger RNA in mouse yolk sac. Dev. Biol., 89:111-116. 439. Tilghman, S. M., and Belayew, A. (1982): Transcriptional control of the murine albumin/a-fetoprotein locus during development. Proc. Natl. Acad. Sci. USA, 79:5254-5257. 440. Sellem, C. H., Frain, M., Erdos, T., and Sala-Trepat, J. M. (1984): Differential expression of albumin and a-fetoprotein genes in fetal tissues of mouse and rat. Dev. Biol., 102:51-60. 441. Krumlauf, R., Hammer, R. E., Tilghman, S. M., and Brinster, R. L. (1985): Developmental regulation of a-fetoprotein genes in transgenic mice. Mol. Cell. Biol., 5:1639-1648. 442. Hammer, R. E., Krumlauf, R., Camper, S., Brinster, R. L., and Tilghman, S. M. (1986): The regulation of a-fetoprotein minigene expression in the germline of mice. J. Embryol. Exp. Morphol. [Suppl.], 97:257-262. 443. Renfree, M. B., and McLaren, A. (1974): Foetal origin of transferrin in mouse amniotic fluid. Nature, 252:159-160. 444. Adamson, E. D. (1982): The location and synthesis of transferrin in mouse embryos and teratocarcinoma cells. Dev. Biol., 91:227-234. 445. Meek, J., and Adamson, E. D. (1985): Transferrin in fetal and adult mouse tissues: synthesis, storage, and secretion. J. Embryol. Exp. Morphol., 86:205-218. 446. Andrews, G. K., Adamson, E. D., and Gedamu, L. (1984): The ontogeny of expression of murine metallothionein: comparison with the a-fetoprotein gene. Dev. Biol., 103:294-303. 447. Hopkins, B., Sharpe, C. R., Baralle, F. E., and Graham, C. F. (1986): Organ distribution of apolipoprotein gene transcripts in 6-12 week postfertilization human embryos. J. Embryol. Exp. Morphol., 97:177-187. 448. Adamson, E. D. (1986): Cell-lineage-specific gene expression in development. In: Experimental Approaches to Mammalian Embry¬ onic Development, edited by J. Rossant and R. A. Pedersen, pp. 321-364. Cambridge University Press, Cambridge. 449. Trelstad, R. L. (editor) (1984): The Role of Extracellular Matrix in Development, 42nd Symposium of the Society for Developmental Biology. Alan R. Liss, New York. 450. Wartiovaara, J., Leivo, I., and Vaheri, A. (1979): Expression of the cell surface-associated glycoprotein, fibronectin, in the early mouse embryo. Dev. Biol., 69:247-257. 451. Semoff, S., Hogan, B. L. M., and Hopkins, C. R. (1982): Local¬ isation of fibronectin, laminin-entactin and entactin in Reichert’s membrane by immuno-electron microscopy. EMBO J., 1:1171-1175. 452. Adamson, E. D., and Ayers, S. E. (1979): The localization and synthesis of collagen in mouse embryos. Cell, 16:953-965. 453. Leivo, I., Vaheri, A., Timpl, R., and Wartiovaara, J. (1980): Ap¬ pearance and distribution of collagens and laminin in the early mouse embryo. Dev. Biol., 76:100-114. 454. Cooper, A. R., and MacQueen, H. A. (1983): Subunits of laminin are differentially synthesized in mouse eggs and early embryos. Dev. Biol., 96:467-471. 455. Wu, T. C., Wan, Y.-J., Chung, A. E., and Damjanov, I. (1983): Immunohistochemical localization of entactin and laminin in mouse embryos and fetuses. Dev. Biol., 100:496-505.

227

456. Sherman, M. I., Gay, R., Gay, S., and Miller, E. J. (1980): As¬ sociation of collagen with preimplantation and peri-implantation mouse embryos. Dev. Biol., 74:470^178. 457. Schnieke, A. A., Harbers, K., and Jaenisch, R. (1983): Embryonic lethal mutation in mice induced by retrovirus insertion into the a 1(1) collagen gene. Nature, 304:315-320. 458. Hogan, B. L. M., Cooper, A. R., and Kurkinen, M. (1980): In¬ corporation into Reichert’s membrane of laminin-like extracellular proteins synthesized by parietal endoderm cells of the mouse embryo. Dev. Biol., 80:289-300. 459. Hogan, B. L. M., Taylor, A., and Cooper, A. R. (1982): Murine parietal endoderm cells synthesize heparan sulphate and 170 K and 145 K sulphated glycoproteins as components of Reichert’s mem¬ brane. Dev. Biol., 90:210-214. 460. Jacob, F. (1979): Cell surface and early stages of mouse embryogenesis. Curr. Top. Dev. Biol., 13:117-135. 461. Sol ter, D., and Knowles, B. B. (1979): Developmental stage-specific antigens during mouse embryogenesis. Curr. Top. Dev. Biol., 13:139165. 462. Wiley, L. M. (1979): Early embryonic cell surface antigens as de¬ velopmental probes. Curr. Top. Dev. Biol., 13:167-197. 463. Johnson, L. V., and Calarco, P. G. (1980): Mammalian preimplan¬ tation development: the cell surface. Anat. Rec., 196:201-219. 464. Johnson, M. H. (1981): Membrane events associated with the gen¬ eration of a blastocyst. Int. Rev. Cytol. [Suppl.], 12:1-37. 465. Richa, J., and Solter, D. (1986): Role of cell surface molecules in early mammalian development. In: Experimental Approaches to Mammalian Embryonic Development, edited by J. Rossant and R. A. Pedersen, pp. 293-320. Cambridge University Press, Cambridge. 466. Webb, C. A. G. (1983): Glycoproteins on gametes and early em¬ bryos. In: Development in Mammals, Vol. 5, edited by M. H. John¬ son, pp. 155-185. Elsevier, Amsterdam. 467. Ducibella, T., and Anderson, E. (1975): Cell shape and membrane changes in the eight-cell mouse embryo: prerequisites for morpho¬ genesis of the blastocyst. Dev. Biol., 47:45-58. 468. Hyafil, F., Morello, D., Babinet, C., and Jacob, F. (1980): A cell surface glycoprotein involved in the compaction of embry¬ onal carcinoma cells and cleavage stage embryos. Cell, 21:927934. 469. Yoshida-Noro, C., Suzuki, N., and Takeichi, M. (1984): Molecular nature of the calcium-dependent cell-cell adhesion in mouse terato¬ carcinoma and embryonic cells studied with a monoclonal antibody. Dev. Biol., 101:19-27. 470. Damsky, C. H., Richa, J., Solter, D., Knudsen, K., and Buck, C. A. (1983): Identification and purification of a cell surface glyco¬ protein mediating intercellular adhesion in embryonic and adult tissue. Cell, 34:455^166. 471. Richa, J., Damsky, C. H., Buck, C. A., Knowles, B. B., and Solter, D. (1985): Cell surface glycoproteins mediate compaction, trophoblast attachment and endoderm formation during early mouse de¬ velopment. Dev. Biol., 108:513-521. 472. Gallin, W. J., Edelman, G., and Cunningham, B. (1983): Charac¬ terization of L-CAM, a major cell adhesion molecule from embryonic liver cells. Proc. Natl. Acad. Sci. USA, 80:1038-1042. 473. Ogou, S.-I., Okada, T. S., and Takeichi, M. (1982): Cleavage stage mouse embryos share a common cell adhesion system with terato¬ carcinoma cells. Dev. Biol., 92:521-528. 474. Kimber, S. J., and Surani, M. A. H. (1982): Spreading of blastomeres from eight-cell mouse embryos on lectin coated beads. J. Cell Sci., 56:191-206. 475. Johnson, L. V. (1985): Wheat germ agglutinin induces compactionand cavitation-like events in 2-cell mouse embryos. Dev. Biol., 113:19. 476. Zvzack, J. S., and Tasca, R. J. (1985): Lectin-induced blockage of developmental processes in preimplantation mouse embryos in vitro. Gamete Res. 12:275-290. 477. Brownell, A. G. (1977): Cell surface carbohydrates of preimplan¬ tation embryos as assessed by lectin binding. J. Supramol. Struct., 7:223-234. 478. Sato, M., and Muramatsu, T. (1985): Reactivity of five A-acetylgalactosamine-recognizing lectins with preimplantation embryos, early postimplantation embryos, and teratocarcinoma cells of the mouse. Differentiation, 29:29-38.

228

/ Chapter 6

479. Solter, D., and Knowles, B. B. (1978): Monoclonal antibody defining a stage-specific mouse embryonic antigen (SSEA-1). Proc. Natl. Acad. Sci. USA, 75:5565-5569. 480. Solter, D., and Knowles, B. B. (1979): Developmental stage-specific antigens during mouse embryogenesis. Curr. Top. Dev. Biol., 13:139— 165. 481. Fox, N., Damjanov, I., Martinez-Hemandez, A., Knowles, B. B., and Solter, D. (1981): Immunohistochemical localization of the early embryonic antigen (SSEA-1) in postimplantation mouse embryos, and fetal and adult tissues. Dev. Biol., 83:391-398. 482. Rastan, S., Thorpe, S. J., Scudder, P., Brown, S. Gooi, H. C., and Feizi, T. (1985): Cell interactions in preimplantation embryos: evi¬ dence for involvement of saccharides of the poly-N-acetyllactosamine series. J. Embryol. Exp. Morphol., 87:115-128. 483. Knowles, B. B., Rappaport, J., and Solter, D. (1982): Murine em¬ bryonic antigen (SSEA-1) is expressed on human cells and structur¬ ally related human blood group antigen I is expressed on mouse embryos. Dev. Biol., 93:54-58. 484. Kapadia, A., Feizi, T., and Evans, M. J. (1981): Changes in the expression and polarization of blood group I and i antigens in post¬ implantation embryos and teratocarcinomas of mouse associated with cell differentiation. Exp. Cell Res., 131:185-195. 485. Shevinsky, L. H., Knowles, B. B., Damjanov, I., and Solter, D. (1982): Monoclonal antibody to murine embryos defines a stagespecific embryonic antigen expressed on mouse embryos and human teratocarcinoma cells. Cell, 30:697-705. 486. Kannagi, R., Cochran, N. A., Ishigami, F., Hakomori, S.-I., An¬ drews, P. W., Knowles, B. B., and Solter, D. (1983): Stage-specific embryonic antigens (SSEA-3 and -4) are epitopes of a unique globoseries ganglioside isolated from human teratocarcinoma cells. EMBO J., 2:2355-2361. 487. Fox, N. W., Damjanov, I., Knowles, B. B., and Solter, D. (1984): Stage-specific embryonic antigen 3 as a marker of visceral extraembryonic endoderm. Dev. Biol., 103:263-266. 488. Willison, K. R., Karol, R. A., Suzuki, A., Kundu, S. K., and Marcus, D. M. (1982): Neutral glycolipid antigens as developmental markers of mouse teratocarcinomas and early embryos: an immu¬ nologic and chemical analysis. J. Immunol., 129:603-609. 489. Willison, K. R., and Stem, P. L. (1978): Expression of a Forssman antigenic specificity in the preimplantation mouse embryo. Cell, 14:785-793. 490. Muramatsu, H., and Muramatsu, T. (1983): A fucosyltransferase in teratocarcinoma cells. Decreased activity accompanying differentia¬ tion to parietal endoderm cells. FEBS Lett., 163:181-184. 491. Shur, B. D. (1983): Embryonal carcinoma cell adhesion: the role of surface galactosyltransferase and its 90K lactosaminoglycan sub¬ strate. Dev. Biol., 99:360-372. 492. Sato, M., Muramatsu, T., andGerger, E. G. (1984): Immunological detection of cell surface galactosyltransferase in preimplantation mouse embryos. Dev. Biol., 102:514-518. 493. Shur, B. D., Oettgen, P., and Bennett, D. (1979): UDP-galactose inhibits blastocyst formation in the mouse. Dev. Biol., 73:178-181. 494. Mulnard, J., and Huygens, R. (1978): Ultrastructural localization of non-specific alkaline phosphatase during cleavage and blastocyst for¬ mation in the mouse. J. Embryol. Exp. Morphol., 44:121-131. 495. Johnson, L. V., Calarco, P. G., and Siebert, M. L. (1977): Alkaline phosphatase activity in the preimplantation mouse embryo. J. Em¬ bryol. Exp. Morphol., 40:83-89. 496. Eddy, E. M. (1975): Germ plasm and the differentiation of the germ cell line. Int. Rev. Cytol., 43:229-280. 497. Eddy, E. M., Clark, J. M., Gong, D., and Fenderson, B. A. (1981): Origin and migration of primordial germ cells in mammals. Gamete Res., 4:333-362. 498. Vorbrodt, A., Konwinski, M., Solter, D., and Koprowski, H. (1977): Ultrastructural cytochemistry of membrane-bound phosphatases in preimplantation mouse embryos. Dev. Biol., 55:117-134. 499. Borland, R. M., Biggers, J. D., and Lechene, C. P. (1976): Kinetic aspects of rabbit blastocoele fluid accumulation: an application of electron probe microanalysis. Dev. Biol., 50:201-211. 500. Benos, D. J. (1981): Ouabain binding to preimplantation rabbit blas¬ tocysts. Dev. Biol., 83:69-78. 501. Benos, D. J., Biggers, J. D., Balaban, R. S., Mills, J. W., and Overstrom, E. G. (1985): Developmental aspects of sodium depen¬ dent transport processes of preimplantation rabbit embryos. In: Reg¬

ulation and Development of Membrane Transport Processes, edited by J. S. Graves, pp. 211-235. Wiley, New York. 502. DiZio, S. M., and Tasca, R. J., (1977): Sodium-dependent aminoacid transport in preimplantation mouse embryos. III. Na+-K+ATPase-linked mechanism in blastocysts. Dev. Biol., 59:198-205. 503. Wiley, L. M. (1984): Cavitation in the preimplantation embryo: Na/K-ATPase and the origin of nascent blastocoel fluid. Dev. Biol., 105:330-342. 504. Searle, R. F., Sellens, M. H., Elson, J., Jenkinson, E. J., and Billington, W. D. (1976): Detection of alloantigen during preim¬ plantation development and early trophoblast differentiation in the mouse by immunoperoxidase labeling. J. Exp. Med., 14:348-359. 505. Warner, C. M., and Spannans, D. J. (1984): Demonstration of H-2 antigens on preimplantation mouse embryos using conventional anti¬ sera and monoclonal antibody. J. Exp. Zool., 230:37-52. 506. Kirkwood, K. J., and Billington, W. D. (1981): Expression of ser¬ ologically detectable H-2 antigens on mid-gestation mouse embryonic tissues. J. Embryol. Exp. Morphol., 61:207-219. 507. Sawicki, J. A., Magnuson, T., and Epstein, C. J. (1982): Evidence for expression of the paternal genome in the two-cell mouse embryo. Nature, 294:450^451. 508. Hakansson, S., and Peterson, P. A. (1976): Presence of beta-2„ microglobulin on the implanting mouse blastocyst. Transplantation, 21:358-360. 509. Heyner, S. (1983): Alloantigen expression on mouse oocytes and early embryos. In: Immunology of Reproduction, edited by T. G. Wegmann and J. J. Gill, pp. 79-99. Oxford University Press, Lon¬ don. 510. Pratt, H. P. M., Ziomek, C. A., Reeve, W. J. D., and Johnson, M. H. (1982): Compaction of the mouse embryo: an analysis of its components. J. Embryol. Exp. Morphol., 70:113-132. 511. Ducibella, T. (1980): Divalent antibodies to mouse embryonal car¬ cinoma cells inhibit compaction in the mouse embryo. Dev. Biol, 79:356-366. 512. Wiley, L. M., and Eglitis, M. A. (1980): Effect of colcemid on cavitation during mouse blastocoele formation. Exp. Cell Res., 127:89-

101. 513. Surani, M. A. H., Barton, S. C., and Burling, A. (1980): Differ¬ entiation of 2-cell and 8-cell mouse embryos arrested by cytoskeletal inhibitors. Exp. Cell Res., 125:275-286. 514. Maro, B., and Pickering, S. J. (1984): Microtubules influence com¬ paction in preimplantation mouse embryos. J. Embryol. Exp Mor¬ phol., 84:217-232. 515. Pratt, H. P. M., Chakraborty, J., and Surani, M. A. H. (1981): Molecular and morphological differentiation of the mouse blastocyst after manipulations of compaction with cytochalasin D. Cell, 26 279292. 516. Petzoldt, U., Burki, K., Illmensee, G. R., and Illmensee, K. (1983): Protein synthesis in mouse embryos with experimentally produced asynchrony between chromosome replication and cell division. Wil¬ helm Roux’s Arch. Dev. Biol., 192:138-144. 517. Petzoldt, U. (1986): Expression of two surface antigens and paternal glucose-phosphate isomerase in polyploid one-cell mouse eggs Dev Biol., 113:512-516. 518. Fleming, T. P., Cannon, P. M., and Pickering, S. J. (1986): The cytoskeleton, endocytosis and cell polarity in the mouse preimplan¬ tation embryo. Dev. Biol., 113:406^419. 519. Johnson, M. H., Chisholm, J. C., Fleming, T. P., and Houliston, E. (1986): A role for cytoplasmic determinants in the development of the mouse early embryo? J. Embryol. Exp. Morphol. ISuppl I 97:97-121. ' 520. Ducibella, T., Ukena, T., Kamovsky, M., and Anderson, E. (1977): Changes in cell surface and cortical cytoplasmic organization during embryogenesis in the preimplantation mouse embryo J Cell Biol 74:153-167. 521. Opas, J., and Soltynska, M. S. (1978): Reorganization of the cortical layer during cytokinesis in mouse blastomeres. Exp. Cell Res 113 208-

211. 522. Lehtonen, E., and Badley, R. A. (1980): Localization of cytoskeletal proteins in preimplantation mouse embryos. J. Embryol Exp Mor¬ phol., 55:211-225. 523. Johnson, M. H., and Maro, B. (1984): The distribution of cyto¬ plasmic actin in mouse 8-cell blastomeres. J. Embryol Exp Mor¬ phol., 82:97-117.

Early Embryogenesis / 524. Sobel, J. S., and Alliegro, M. A. (1985): Changes in the distribution of a spectrin-like protein during develoment of the preimplantation mouse embryo. J. Cell Biol., 100:333-336. 525. Sobel, J. S. (1983): Cell-cell contact modulation of myosin organ¬ ization in the early mouse embryo. Dev. Biol., 100:207-213. 526. Sobel, J. S. (1983): Localization of myosin in the preimplantation mouse embryo. Dev. Biol., 95:227-231. 527. Sobel, J. S. (1984): Myosin rings and spreading in mouse blastomeres. J. Cell Biol., 99:1145-1150. 528. Steinert, P. M., Steven, A. C., and Roop, D. R. (1985): The mo¬ lecular biology of intermediate filaments. Cell, 42:411-419. 529. Brulet, P., Babinet, C., Kemler, R., and Jacob, F. (1980): Mono¬ clonal antibodies against trophectoderm-specific markers during mouse blastocyst formation. Proc. Natl. Acad. Sci. USA, 77:4113-4114. 530. Brulet, P., and Jacob, F. (1982): Molecular cloning of a cDNA sequence encoding a trophectoderm-specific marker during mouse blastocyst formation. Proc. Natl. Acad. Sci. USA, 79:2328-2332. 531. Oshima, R. G. (1981): Identification and immunoprecipitation of cytoskeletal proteins from murine extra-embryonic endodermal cells. J. Biol. Chem., 256:8124-8133. 532. Oshima, R. G. (1982): Developmental expression of murine extraembryonic endodermal cytoskeletal proteins. J. Biol. Chem., 257:34143421. 533. Oshima, R. G., Howe, W. E., Klier, F. G., Adamson, E. D., and Shevinsky, L. H. (1983): Intermediate filament protein synthesis in preimplantation murine embryos. Dev. Biol., 99:447^-55. 534. Duprey, P., Morello, D., Vasseur, M., Babinet, C., Condamine, H. , Brulet, P., and Jacob, F. (1985): Expression of the cytokeratin endo A gene during early mouse embryogenesis. Proc. Natl. Acad. Sci. USA, 82:8535-8539. 535. Trevor, K., and Oshima, R. G. (1985): Preimplantation mouse em¬ bryos and liver express the same type I keratin gene product. J. Biol. Chem., 260:15885-15891. 536. Bissell, M., Hall, H. G., and Parry, G. (1982): How does the ex¬ tracellular matrix direct gene expression? J. Theor. Biol., 99:31-68. 537. Haeuptle, M.-T., Suard, Y. L. M., Bogenmann, E., Reggio, H., Racine, L., and Kraehenbuhl, J.-P. (1983): Effect of cell shape change on the function and differentiation of rabbit mammary cells in culture. J. Cell Biol., 96:1425-1434. 538. Suard, Y. M. L., Haeuptle, M.-T., Farinon, E., and Kraehenbuhl, J.-P. (1983): Cell proliferation and milk protein gene expression in rabbit mammary cell cultures. J. Cell Biol., 96:1435-1442. 539. Unemori, E. N., and Werb, Z. (1986): Reorganization of polymer¬ ized actin: a possible trigger for induction of procollagenase in fi¬ broblasts cultured in and on collagen gels. J. Cell Biol., 103:1021 — 1031. 540. Adamson, E. D. (1983): Growth factors in development. In: The Biological Basis of Reproductive and Developmental Medicine, ed¬ ited by J. B. Warshaw, pp. 307-336. Elsevier, Amsterdam. 541. Twardzik, D. R. (1985): Differential expression of transforming growth factor-a during prenatal development of the mouse. Cancer Res., 45:5413-5416. 542. Stromberg, K., Pigott, D. A., Ranchalis, J. E., and Twardzik, D. R. (1982): Human term placenta contains transforming growth factors. Biochem. Biophys. Res. Commun., 106:354—364. 543. Merimee, T. J., Grant, M., and Tyson, J. E. (1984): Insulin-like growth factors in amniotic fluid. J. Clin. Endocrinol. Metab., 59:752767. 544. Tabarelli, M., Kofler, R., and Wick, G. (1983): Placental hormones. I. Immunofluorescence studies of the localization of chorionic gonadotropin placental lactogen and prolactin in human and rat pla¬ centa and in the endometrium of pregnant rats. Placenta, 4:379388. 545. Soares, M. J., Julian, J. A., and Glasser, S. R. (1985): Trophoblast giant cell release of placental lactogens: temporal and regional char¬ acteristics. Dev. Biol., 107:520-526. 546. Boime, I., Boothby, M., Hoshina, M., Daniels-McQueen, S., and Darnell, R. (1982): Expression and structure of human placental hormone genes as a function of placental development. Biol. Reprod., 26:73-91. 547. Hoshina, M., Boothby, M., and Boime, I. (1982): Cytological localization of chorionic gonadotropin ot and placental lactogen mRNAs during development of the human placenta. J. Cell Biol., 93: 190-198.

229

548. Adamson, E. D., and Meek, J. (1984): The ontogeny of epidermal growth factor receptors during mouse development. Dev. Biol., 103:6271. 549. Hortsch, M., Schlessinger, J., Gootwine, E., and Webb, C. (1983): Appearance of functional EGF receptor kinase during rodent embry¬ ogenesis. EMBO J., 2:1937-1941. 550. Nexo, E., Hollenberg, M. D., Figueroa, A., and Pratt, R. M. (1980): Detection of epidermal growth factor-urogastrone and its receptors during mouse fetal development. Proc. Natl. Acad. Sci. USA, 77:27822785. 551. Muller, R., and Verma, I. M. (1984): Expression of cellular onco¬ genes. Curr. Top. Microbiol., 112:73—115. 552. Slamon, D. J., and Cline, M. J. (1984): Expression of cellular on¬ cogenes during embryonic and fetal development of the mouse. Proc. Natl. Acad. Sci. USA, 81:7141-7145. 553. Sherr, C. J., Rettenmier, C. W., Sacca, R., Roussel, M. F., Look, A. T., and Stanley, E. R. (1985): The c-fms proto-oncogene product is related to the receptor for the mononuclear phagocyte growth factor, CSF-1. Cell, 41:665-676. 554. Propst, F., and Vande Woude, G. F. (1985): Expression of c-mos proto-oncogene transcripts in mouse tissues. Nature, 315:516-518. 555. Muller, R., Slamon, D. J., Adamson, E. D., Tremblay, J. M., Muller, D., Cline, M. J., and Verma, I. M. (1983): Transcription of cellular oncogenes c-rasK' and c-fms during mouse development. Mol. Cell. Biol., 3:1062-1069. 556. Muller, R., Tremblay, J. M., Adamson, E. D., and Verma, I. M. (1983): Tissue and cell type-specific expression of two human c-one genes. Nature, 304:454-456. 557. Muller, R. (1983): Differential expression of cellular oncogenes dur¬ ing murine development and in teratocarcinoma cell lines. In: Teratocarcinoma Stem Cells, Vol. 10, edited by L. M. Silver, G. R. Martin, and S. Strickland, pp. 451—468. Cold Spring Harbor Lab¬ oratory, Cold Spring Harbor, N.Y. 558. Muller, R., Verma, I. M., and Adamson, E. D. (1983): Expression of c-one genes: c-fos transcripts accumulate to high levels during development of mouse placenta, yolk sac, and amnion. EMBO J., 2:679-684. 559. Pfeifer-Ohlsson, S., Goustin, A. S., Rydnert, J., Wahlstrom, T., Bjersing, L., Stehelin, D., and Ohlsson, R. (1984): Spatial and temporal pattern of cellular myc oncogene expression in developing human placenta: implications for embryonic cell proliferation. Cell, 38:585-596. 560. Pelham, H. R. B. (1986): Speculations on the functions of the major heat shock and glucose-regulated proteins. Cell, 46:959-961. 561. Howlett, S. K. (1986): A set of proteins showing cell-cycle dependent modification in the mouse early embryo. Cell, 45:387-396. 562. Wittig, S., Hensse, S., Keitel, C., Eisner, C., and Wittig, B. (1983): Heat shock gene expression is regulated during teratocarcinoma cell differentiation and early embryonic development. Dev. Biol., 96:507514. 563. Morange, M., Diu, A., Bensaude, O., and Babinet, C. (1984): Altered expression of heat shock proteins in embryonal carcinoma and mouse early embryonic cells. Mol. Cell. Biol., 4:730-735. 564. Heikkila, J. J., and Schultz, G. A. (1984): Different environmental stresses can activate the expression of a heat shock gene in rabbit blastocysts. Gamete Res., 10:45-56. 565. Ulberg, L. C., and Sheean, L. A. (1973): Early development of mammalian embryos in elevated temperatures. J. Reprod. Fertil. [Supplf, 19:155-161. 566. Bellve, A. R. (1976): Incorporation of [3H]uridine by mouse embryos with abnormalities induced by parental hyperthermia. Biol. Reprod., 15:632-646. 567. Kim, K., andMarkert, C. L. (1987): Are heat shock proteins involved in development? In: Developmental Toxicology: Mechanisms and Risk, edited by J. McLachlan, R. M. Pratt, and C. L. Markert. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. (in press). 568. Nusslein-Volhard, C., and Wieschaus, E. (1980): Mutations af¬ fecting segment number and polarity in Drosophila. Nature, 287:795801. 569. Garcia-Bellido, A. (1975): Genetic control of wing development in Drosophila. In: Cell Patterning, Ciba Foundation Symposium 29, pp. 161-178. Associated Scientific, Amsterdam. 570. Lewis, E. B. (1978): A gene complex controlling segmentation in Drosophila. Nature, 276:565-570.

230

/ Chapter 6

571. Gehring, W. J. (1975): The homeo box: a key to the understanding of development. Cell, 40:3-5. 572. McGinnis, W., Garber, R. L., Wirz, J., Kuroiwa, A., and Gehring, W. J. (1984): A homologous protein-coding sequence in Drosophila homeotic genes and its conservation in other metazoans. Cell, 37:403408. 573. Hart, C. P., Awgulewitsch, A., Fainsed, A., McGinnis, W., and Ruddle, F. H. (1985): Homeo box gene complex on mouse chro¬ mosome II: molecular cloning, expression in embryogenesis, and homology to a human homeo box locus. Cell, 43:9-18. 574. Jackson, I., Schofield, P., and Hogan, B. (1985): A mouse homeo box gene is expressed during embryogenesis and in the adult kidney. Nature, 317:745-748. 575. Joyner, A. L., Komberg, T., Coleman, K. G., Cox, D. R., and Martin, G. R. (1985): Expression during embryogenesis of a mouse gene with sequence homology to the Drosophila engrailed gene. Cell, 43:29-37. 576. Rabin, M., Hart, C. P., Ferguson-Smith, A., McGinnis, W., Levine, M., and Ruddle, F. H. (1985): Two homeo box loci mapped in evolutionarily related mouse and human chromosomes. Nature, 314:175-178. 577. McGinnis, W., Hart, C. P., Gehring, W. J., and Ruddle, F. H. (1984): Molecular cloning and chromosome mapping of a mouse DNA sequence homologous to homeotic genes of Drosophila. Cell, 38:675-680. 578. Wolgemuth, D. J., Engelmyer, E., Duggal, R. N., Gizang-Ginsberg, E., Mutter, G. L., Ponzetto, C., Niviano, C., and Zakeri, Z. F. (1986): Isolation of a mouse cDNA coding for a developmentally regulated, testis-specific transcript containing homeo box homology EMBO J., 5:1229-1235. 579. Rubin, M. R., Toth, L. E., Patel, M. D., D’Eustachio, P., and Nguyen-Huu, M. C. (1986): A mouse homeo box gene is expressed in spermatocytes and embryos. Science, 233:663-667. 580. Hauser, C. A., Joyner, A. L., Klein, R. D., Learned, T. K., Martin, G. R., and Tjian, R. (1985): Expression of homologous homeo-boxcontaining genes in differentiated human teratocarcinoma cells and mouse embryos. Cell, 43:19-28. 581. Joyner, A. L., Lebo, R. V., Kan, Y. W., Tjian, R., Cox, D. R., and Martin, G. R. (1985): Comparative chromosome mapping of a conserved homeo box region in mouse and human. Nature, 314:173175. 582. Colberg-Poly, A. M., Voss, S. D., Chowdhury, K., and Grass, P. (1985): Structural analysis of murine genes containing homeo box

sequences and their expression in embryonal carcinoma cells. Nature, 314:713-718. 583. Colberg-Poley, A. M., Voss, S. D., Chowdhury, K., Stewart, C. L., Wagner, E. F., and Grass, P. (1985): Clustered homeo boxes are differentially expressed during murine development. Cell, 43:39-45. 584. Carrasco, A. E., McGinnis, W., Gehring, W. J., and DeRobertis, E. M. (1984): A homologous protein-coding sequence in Drosophila homeotic genes and its conservation in other metazoans. Cell, 37:409414. 585. Muller, M. M., Carrasco, A. E., and DeRobertis, E. M. (1984): A homeo-box-containing gene expressed during oogenesis in Xenopus. Cell, 39:157-162. 586. White, R. A. H., and Wilcox, M. (1984): Protein products of the bithorax complex in Drosophila. Cell, 39:163-171. 587. Beddington, R. S. P. (1985): The development of 12th to 14th day foetuses following reimplantation of pre- and early-primitive-streakstage mouse embryos. J. Embryol. Exp. Morphol., 88:281-291. 588. New, D. A. T. (1978): Whole-embryo culture and the study of mammalian embryos during organogenesis. Biol. Rev., 53:81-122. 589. Wiley, L. M., and Pedersen, R. A. (1977): Morphology of mouse egg cylinder development in vitro: a light and electron microscopic study. J. Exp. Zool., 200:389-402. 590. Hsu, Y. C. (1978): In vitro development of whole mouse embryos beyond the implantation stage. In: Methods in Mammalian Repro¬ duction, edited by J. C. Daniel, Jr., pp. 229-245. Academic Press, New York. 591. Mintz, B. (1971): Allophenic mice of multi-embryo origin. In: Meth¬ ods in Mammalian Embryology, edited by J. C. Daniel, Jr., pp. 186— 214. W. H. Freeman, San Francisco. 592. Gardner, R. L. (1971): Manipulations on the blastocyst. In: Advances in the Biosciences, Vol. 6, edited by G. Raspe, pp. 279-296. Pergamon, Oxford. 593. Pierce, B. (1983): The cancer cell and its control by the embryo. Am. J. Pathol., 113:117-124. 594. Gardner, R. L. and Papaioannou, V. E. (1975): Differentiation in the trophectoderm and inner cell mass. In: The Early Development of Mammals, British Society for Developmental Biology, Symp. 2, edited by M. Balls and A. E. Wild, pp. 107-132. Cambridge Uni¬ versity Press, Cambridge.

The Physiology of Reproduction, edited by E. Knobil and J. Neill et al.. Raven Press, Ltd., New York © 1988.

CHAPTER

7

Biology of Implantation H. M. Weitlauf 242 • Embryonic Signals, 244 • Early Pregnancy Fac¬ tor, 249 • Influence of the Uterine Environment on Blas¬ tocyst Development, 250

Cellular Aspects of Implantation, 231 Attachment, 231 • Penetration of the Epithelium, 234 • Localized Changes in the Stroma, 235 • Decidualization, 235 • Preparation of the Endometrium, 236 • Transformation of the Endometrium, 240 • Initiation,

Summary, 251 References, 251

findings that are most consistent and that seem to reflect common elements in various species are emphasized in this chapter. Several extensive reviews dealing with specific as¬ pects of implantation should be consulted for further details regarding the controversial issues (1-11).

Implantation of embryos in the wall of the uterus is a basic feature of mammalian reproduction. It is the result of a complex series of interactive steps beginning with fixation of the blastocyst in the uterus and ending with formation of a definitive placenta. Details of the intervening steps vary in different species, but the fundamental elements of embryo attachment and penetration of the epithelium with invasion into the endometrium, as well as formation of decidual tissue by the uterus, are features common to many animals. Fur¬ thermore, in the species studied so far, it appears the em¬ bryos and the uterus must be “synchronized”; typically, this means that the embryos have reached the expanded blas¬ tocyst stage and that the endometrium has undergone certain hormone-dependent changes that cause it to become “re¬

CELLULAR ASPECTS OF IMPLANTATION Details of implantation vary in different species. How¬ ever, in all animals it ultimately involves a direct interaction of the trophoblast with the luminal epithelium of the uterus. This basic step has been described in detail from a morpho¬ logical perspective and provides a framework upon which physiological and biochemical observations can be organ¬

ceptive” to the embryo. Because of its complexity, the process of implantation is difficult to study in toto, and most research has been carried out on those component steps that can be dissected out for description or definition. This approach has been quite suc¬ cessful, and a vast literature has grown up over the last 50 years in which detailed accounts have been compiled de¬ scribing the processes of embryo attachment, the penetration of the epithelium, the endocrine basis for uterine receptivity, and the chemical and cytological changes associated with initiation of the decidual reaction and differentiation of de¬ cidual tissue. On the other hand, because they have focused on selected facets of the process, often in only one species, investigators have sometimes been tempted to look for and champion single mechanisms as the “cause of implantation. But because implantation is not an isolated event established at a moment in time, or even necessarily involving the same mechanisms in different species, there is no single cause or mechanism and the resulting controversies tend to dominate the literature. Because it is difficult to develop an overview of implantation from the perspective of the conflicts, those

ized.

Attachment For descriptive purposes it has been useful to consider that the attachment of embryos to the uterine epithelium occurs in two phases, apposition and adhesion (8); the pro¬ cesses involved are quite distinct. The term apposition de¬ notes the progressively increasing intimacy of contact be¬ tween the trophoblast and the uterine epithelium. In mice and rats the uterine lumen closes down around the embryos in the earliest phase of implantation and thus the uterus has the appearance of “clasping” the blastocysts (12). On the other hand, in animals such as the rabbit, the blastocyst enlarges to fill the uterine lumen and hence brings the trophoblast into apposition with the epithelium without gen¬ eral obliteration of the lumen (8,13). In either case, investment of embryos with maternal epithelium seems to provide the initial mechanism for their immobilization within the uterus. At this stage the blastocyst can be dislodged without damage

231

232

/

Chapter 7

by gentle perfusion of the uterus, and hence the earliest fixation of the embryo is functional rather than involving structural connections between the two individuals. During this phase there is a progressive interdigitation of microvilli and an increasingly intimate association between the mem¬ branes of the blastocyst and endometrium. The microvilli become shorter, more blunted, and irregular (14-19), and there is the appearance of large bulbous cytoplasmic pro¬ jections, particularly on the antimesometrial surface (2,20). The obliteration of the uterine lumen with progressively closer apposition of the apical ends of the endometrial ep¬ ithelial cells and the trophoblast has been referred to as the attachment reaction (15). The mechanisms responsible for the development of ap¬ position, clasping, and the attachment reaction have not been worked out in detail, although in mice, rats, hamsters, and guinea pigs the changes appear to be associated with a more generalized closure of the uterine lumen that is de¬ pendent on the endocrine status of the mother (13,16-18, 21-27). For instance, the uterine lumen remains relatively open in ovariectomized mice given no replacement hor¬ mones and becomes obliterated only after the animals are injected with progesterone (28,29). Following treatment with progesterone alone, the process of closure tends to be ar¬ rested at the stage characterized by simple interdigitation of microvilli and does not show the progressively closer ap¬ position characteristic of normal pregnancy. This “first stage” of closure can be maintained with progesterone for pro¬ longed periods in castrated animals and is induced to pro¬ gress to the second stage” only by adding estrogen to the regimen (30). As might be expected then, some degree of closure and clasping has been reported to occur in mice with lactational or experimentally induced delayed implantation (24,31). In those situations, the interaction between uterine epithelium and trophoblast is characterized by simple in¬ terdigitation of the microvilli as long as the animals are nursing or are maintained with progesterone. However, after the addition of estrogen the association between the embryo and the uterus becomes more intimate, with a reduction in the microvilli and progression to the typical attachment re¬ action (29). With the development of more intimate appo¬ sition in response to estrogen, there is apparently increased pressure on the blastocysts as their surfaces become marked and distorted by corresponding irregularities in the uterine epithelium (32). There is some degree of clasping of delayed-implanting rat embryos (23), but the preponderance of evidence indicates that, in that species also, estrogen must be added for a typical attachment reaction (33,34). It is now generally assumed that uterine closure and clasp¬ ing of embryos or other objects (35,36) involves both en¬ docrine-dependent resorption of fluid from the lumen, pre¬ sumably by way of the irregular cytoplasmic projections referred to as pinopods (37-39), and a mild generalized edema that occurs throughout the endometrium in response to estrogen (40,41). However, associated changes in the

apical membranes of the uterine epithelial cells (42) and the occurrence of increased membrane turnover (43) indicate that processes associated with uterine closure are not limited to the removal of luminal fluid. Furthermore, although clo¬ sure and clasping appear to be primarily hormone-dependent uterine functions, it has been reported that the presence of blastocysts (or oil droplets) in the lumen hastens the second stage of closure and the appearance of the attachment re¬ action (44), presumably because they provide some addi¬ tional stimulus to the epithelium. The significance of closure and the attachment reaction in mice and rats is presumably that a period of stability is provided during which adhesion between the embryo and the uterus can develop. However, in animals such as the rabbit, the mink, or the rhesus monkey, there does not appear to be a typical closure of the uterine lumen or clasping of the embryos (13,45), and it seems to be the expansion -of the blastocyst that is primarily responsible for establishing the initial apposition between trophoblast and epithelium. It is interesting in this regard that there is no evidence for pinopod-mediated uptake of fluid from the lumen of the rabbit uterus (46). In large domestic species also, with their strated the exsuperficial epitheliochorial-type implantation, the interdigitation of microvilli is particularly extensive and the embryos appear to be firmly fixed in the uterus even though a typical attachment reaction with progressive loss of microvilli is not observed (47-49). Adhesion of the trophoblast cells to the apical ends of the uterine epithelial cells develops as the apposition phase progresses. Cytologic evidence most often cited as dem¬ onstrating the development of adhesiveness is twofold. First, with tissue taken from progressively later stages in the early phase of implantation, there is an increase in the amount and frequency of distortion of one or both surfaces. This is thought to occur as a normal part of the attachment process and to be exaggerated as a differential shrinkage artifact when the tissues are fixed for microscopy. Second, not only are there extensive areas where closely apposed membranes of trophoblast and apical ends of the uterine cells are parallel and separated by less than 150 to 200 A, but primitive junctional complexes are established (8,9). Although some descriptions of these regions have indicated the existence of mature septate-type desmosomal junctions and areas of cytoplasmic confluence (18,23), the usual observation is of less specialized and more primitive types of junctional com¬ plexes and the absence of cytoplasmic confluence (8,9,12, 24,50). Once the epithelium is penetrated, however, typical mature junctional complexes are often observed to be shared between uterine epithelium and the invading trophoblast (50). The molecular basis for initial adhesion of trophoblast and uterine epithelium at implantation has not yet been determined. However, the expression of complementary surface glycoproteins and the elaboration of aggregation molecules and extracellular matrix material are known to

Biology of Implantation be involved in cell recognition and adhesion in some simple multicellular organisms, and these mechanisms are thought to be generally applicable to vertebrates as well (51). There¬ fore, most studies of the acquisition of adhesiveness at im¬ plantation have been concerned with documenting the changes in cell-surface molecules during the peri-implantation pe¬ riod. The several changes observed either at the time of normal implantation or after termination of the dormant phase associated with delayed implantation may be relevant to the development of adhesiveness. Thus, Enders and Schlafke (52), using a combination of colloidal thorium, ruthenium red, and concanavalin A-peroxidase, demon¬ strated the extensive distribution of negative charges and acidic glycoproteins on the surface of mouse uterus during the fourth and fifth days of normal pregnancy and on the seventh day of lactation-delayed implantation. Although those workers were the first to suggest that there is a reduction in the thickness of the glycocalyx at implantation, the meth¬ ods used were not considered to be quantitative. Others, however, have subsequently reported that there are both a generalized loss of anionic sites and a reduction in thickness of the uterine glycocalyx at implantation: rabbit (53), rat (54), and mouse (55). Interestingly, these changes are ap¬ parently not dependent on the presence of embryos, since they occur in pseudopregnant as well as pregnant animals. In the ferret, however, the uterine glycocalyx does appear to be reduced in thickness initially at the specific points of contact between the embryo and uterus, indicating that, in that species at least, the embryo plays a role in the process of modifying the uterine surface (56). A reduction in the thickness of the glycocalyx could be associated with the removal of masking residues and thus be responsible for opening a variety of specific binding sites. Indeed, Chavez and Anderson (55) have reported that binding sites for the Ricinus communis lectin (i.e., binds D-galactose) appear at both implantation and interimplantation areas in mouse uterus in the peri-implantation period. They suggest that lectin¬ like receptors for galactose, appearing on the embryos, could bind to those sites and act as part of the adhesion mechanism at implantation. Characteristics of surface molecules on the embryos have also been observed to change at the time of attachment. For instance, histocompatibility antigens (H-2) on dormant delayed-implanting mouse blastocysts decrease during reac¬ tivation (57,58), suggesting that these glycoprotein factors are either removed or masked; there is a reduction in binding of positively charged colloidal iron to the surface of these blastocysts as well, indicating a decrease in the density of negative charges (59—61). This histochemical finding was confirmed by the results of experiments utilizing free-zone electrophoresis (62,63). Such changes in the embryos are compatible with removal of terminal sialic acid residues or larger moieties of the oligosaccharide components of surface glycoproteins at the time of attachment and at the initiation of implantation.

/

233

In related experiments, lectins have been used to probe the glycocalyx of the embryo for changes in the “accessi¬ bility” of specific sugars at the time of implantation. Because it appears, in many species, that the initial attachment of embryos occurs on specific limited regions of their surfaces (52,56,64,65), it might be anticipated that those regions would acquire adhesiveness earlier than other areas on the embryos and, therefore, that changes in lectin binding which are relevant to implantation would have corresponding tem¬ poral and regional patterns in the peri-implantation period. Although binding of various lectins has been observed by several investigators, in most cases the appropriate temporal and regional changes have not been demonstrated. Thus, it was shown with the electron microscope that concanavalin A-peroxidase binds to the surface of both the embryos and the uterus of mice, but there was no evidence of a change with implantation or of regional differences in distribution on the blastocysts (52,66). Chavez and Enders (67), using ferritin-conjugated lectins on days 5 and 6 of normal preg¬ nancy, as well as during delayed implantation, found stagespecific changes in the binding of peanut agglutinin and the agglutinin from Ricinus communis to the surface of blas¬ tocysts as they prepared to implant (these lectins bind pref¬ erentially to A-acetylgalactosamine and D-galactose, re¬ spectively), but neither change could be implicated as a factor in the acquisition of adhesiveness because the binding pattern was not different in nonadhesive delayed-implanting embryos. On the other hand, those same workers (68) re¬ ported that the lectin from Dolichos biflorus (specific for Nacetylglucosamine) did bind to delayed-implanting embryos but did not bind to embryos that were reactivated and, al¬ though regional differences were not seen, they suggested that the loss of binding sites is associated with the acquisition of adhesiveness. With concanavalin A bound to red blood cells, Sobel and Nebel (69,70) observed increased agglu¬ tination over the abembryonic pole of mouse blastocysts and, similarly, concanavalin A attached to latex beads was shown to bind preferentially to the abembryonic end of delayed-implanting blastocysts (71). However, again, there was no change in this regional pattern of binding upon reactivation and subsequent implantation. From such ob¬ servations it cannot be determined whether changes in the glycocalyx which lead to changes in lectin binding are sim¬ ply not related to the acquisition of regional adhesiveness or whether the methods used are too insensitive to dem¬ onstrate subtle differences in binding. By contrast, it has been observed that binding of molecular [3H]concanavalin A to delayed-implanting embryos decreases in the abem¬ bryonic region during the process of reactivation and sug¬ gested that there is a regional change in availability of man¬ nose-like sugars, or membrane fluidity, on the embryo that is related to the development of adhesiveness (72). A similar temporal and regional change in the glycocalyx during reac¬ tivation of delayed-implanting mouse embryos was inferred from increased staining of the abembryonic pole with Alcian

234

/

Chapter 7

blue (71). Although the precise nature of the changes in the glycocalyx cannot be deduced from such information, the observation of such temporal and regional patterns is com¬ patible with the interpretation that they are related to the acquisition of adhesiveness at implantation. Pinkser and Mintz (73) are often credited with the first attempts to establish qualitative differences in composition of surface glycoproteins in preimplantation mouse embryos. Those workers labeled two- to four-cell embryos or morulaearly blastocysts differentially with [3H]- and [‘^glu¬ cosamine, combined them, and removed the glycocalyx with trypsin; the trypsinate was treated with pronase and the digest was subjected to gel filtration. They found more label and generally larger fragments in the material from blas¬ tocysts, but were unable to determine whether the change was due to synthesis of qualitatively new glycoproteins or due to differentially regulated expression of those present in the earlier stages. In either case, they did not examine blastocysts at the time they acquire adhesiveness; therefore, their results do not provide information strictly about changes associated with implantation. The reports that mouse blastocysts are capable of syn¬ thesizing a variety of glycoproteins that could be involved in changes in embryo adhesiveness (74), the suggestion that type-III collagen is present on the surface of late blastocysts and may be the material in the intercellular ridges (75), and the report that fibronectin is synthesized by the inner cell mass but not trophoblast of mouse blastocysts (76) have apparently not been followed up. If confirmed, the obser¬ vations that such extracellular factors are synthesized would have potentially important implications for the acquisition of adhesiveness at implantation (51). The fact that changes occur in both the thickness of the glycocalyx and the capacity of embryos and uterus to bind specific lectins demonstrates that there are alterations in surface glycoproteins in the peri-implantation period. Al¬ though it must be emphasized that the precise nature of the changes cannot be deduced from such information and that it cannot be assumed that the changes are responsible for the acquisition of adhesiveness, it seems probable that they are involved. The further possibility that aggregation factors and secreted extracellular matrix materials are involved in attachment of the embryos remains to be examined criti¬ cally.

Penetration of the Epithelium It is clear that, with the exception of those species having a superficial (i.e., epitheliochorial) type of placentation, all mammalian embryos penetrate the uterine epithelium and its associated basal lamina to establish a definitive vascular relationship with the mother. However, the process varies considerably from species to species in terms of both timing and precise cytological features, and the early literature was replete with what appeared to be conflicting observations.

Some conceptual order was brought to this complex problem when Schlafke and Enders (9) pointed out that there are really three general strategies for penetrating the uterine epithelium and that various animals have adopted one or another of these approaches. Many of the apparent incon¬ sistencies disappear when comparisons are confined to a single type of penetration. A brief description of the essential cytological details in each of the three categories is useful to demonstrate the various ways in which different species have solved the problem of breaching the epithelial barrier. Intrusive Penetration In species that have adopted the intrusive approach to penetration of the epithelium, the embryos are generally considered to be highly “invasive”; the ferret provides an excellent example (56,77). Attachment initially occurs at ^specialized regions (i.e., ectoplasmic pads) of developing syncytial trophoblast. Whereas the broader regions of syn¬ cytial plaque generally follow the contour of the apical end of the epithelial cells, the ectoplasmic pads tend to indent them and appear to provide the initial points of attachment. The first penetration of epithelium is seen as the projection of a thin fold of syncytial trophoblast between adjacent epithelial cells. Initially the processes are ectoplasmic, but as they enlarge and progress to the basal lamina, the cy¬ toplasm is found to contain the usual array of organelles. The trophoblastic membrane is observed to share both apical junctional complexes and punctate desmosomes with the lateral membrane of adjacent epithelial cells. It is not known how the original epithelial apical junctional complexes are breached, but the process typically occurs at many sites that are separated by only a few cells. Although the trophoblast eventually surrounds large numbers of epithelial cells, and there is evidence of cell death and phagocytosis, the over¬ whelming impression is of apparently undisturbed epithelial cells adjacent to the trophoblast. Indeed, it has been sug¬ gested that healthy epithelium is necessary for anchoring the trophoblast as it penetrates deeper into the endometrium (56). The trophoblastic processes pause at the basal lamina and are disposed along it for a brief time, then proceed to invade the stroma where they surround but do not penetrate the basal lamina of the capillaries. Other species having the intrusive type of penetration include the guinea pig (50,78,79) and the rhesus monkey (45). There are subtle but potentially important differences in detail in these animals. For ex¬ ample, the syncytial trophoblastic processes of both the monkey and the ferret typically pause at the epithelial basal lamina; in the monkey, they then go on to invade the basal lamina of endometrial blood vessels, something which is not seen in the ferret. The intrusion of the cellular trophoblast to form the iso¬ lated endometrial cups responsible for secretion of gonad¬ otropin in the horse provides an interesting variation in this form of epithelial penetration that appears to be unrelated to the process of implantation per se (80).

Biology of Implantation Displacement Penetration The rat and the mouse provide typical examples of dis¬ placement penetration (12,81). As the apposition phase pro¬ ceeds and the first signs of decidualization appear in the subjacent stroma, there is typically evidence of cell death in the epithelial layer and of detachment of cells either singly or in groups. With the light microscope, these cells occa¬ sionally appear as dark masses (W-bodies) between the em¬ bryo and the endometrium and were originally thought to reflect the passage of some material from the embryo to the uterus (82). However, it was subsequently shown with the electron microscope that these masses are really dead epi¬ thelial cells that are being phagocytized by the trophoblast (81). As the trophoblast comes into contact with the basal lamina, it pauses and sends out processes that undermine adjacent cells and thus extends the epithelial defect and increases the area available for contact with the embryo. The basal lamina is then breached, apparently not by the trophoblast but rather by ectoplasmic processes of under¬ lying decidual cells (83). The mechanism responsible for death and detachment of the epithelium in this type of implantation is not known. However, it does appear to be intrinsic to the uterus because it occurs in the “implantation chambers” associated with an oil-induced decidual reaction in pseudopregnant mice (8487) and is blocked by administration of actinomycin D (88). Considerable histochemical evidence has been presented which indicates that the activity of various lysosomal enzymes decreases in the epithelial cells adjacent to an implanting embryo but not in interimplantation areas or in pseudo¬ pregnant animals (89-94), and it has been argued on this basis that the cells are undergoing autolysis in response to an embryonic signal (95). When the sloughing of epithelium was prevented by actinomycin D, mouse blastocysts became attached to the epithelium but were unable to penetrate to the basal lamina or stroma and thus could not truly implant (88). Hence, mouse blastocysts seem to be only weakly invasive, and an intact uterine epithelium may be an effec¬ tive barrier to implantation. Therefore, autolytic destruction of the epithelium seems to be an important precondition for implantation in that species at least, and the finding that mouse blastocysts transferred to the uteri of cyclic females implant only if the epithelium is disrupted (96) supports this suggestion.

/ 235

of trophoblast extending to the basal lamina. The original nucleus is present for some time, and the lateral plasma membrane retains the original junctional complexes with its apparently normal neighboring cells. There may be more than one peg per trophoblastic knob, and after some delay the basal lamina is penetrated at the locations of these pegs and the trophoblastic processes proceed to penetrate the endometrial blood vessels. Subsequently, there is wide¬ spread formation of epithelial symplasma, and fusion occurs in areas between the trophoblastic knobs (99).

Localized Changes in the Stroma In rats and mice the endometrial stroma eventually under¬ goes dramatic cytological changes to form a decidua in response to the implanting embryo. This process is discussed in detail later, but it should be noted here that subtle changes occur locally in the endometrium as an early part of the decidual response. The most obvious of these are the in¬ creases in alkaline phosphatase activity (85) and vascular permeability (100-102), which occur in the stroma imme¬ diately adjacent to the embryos even before they have pen¬ etrated the epithelium. It is clear, even from these strictly morphological ac¬ counts of early pregnancy, that at least two different sorts of changes occur in the peri-implantation uterus. First, there are certain hormone-dependent changes in the endometrium that make subsequent steps possible. These “enabling” changes, such as closure of the lumen and the acquisition, by epithelium, of the potential for self-destruction, occur throughout the endometrium, apparently without regard to the presence of an embryo. By contrast, there are other changes in the endometrium that must be provoked by a stimulus, from either an embryo or a suitable experimental substitute. These “evoked” responses, such as autolysis of epithelial cells or decidual transformation of stromal cells, are typically localized to the endometrium adjacent to the embryo or the site of experimental stimulation. Although the actual roles of many of these changes have not been established with certainty, it is presumed that they are im¬ portant for the implantation process; a great deal of work has been directed at defining their molecular and cellular basis. The process of decidual transformation, and the as¬ sociated enabling changes leading to acquisition of sensi¬ tivity by the endometrium, may be the most thoroughly studied of all.

Fusion Penetration The rabbit provides an example of fusion penetration (50,64,97,98). First attachment occurs between syncytial knobs of the abembryonic trophoblast and individual epi¬ thelial cells. The apical membrane of the epithelial cell fuses with that of the trophoblastic knob, resulting in cytoplasmic confluency. As the epithelial cell is converted to syncytium, it becomes cytologically distinct from its neighbors and appears, using the light or electron microscope, as a “peg”

Decidualization Decidual transformation of the endometrium in response to an implanting embryo results in grossly observable in¬ creases in the size and weight of the uterus. This growth is due not only to proliferation and differentiation of the en¬ dometrial stromal cells, but also to swelling of the tissue caused by localized increases in vascular permeability and

236

/

Chapter 7

the development of tissue edema. Thus, following attach¬ ment the embryo typically becomes embedded in an en¬ larging mass of decidual tissue, each so-called “nidus” being separated from the others by intervening areas of nontransformed endometrium. Formation of the decidua is a con¬ spicuous part of the process of implantation, and although the function of this specialized tissue has not been deter¬ mined with certainty, it is generally believed to be a critical component of the mother’s response to the embryo. The discovery by Loeb (103-105) nearly 80 years ago, that a similar transformation of the endometrium could be induced by indifferent stimuli in the uteri of animals that were suitably prepared with ovarian steroid hormones, has led to the development of an experimental model for this maternal response to the implanting embryo. Several in¬ vestigators have pointed to subtle differences between the “deciduomata” of the experimental model and naturally oc¬ curring decidua, particularly with respect to the timing of development and minor morphological details (106-108). However, the basic processes responsible for decidual trans¬ formation in response to the embryo appear to be the same as those leading to formation of experimental deciduomata, and the model has come to be widely accepted (109). One of the most important observations made with the experimental model has been that the decidual response can be obtained only during a limited time in pregnancy or pseudopregnancy (110,111) and that this period varies with the nature of the experimental stimulus. Thus, a grossly traumatic stimulus such as crushing or cutting the uterus was found to be effective during a period of 3 or 4 days early in pseudopregnancy, whereas less traumatic stimuli such as intraperitoneal injection of pyrathiazine or the in¬ traluminal instillation of various chemical substances can elicit a response only during a period of a few hours (84,1 ti¬ ll 8). Furthermore, this period of maximum sensitivity to the so-called “nontraumatic stimuli” was found to corre¬ spond to the period of uterine receptivity for blastocysts as established by asynchronous embryo transfer experiments (119-126). With this information in hand, it then became possible to determine the endocrine basis for developing uterine sen¬ sitivity. In an extensive series of experiments with mice and rats, it was eventually shown that although progesterone alone would support the development of a deciduoma in response to traumatic stimuli, it is estrogen acting on the endometrium after preconditioning with progesterone for at least 2 days that is responsible for entraining the pattern of sensitivity and subsequent refractoriness characteristic of the peri-implantation period (1-3,109,118). As might be antic¬ ipated, this endocrine regimen is essentially the same as that necessary to obtain implantation of blastocysts in ovariectomized rats and mice (100,127,128). Once it was established that formation of this new “de¬ cidual organ” (129) was dependent on appropriate hormonal conditioning and the application of a suitable stimulus dur¬ ing a limited time, the emphasis of investigators shifted to defining those chemical and cytological changes within the

endometrium that are associated with establishing receptiv¬ ity and the differentiation of decidual tissue (1-7).

Preparation of the Endometrium Hormone-dependent changes in cell proliferation and dif¬ ferentiation occur in all compartments of the endometrium and, although it has not been possible to determine how the changes are related to the acquisition of receptivity or to the development of the decidual reaction, it is generally thought that they are essential.

Cell Proliferation Changes in the rates of proliferation of endometrial com¬ ponents have been examined in intact animals during the ^strous cycle and early pregnancy and pseudopregnancy: mice (130-132); rats (133-136); guinea pig (137-139); and hamster (140). Luminal epithelium. Mitotic activity in luminal epithe¬ lium of mice varies during the ovarian cycle, with a peak at about the time of ovulation and a smaller secondary peak 3 days later. The secondary peak falls on the third day of pregnancy if mating has occurred, and there is little or no mitotic activity in the luminal epithelium thereafter (130,131). A similar pattern appears in the rat, with the first peak at about the time of ovulation and a second one 2 days later on the second day of pregnancy or pseudopregnancy (133— 135). It has been reported similarly that in guinea pigs there is mitotic activity at about the time of ovulation and a sec¬ ondary peak 2 to 3 days later (137-139). With hamsters also, mitotic activity is found in the epithelium at the time of ovulation, with a marked increase 2 days later (140). Synthesis of DNA, as demonstrated by incorporation of [3H]thymidine and autoradiographic or scintillation counting techniques, follows the same pattern. Incorporation of [3H]thymidine increases in luminal epithelium of the mouse on the second day of pseudopregnancy and decreases there¬ after (131,141). In rats also, the epithelium is labeled with [3H]thymidine on the second and third days of pseudopreg¬ nancy (142). Glandular epithelium. Proliferation of endometrial glands tends to be low in mice at and before ovulation but shows a marked increase 3 days later, coincident with the second peak of epithelial mitosis (130,132). The pattern appears to be similar in the rat, with a peak on the second day of pregnancy (134,135,143); a corresponding incorporation of [3H]thymidine has been observed on the second day of preg¬ nancy (144). Mitotic activity in the glands of the guinea pig is of relatively greater extent than that in the mouse and the rat and appears somewhat later, with a peak on the fourth day of the estrous cycle (137-139). Stroma. A small amount of mitotic activity is observed in the stroma of the mouse just before ovulation and is minimal thereafter until a marked increase occurs on the fourth day of pseudopregnancy or pregnancy; the high levels

Biology of Implantation

of mitotic activity carry over into the fifth day, when im¬ plantation occurs (130,131). There appears to be some pref¬ erential distribution of the active cells near the luminal ep¬ ithelium, although mitosis occurs throughout the endometrium. A similar pattern is observed in the rat, with stromal mitosis appearing late on the third day and peaking on the fourth day, and the active cells appear to be con¬ centrated in the subepithelial antimesometrial stroma (134136). The pattern in the guinea pig seems to peak between the sixth and seventh days of the cycle (137-139). Incorporation of [3H]thymidine into DNA is observed in the stroma of mice and rats after epithelial mitosis has ceased. It is maximal on the fourth and fifth days of pseudopreg¬ nancy (131,141,142) in the subepithelial cells of the anti¬ mesometrial region (145) and appears to be a consequence of estrogen action after preparation by progesterone (146). The endocrine regulation of cell division in the various tissue compartments of the endometrium has been examined critically using ovariectomized animals given replacement therapy: mice (147-152); rats (136,146,153); guinea pig (137,138); and rabbit (154,155). The relationships between changes in the endocrine milieu and mitotic activity in the luminal epithelium, the glands, and the stroma have been most completely established for mouse endometrium. A description of these relationships in the mouse therefore seems useful as a basis for comparison. The observations on intact mice fit well with what has been discovered about endocrine regulation of these tissues in replacement exper¬ iments with ovariectomized animals; these observations also fit well with what is known about the changing levels of ovarian steroid hormones in the estrous cycle and early pregnancy (3,4). Luminal epithelium. Mitotic activity is nil in the luminal epithelium of ovariectomized mice given no hormone re¬ placement. However, following a single injection of estro¬ gen, there is an increase in mitotic activity; the response is biphasic with peaks at about 24 and 36 hr (147-150, 152,156-160). If estrogen is administered continuously, the effect on mitotic activity in epithelial cells is maintained for 2 days and then drops off (151). Progesterone administered by itself has no stimulatory effect on luminal epithelium but markedly reduces the effect of estrogen if the two hormones are administered simultaneously; typically, the first peak of mitotic activity is reduced and the second peak is blocked completely. The effect of progesterone can be obtained up to about 17 hr after estrogen (161,162). On the other hand, if progesterone is given for 3 days, a subsequent injection of estrogen has no effect on mitotic activity in the epithelium (163). The rat is apparently similar to the mouse in that estrogen stimulates mitotic activity in luminal epithelial cells within 24 hr; pretreatment with progesterone suppresses that effect (136,153). The rabbit seems to respond differently in that estrogen stimulates epithelium only mildly whereas pro¬ gesterone alone (in sufficiently high doses) is the more ef¬ fective stimulus; it has been reported, however, that priming with estrogen reduces the amount of progesterone needed for the mitogenic effect (154,155). In the ovariectomized

/

237

guinea pig, estrogen stimulates epithelial mitosis (137,138); pretreatment with progesterone prevents this effect (137). Glandular epithelium. A single injection of estrogen has no effect on mitosis in uterine glands of mice (150). How¬ ever, with continuous administration of estrogen a wave of mitosis is seen in the glands at 72 hr; this is followed by a second wave of mitotic activity 72 hr after terminating treat¬ ment (151). Progesterone given beforehand suppresses both waves of activity, whereas progesterone given with the es¬ trogen, or following estrogen treatment, blocks only the second wave. In the rat, estrogen causes an increase in mitosis in the glands within 24 hr and causes a further increase at 48 hr; again, pretreatment with progesterone suppresses that effect (136,153). In rabbits, there is reported to be some mitotic activity in the glands following treatment with estrogen alone, but it is much greater with progesterone (155). There is a question regarding control of mitosis in uterine glands of the guinea pig because some investigators have achieved proliferation with estrogen alone and suggest that the guinea pigs are like mice and rats (137); others have reported that mitotic activity is stimulated by progesterone after estrogen priming and suggest, therefore, that guinea pigs are similar to rabbits (138). Stroma. Neither estrogen nor progesterone alone appears to have an effect on mitotic activity in stroma in mice. However, progesterone is able to induce stromal mitosis if a priming dose of estrogen has been given previously (164). Furthermore, if progesterone is given for 3 days, the stroma undergoes marked hypertrophy in response to a subsequent injection of estrogen regardless of whether priming estrogen was given (150,163). Priming with estrogen appears to com¬ press the progesterone-induced changes into just 2 days (165). Interestingly, a second injection of estrogen given to progesterone-treated animals at 48 hr will cause a second increase in mitotic activity, but if it is given between 12 and 36 hr after the first injection, the endometrium is re¬ fractory and the estrogen is redirected to affect the epithe¬ lium (166). In the rat also, estrogen has no effect on stromal mitosis unless there has been pretreatment with progesterone (136,153). In addition, it appears the mitotic activity in the rat is concentrated in the antimesometrial stroma adjacent to the luminal epithelium (153). With the rabbit, estrogen alone causes a modest increase in stromal mitosis (155). Again, there appears to be a question in the guinea pig because some investigators find that progestrone alone stim¬ ulates little mitotic activity in the stroma but when estrogen is added it increases dramatically as it does in the mouse (137), whereas others report that progestrone after estrogen priming causes massive increases in stromal mitosis and suggest that the guinea pig is like the rabbit with regard to the regulation of proliferation in the endometrium (138). Associated with the changes in proliferation during the peri-implantation period there are marked changes in the levels of steroid receptors (167-171). However, it has been difficult to evaluate the significance of the changes with respect to development of endometrial receptivity and mi¬ totic activity, not only because of the heterogeneity within

238

/

Chapter 7

the receptor populations (172,173) but also because in many cases the measurements were made on whole uterus rather than in the various tissue compartments or cell populations of the endometrium. Because the myometrium accounts for as much as 90% of the mass of the uterus, measurements of the whole organ tend to reflect changes in that compart¬ ment rather than in the endometrium. The significance of this distinction was demonstrated by Glasser and Mc¬ Cormack (7), who observed that, as measured in whole uterus, both nuclear and cytosol estrogen receptor concen¬ tration decreases from the second or third day of pregnancy onward while in stroma the content actually increases. Mar¬ tel and Psychoyos (174) likewise found differential changes in cytosol estrogen receptor levels in the different uterine compartments in response to progesterone, with increases occurring in stroma after 48 hr accompanied by simulta¬ neous decreases in the epithelium. Although the concentra¬ tions of estrogen receptors therefore are correlated with the mitogenic effect of estrogen in the peri-implantation period, levels of receptor do not appear to be the limiting factor (174); also, the acquisition of sensitivity to estrogen after 48 hr of progesterone appears to involve other regulatory factors. In general, then, estrogen stimulates proliferation of both luminal and glandular epithelium, whereas progesterone in¬ hibits the mitogenic effect of estrogen on epithelium. Fur¬ thermore, estrogen given after several days of progesterone treatment causes the stroma, rather than epithelium, to pro¬ liferate. Pretreatment of the uterus with estrogen reduces the time necessary for this effect of progesterone to become manifest. The changes in mitotic activity are reflected, as would be expected, by parallel changes in the incorporation of [3H]thymidine into DNA. In light of these findings, the patterns of mitotic activity in cyclic and pregnant mice and rats have led to the hypothesis that it is the estrogen released at about the time of ovulation that initiates the first wave of epithelial proliferation, its withdrawal being responsible for a wave of activity in the glands on the third day of pregnancy. According to this hypothesis, progesterone from the corpus luteum then conditions the stroma, and the nidatory estrogen induces proliferation of the stroma on the fourth and fifth days in preparation for implantation. Thus, the hormone regimen necessary to establish a typical periimplantation pattern of mitotic activity in the endometrium of ovariectomized mice appears to be the same as that which is required to develop maximum uterine sensitivity to a nontraumatic intraluminal stimulus for decidualization (118). It is not known whether this pattern of proliferative activity enables the endometrium to respond to a decidual stimulus or merely accompanies the acquisition of sensitivity.

Cell Differentiation Morphological changes. Results of morphological ex¬ amination of the endometrium in cyclic and pregnant ani¬ mals, or castrated animals given various hormone replace¬

ments, demonstrate clearly again that changes occur in all compartments of the endometrium in response to ovarian steroid hormones. The cytological changes indicate that there are significant differences in overall synthetic activity in the stromal cells as the endometrium moves from presensitivity through the sensitive phase and becomes refractory. Luminal Epithelium; Uterine luminal epithelium be¬ comes atrophic after removal of the ovaries. The typical high columnar cells quickly change to a more cuboidal shape, with the nucleus located in a middle position. The cytoplasm becomes weakly basophilic, with meager amounts of en¬ doplasmic reticulum, and the Golgi complex is small. Nu¬ merous lipid droplets accumulate in the basal regions of the cells; and the apical microvilli become short, with little evidence of surface coat material. Following the injection of estrogen, the cells increase in height, the cytoplasm be¬ comes intensely basophilic, and there is an increase in the amount and prominence of the endoplasmic reticulum. The nuclei come to occupy a more basal position, the Golgi complex increases markedly and may occupy the lateral as well as the supranuclear area of the cells, and the apical microvilli increase in length. It is estimated that the apical surface area of the cells increases by as much as 50% and becomes covered with a “fuzz” (156-158,175). It is gen¬ erally presumed that this extracellular material is a typical glycocalyx composed of negatively charged acidic glyco¬ proteins (23,24,52,176). Other consistent findings follow¬ ing the administration of estrogen are the rapid dissipation of basal lipid droplets (177-181) and increases in several enzymatic activities, including alkaline phosphatase and nonspecific esterase (182-185). Progesterone, given by itself to ovariectomized animals, also leads to an increase in cell height. However, the cy¬ toplasm remains pale-staining and there is increased accu¬ mulation of lipid in the basal area of the cells (28). Although cathepsin D activity increases in the luminal epithelium upon stimulation with progesterone (186), there is no increase in alkaline phosphatase activity (184). However, probably the most dramatic cytologic change that takes place in epithe¬ lium treated for several days with progesterone is the de¬ velopment of extensive interdigitation of the apical micro¬ villi on the apposed luminal surfaces (28-30) and the formation of pinopods (20,65,187,188). It will be recalled that these features are characteristic of the first phase of uterine closure and are, therefore, associated with “presensitive” endo¬ metrium. The addition of estrogen in this situation leads to loss of the lipid droplets, increased prominence of the rough en¬ doplasmic reticulum, and a reduction in the number and size of the microvilli. Subsequently the apical membranes are more closely apposed and the surface becomes irregular, with attainment of the second stage of uterine closure char¬ acteristic of postsensitive or refractory endometrium (30). Although the transition from first to second stage of closure appears to be dependent on ovarian steroid hormones, the fact that it occurs more rapidly in the presence of an embryo

Biology of Implantation

or oil droplet in the lumen indicates that some additional factors are involved (44). The luminal epithelium of intact cyclic, or pregnant, animals shows corresponding changes in lipid and in the activity of several enzymes, presumably in response to changing levels of ovarian steroid hormones (186,189-193). Uterine Glands: There is relatively little information about endocrine-dependent cytodifferentiation of the glandular ep¬ ithelium in ovariectomized animals, and most attention has been directed at the hormonal basis for secretory activity. It is reported that continuous treatment of ovariectomized mice with progesterone (with or without estrogen priming) leads to increased glandular secretion after 8 days; simul¬ taneous injection of progesterone and estrogen leads to more massive secretory activity in just 5 days, whereas injection of estrogen after 3 days of progesterone leads to maximum secretion 48 hr later (194). Cells of the uterine glands in mice are typically more cuboidal in shape than luminal epithelium; they also have less lipid and appear to respond more slowly to ovarian steroid hormones (195). By the fourth day of pregnancy the gland cells are found to have large lucent apical vesicles that disappear on the fifth and sixth day, as the lumen becomes distended (196). With cytochemical methods it has been shown that this material contains carbohydrate and appears first in multivesicular bodies and on the concave, but not the convex, surface of the Golgi complex. The lumen is narrowed during the pro¬ gesterone domination characteristic of delayed implantation; the Golgi complex is typically located lateral to the nucleus, although there is abundant smooth and rough endoplasmic reticulum (197). The addition of estrogen after several days of progesterone domination leads to the second stage of uterine closure within 24 hr, and the glands start to become distended, although there are no significant ultrastructural changes in the cells. However, by 48 hr the glandular lumen becomes filled with carbohydrate-rich material, and elec¬ tron-dense granules are found at the apical border of the cells; the Golgi complex is located apical to the nucleus, and rough endoplasmic reticulum is present in increased amounts. That the glands are normally active on the sixth to seventh day of pregnancy or pseudopregnancy (i.e., 48 hr after nidatory estrogen is imposed on progesterone dom¬ ination) correlates well with the observed changes in en¬ docrine replacement studies. Stroma: The stromal cells also show hormone-dependent changes, particularly in the nucleoli (198). Tachi et al. (199) demonstrated, with the electron microscope, that following ovariectomy (a) the nucleoli in stromal cells are small and the fibrous component predominates, (b) there is a reduction in the amount of cytoplasm, and (c) the rough endoplasmic reticulum is less prominent. Following treatment with es¬ trogen alone there is only a slight increase in the granular component of the nucleoli and no significant change in cy¬ toplasmic features. By contrast, after treatment with pro¬ gesterone there is a marked enlargement of the nucleoli, with augmentation of the granular component. There are

/

239

also increases in the rough endoplasmic reticulum, which becomes distended, and the cytoplasm has many polyri¬ bosomes. If estrogen is added after progesterone pretreat¬ ment, all aspects of these changes are accentuated. Another prominent hormone-dependent feature in presensitive stroma is the development of a generalized edema that leads, in part, to the first phase of uterine closure (40,41,127,200). Metabolic changes: The results of extensive studies of uterine metabolism demonstrate that there are hormone-de¬ pendent changes at the molecular level which correlate with the changes in proliferation and cytodifferentiation observed in the peri-implantation period. The metabolism of uterine RNA in early pregnancy has been examined in some detail. Incorporation of [3H]uridine into RNA increases between the second and third day of pregnancy or pseudopregnancy in mice and rats (201-203). In rats the increase is biphasic, with a small peak on the third day and a more sustained increase on the fifth day, which then decreases unless decidualization occurs (204,205). This latter increase appears to be largely in the nuclear fraction, is thought to be associated with increased pro¬ cessing of ribosomal RNA (206,207), and is correlated tem¬ porally with the increased numbers of polyribosomes ob¬ served in stroma on the fifth day (136). Measurements of DNA template activity with bacterial RNA polymerase in vitro generally confirm the biphasic nature of the pattern of RNA synthesis, with the greatest activity in chromatin pre¬ pared from uteri on the third and fifth day (6,208). Fur¬ thermore, it has been inferred from DNA/RNA competitive hybridization studies that there are species of RNA synthe¬ sized on the fifth day of pregnancy that are not present on the second or the seventh day (208). Although increases in the synthesis of RNA were found in all compartments of the endometrium, the greatest increase was in the stroma (209). It is of particular interest that the activity was found to be greater in pregnant horns than in pseudopregnant horns, even at this time before attachment of the embryo. Ornithine decarboxylase activity increases within 4 hr of estrogen administration, and this change is prevented by cycloheximide (210). There is a biphasic peak in ornithine decarboxylase activity, with increases on the third and fifth day; this persists in implantation sites but not in interim¬ plantation areas (211,212). Synthesis of uterine proteins, as inferred from the incor¬ poration of labeled amino acids, also has a biphasic pattern in the peri-implantation period. Thus, incorporation of label increases on the third day of pregnancy, decreases on the fourth day, and increases again on the fifth (213,214). The increased rates of synthesis are maintained beyond the fifth day in implantation sites but not in interimplantation areas. With estrogen alone the increases are greatest in epithelium; with progesterone alone the increase is more dramatic in stroma (215). Along with the increase in overall protein synthesis, specific and apparently new proteins appear. These migrate in the post-transferrin region during electrophoresis on polyacrylamide gels (216,217). Interestingly, the pattern

240

/

Chapter 7

in pseudopregnant animals is different in that there is no increase in protein synthesis on the fifth day, again sug¬ gesting that the embryos provide some stimulus to the uterus even before attachment (214). Chemical analysis of endometrium for lipid throughout the peri-implantation period confirms the cytologic obser¬ vations. There are changes in both total lipid and the relative proportions of neutral triglycerides (218). The studies have been undertaken with both intact cyclic, or pregnant, ani¬ mals as well as with ovariectomized animals given replace¬ ment ovarian steroid hormone therapy. Although there are some species differences, it appears in general that estrogen reduces total lipid, primarily by influencing neutral triglyc¬ erides, whereas progesterone causes an increase; proges¬ terone is particularly effective in this regard when it has been preceded by estrogen (219 -223). Neutral triglycerides tend to accumulate in the endometrium, both epithelium and stroma, in the first 4 days of pregnancy, presumably because of the effect of luteal progesterone. Interestingly, lipids decrease in the implantation sites, but not interimplantation areas, on days 6 and 7 as decidualization proceeds (224). Although various investigators have suggested that neutral lipids provide an energy source for the developing embryo and that phospholipids may be important for synthesis of such components as cell membrane and prostaglandins dur¬ ing decidualization, it has not been possible to directly relate changes in lipids to regulation of proliferation or differen¬ tiation of the endometrium. Endocrine-dependent changes have also been observed in the intermediary metabolism of the endometrium during the estrous cycle and in the early progestational period in the rat. Attempts have been made to relate differing patterns of intrauterine oxygen tension, activity of glycolytic enzymes, uterine respiration, and glucose utilization to the shift in stromal cells away from mitotic activity and toward cytodifferentiation and the development of progestational sen¬ sitivity (5,189,225-233). Although it is clear that hormonedependent changes in cell proliferation, in cytodifferentiation, and in the various metabolic and synthetic patterns must ultimately be responsible for the development of uterine receptivity, none of the observations has provided a clear understanding of the process at the molecular level. However, an interesting suggestion has been made re¬ cently that may begin to explain the mechanisms of the process. Yochim and his colleagues have shown that the ovarian steroid hormones regulate availability of the pyri¬ dine nucleotide cofactor nicotinamide adenine dinucleotide phosphate and thus ultimately control pentose shunt activity in the endometrium (233-236). It is hypothesized by these workers that progestational sensitivity develops in the en¬ dometrium because increased levels of cofactor lead to in¬ creased pentose shunt activity and thus provide endometrial cells with increased capacity to recycle cofactors for re¬ ductive biosynthesis and to provide sugars for nucleic acid synthesis. This situation would facilitate cytodifferentiation, whereas the consequent reduced flux in the nicotinamide

adenine dinucleotide salvage path would limit production of ADP-ribose and thus may impair DNA synthesis and cell division (237-239). Transformation of the Endometrium Sensitization of the endometrium is essential for the sub¬ sequent decidual transformation in response to an appro¬ priate stimulus. Decidualization per se involves profound changes in both morphologic and biochemical characteris¬ tics of the endometrial stroma. Although details of timing and of the extent of endometrial involvement vary between species, the basic cytological changes are similar in different animals, and comprehensive descriptions of the process at the light microscopic level are to be found in several early papers (200,240-243). The precursors, from which decidual tissue forms, are Tibroblast-like cells in the stroma (142,145). In rats the process typically begins in compact areas of the subepithelial antimesometrial stroma (153), although this is apparently not critical because the initial changes occur throughout the antimesometrial region in the mouse (244) Among the ear¬ liest changes to be observed are the development of edema (26,101,123,124,245) and an increase in alkaline phospha¬ tase activity in the fibroblast-like precursor cells (85). Sub¬ sequently, these cells become slightly enlarged, the nuclei become rounded, and there is typically an increase in the prominence of the nucleoli. The cells tend to align them¬ selves into a continuous layer beneath the epithelium (12,18); and while mitotic activity in the central area declines and finally ceases, there is continued proliferation of cells in the areas adjacent to the developing nidus (130,136,246,247). As the process continues, the cells assume an epithelioid appearance (i.e., a characteristically rounded shape) and typically contain two or more polyploid nuclei (248). At the ultrastructural level, differentiation of decidual cells is characterized by the accumulation of glycogen, often in association with lipid droplets, and progessively increas¬ ing amounts of fibrillar material organized into parallel ar¬ rays and appearing as bundles in the cytoplasm. There is an increase in the number of polyribosomes, as well as in the amount and distention of rough endoplasmic reticulum; the nucleoli become enlarged, with an increase in the gran¬ ular component (12,26,108,249-254). It has recently been reported that the amount of fibronectin on the surface of these cells decreases as they undergo differentiation and that this change may be important in the development of their characteristic rounded shape (255). A consistent finding in the various electron microscopic studies of decidualization is that the cells are crowded closely together, with finger¬ like projections and numerous junctions of the adherens and gap types (12,249,256,257). The temporal aspect of the development of these junctions has been described in detail (250,258). Localized changes in vascular permeability is one of the earliest responses of the sensitive endometrium to any kind

Biology of Implantation

of deciduogenic stimulus and has most often been demon¬ strated by the extravascular accumulation of intravenously injected macromolecular dye (100,101). The development of fenestrations and gaps in the endothelium of endometrial blood vessels at implantation sites, but not between im¬ plantations, has been described and provides an explanation at the ultrastructural level for the so-called “Pontamine Blue reaction” in those areas (252,259). Although Lundkvist and Ljungkvist (250) argue that some cytological changes pre¬ cede the appearance of overt edema and the Pontamine Blue reaction, it has subsequently been shown, in the rat, that labeled albumin leaks from the vessels within 15 min of a decidualizing stimulus (260). Thus, the development of edema is still one of the earliest changes known to be associated with the endometrial response to a decidualizing stimulus. These changes can occur while blastocysts are still enclosed in the zona pellucida, and thus the signal does not require direct contact between trophoblast and uterine epithelium (102,261). Development of mature decidual tissue involves both pro¬ liferation and differentiation of the stromal cells. It is not surprising, therefore, that significant changes in both the total content and rates of synthesis of DNA, RNA, and protein have been observed. As indicated earlier, the syn¬ thesis of DNA can be demonstrated in endometrial stromal cells on the fourth and fifth days of pseudopregnancy or pregnancy by incorporation of [3H]thymidine (131,142,146). This increase in synthetic activity in the stroma follows the decline of proliferation in the epithelium. It occurs in re¬ sponse to the nidatory estrogen and accompanies the tran¬ sition through the receptive phase and into the refractory phase (141). The synthesis of DNA continues on the sixth day in areas undergoing decidualization, but not in those areas between implantations (144). However, it appears that once the decidual cells have become differentiated they no longer synthesize DNA; rather it is those cells peripheral to the forming nidus that continue to incorporate the labeled thymidine (145). It has been observed in mice that there are two populations of cells that begin DNA synthesis at about 11 to 15 hr after the decidualizing stimulus; one differen¬ tiates into mature decidual cells without dividing, the other goes on to divide before differentiating (262). The peak in DNA synthesis occurs about 30 hr after application of the decidual stimulus (263); chemical measurements of total DNA content revealed that changes are substantial, with increases of up to 70% per day in the second and third day after the experimentally applied stimulus (264). As might be expected, this increase in DNA content did not occur when decidualization was prevented by the antiestrogen MER25 (265). The decidual cells typically become binucleate and poly¬ ploid (246,247). Production of these nuclei, some contain¬ ing as much as 32n DNA in rats and 64n DNA in mice, involves endoreplication rather than fusion (248) and reaches a maximum at about 96 hr after application of the deci¬ dualizing stimulus (266). With rats it has been possible to

/

241

separate decidual cells on the basis of ploidy by means of differential sedimentation velocity on serum albumin gra¬ dients (267). It appears that those cells that are destined to develop the highest ploidy will synthesize DNA on the fourth day of pseudopregnancy, whereas those engaged in synthe¬ sis of DNA on the fifth day will typically develop lower ploidy (268,269). On the other hand, it appears that it is those cells that are synthesizing DNA early in the process of transformation which remain in the 2n to 4n population, whereas those synthesizing DNA later tend to end up in the 6n to 8n range. Thus, Moulton (268) finds evidence to support the concept that there are two populations of stromal cells in rats: one population that differentiates without di¬ viding, and a second population that divides before undergo¬ ing differentiation. It appears, then, that the rat is similar to the mouse (262) in this regard. The continued synthesis of DNA and RNA by decidual cells in ovariectomized an¬ imals is dependent on progesterone replacement, apparently more so in the cells at 4n to 8n DNA than in smaller cells. In contrast, synthesis of proteins is more progesterone-dependent in smaller cells than it is in the 4n to 8n population (268). The hormone-dependent nature of these various changes led some investigators to examine differences in steroid receptors in the endometrium. Martel and Psychoyos (270) report that the amount of estrogen receptor in implantation sites decreases relative to DNA and protein, and that there is little evidence of receptor in the nucleus. This seems to be compatible with the observation of decreased uptake of [3H]estradiol by implantation versus interimplantation sites (271,272). The conflicting report by Logeat et al. (169), that estrogen receptor increases markedly in implantation sites identified by Trypan Blue dye, appears to be based on the artifactual binding of steroids by the dye (270). Moulton and Koenig (273) have, however, reported that the number of estrogen receptors in cells with high ploidy increases relative to DNA, whereas McConnell et al. (274) report that progesterone receptors in the same cells decrease relative to DNA. These several observations demonstrate that a marked degree of heterogeneity exists in endometrial cells and de¬ veloping decidual tissue. Although it seems probable that these profound cellular differences are related to various aspects of the process of implantation and the specialized functions of decidual tissue, interpretation of the findings has been difficult and little progress has been made toward determining what role those differences actually play. Although synthesis of RNA occurs in stroma of pregnant and pseudopregnant mice and rats before the acquisition of sensitivity, further increases are observed with decidual transformation. Synthesis and accumulation of uterine RNA change dramatically upon formation of mature decidual tis¬ sue, with increases in content being observed as little as 5 hr after the systemic injection of pyrathiazine as the decid¬ ualizing stimulus (242). This early change in RNA was localized, by histochemical means, to the superficial cells of the antimesometrial stroma in rat (243). The increase is

242

/

Chapter 7

limited to decidualizing tissue and is not observed in those areas that fail to become transformed (203-205); it can be detected within 8 hr of the intraluminal application of the decidualizing stimulus (275). The overall increase in RNA content of decidual tissue has been estimated at 95% to 110% on the first day after stimulation (264,275). A cor¬ responding increase in uridine incorporation is observed and thought to reflect largely increased processing of ribosomal RNA (207,208); however, increases in activity of both RNA polymerase types I and III have been reported in the mouse (276), and template activity is significantly increased in decidualizing tissue on the sixth day of pregnancy, sug¬ gesting that there are substantial changes in the synthesis of all classes of RNA (6,209). Ornithine decarboxylase increases in a biphasic manner early in decidual transformation, presumably to support nu¬ cleic acid synthesis, and the increases are blocked by cycloheximide, actinomycin D (277), and indomethacin (278). Protein content in the rat deciduoma increases by as much as 70% per day in the second and third days after stimulation (264). In the mouse there is a biphasic increase in the protein content: a two- to threefold increase in the first 24 hr, and a secondary and more sustained increase of four- or fivefold in the third and fourth day (263). Rates of protein synthesis, as measured by incorporation of single radiolabeled amino acids, increase on the third day and fall on the fourth day. A larger and more sustained increase that is localized in implantation sites occurs on the fifth day (214,279). In¬ creases in protein synthesis are dependent on appropriate hormonal preparation of the endometrium as well as on the decidualizing stimulus and can be blocked with antiestro¬ gens (MER-25; 265), tamoxifen (215), and cycloheximide (279). These treatments also block the decidual transfor¬ mation. Progesterone appears to be necessary for continued high levels of protein synthesis by decidual tissue and may be acting primarily on the smaller stromal cells (i.e., those with 2-4n DNA; 268). Because decidual transformation is associated with significant increases in tissue mass, it is not surprising that there are marked increases in synthesis of DNA, RNA, and protein. However, although it has been essential to document the changes in macromolecular syn¬ thesis, the observations have not provided insights into either the nature of this unique process or the potential functional significance of the new tissue. As it became clear that increases in general protein syn¬ thesis accompanied the decidual transformation, efforts were made to determine whether decidual cell-specific proteins could be identified that might provide a clue to the function of this developing organ. Yoshinaga (280) prepared rabbit antiserum to crude extracts of rat deciduomata and found that it would prevent decidualization in both the rat and mouse (280,281); he suggested that some proteins in de¬ cidual tissue of these species have similar immunological characteristics. Similarly, Joshi et al. (282) reported the existence of a decidua-specific antigen in the baboon, and Sacco and Mintz (283) reported a uterine-specific antigen

on the fourth day of pregnancy in mice. Although none of these antigens has been characterized further, their existence demonstrates the potential for unique function of proteins in this tissue. Several investigators have demonstrated the existence of unique decidual proteins by resolving dual radiolabeled pro¬ teins on polyacrylamide gels (284-290). Again, however, none of these proteins has been described in sufficient detail to determine if they are the same between species. Denari et al. (286) observed a unique protein within 1 hr of the stimulus for decidualization in the rat. In terms of electro¬ phoretic mobility, this protein (protein A) was similar to estrogen-induced uterine protein (i.e., IP; 291-293). How¬ ever, it (i.e., protein A) was not increased in animals treated with estrogen alone and was not observed in nonstimulated uteri; it seems unlikely that this protein is the IP. Another protein with characteristics similar to those of IP was found 4o be synthesized maximally on the fourth and sixth days of pregnancy and depressed on the fifth day; the investigators suggested that it is associated with regulation of cell division (290). Although this protein was presumed to be IP because of its electrophoretic mobility, it was found to be greatly increased in deciduomata when there were no concomitant increases in estrogen; and because it was shown that IP is induced by estrogen in all cell layers (294), it seems unlikely that this protein is the IP. In addition, a pregnancy-asso¬ ciated protein was found in the post-transferrin region of the same gels. Its synthesis increased from the fourth through the sixth day of pregnancy and remained elevated in im¬ plantation sites but not in interimplantation areas (218,287,288). The significance of these various proteins is obscure. Two-dimensional polyacrylamide gel electrophoresis has recently been used because of the significant increase in resolution over conventional electrophoresis, and it has been possible to demonstrate in the rat that at least four new peptides appear with decidualization and that several others decrease (295). More recently, with hamster uterus, it has been shown that several decidua-specific nuclear and cy¬ toplasmic proteins are modulated both positively and neg¬ atively by progesterone (296), and more significantly, that decidual cells continue to produce these distinctive proteins in vitro (297). Because it should now be possible to purify these proteins, there is reason to expect that this approach will provide the breakthrough necessary to determine what the decidual cells are doing. Initiation That transformation of sensitized endometrium can be initiated by a variety of stimuli and will proceed in the absence of an embryo implies not only that elements nec¬ essary for decidualization are intrinsic to the uterus, but also that events entrained by the various stimuli converge at a common physiological point. It has been proposed that his¬ tamine and prostaglandins, released locally in the uterus in

Biology of Implantation

response to the various stimuli, provide the common locus and that these factors initiate the vascular and cellular changes of decidualization. Histamine The hypothesis that it is histamine released from uterine mast cells by the nidatory surge of estrogen that is respon¬ sible for initiating the decidual reaction was formulated over a period of several years by Shelesnyak and his colleagues, based on the following observations: (a) Histamine antag¬ onists instilled into the uterine lumen prevent the formation of deciduomata and reduce the number of implantations in rats (298,299); (b) histamine injected intraluminally, as well as histamine releasers administered systemically, induced decidual reactions in pseudopregnant rats (298-300); and (c) histamine content of rat uteri is reduced at the time of implantation (301), as well as in the uteri of ovariectomized rats following the injection of estrogen (302,303). Several investigators have disputed this hypothesis (109,304) and raise the following objections: (a) Intraluminal stimulators of the decidual response may be nonspecific (114,115,305); (b) systemic antihistamines are not particularly effective in blocking the decidual response (306-308); (c) in the hands of other investigators, instillation of histamine in the uterine lumen has not elicited greater responses than the vehicle alone, nor can a dose-response relationship be demonstrated (304,309,310); (d) depletion of histamine in mast cells with 48/80 does not prevent the decidual reaction (109); and (e) the decidual reaction normally occurs only in the vicinity of the embryo, or as discrete foci following administration of a systemic stimulus, and it would be expected that a generalized release of mast cell histamine in response to nidatory estrogen would result in a response throughout the uterus (109). Although many of the objections can be argued away, the failure of systemic antihistamines to block deciduali¬ zation was seen as damaging; thus the hypothesis was not universally accepted. The subsequent finding that there are two types of histamine receptors (Hi and H2), and that both may have to be blocked for a complete antihistamine effect, suggested that the early failures (i.e., typically with blockers of H2 receptors) did not constitute evidence against a role for histamine in implantation. Thus, interest was renewed when Brandon and Wallis (311) reported that implantation and decidualization were reduced in rats treated with a com¬ bination of blockers of Hi and H2 receptors (pyrilamine and burimamide, respectively). This finding seemed even more significant when coupled with the demonstration that rabbit blastocysts have H2 receptors whereas endometrium has the Hj type (312). However, more recently Brandon and Raval (313) were unable to block the attachment of embryos with another specific and more potent blocker of the H2 receptor (i.e., metiamide), and it has now been questioned whether the earlier effect with burimamide (311) was actually me¬ diated through an effect on H2 receptors (314).

/

243

Although the question of histamine receptors is still open, other evidence seems to support the hypothesis that hista¬ mine from mast cells is important in the uterine response. Thus, Ferrando and Nalbandov (315) depleted areas of the endometrium of mast cells by localized freezing and found that although this prevented implantation and decidualiza¬ tion, the effect could be overcome by instillation of hista¬ mine into the uterine lumen. Furthermore, Dey et al. (316) found that inhibition of histamine release from mast cells by means of intraluminal instillation of disodium cromoglycate prevented implantation and the decidual reaction in the rabbit and concluded that mast cell histamine plays a critical role in decidualization, and thus in implantation. From these observations, as well as those of Shelesnyak and his colleagues, it seems probable that histamine has something to do with decidual transformation and implan¬ tation. However, many of the objections raised by DeFeo (109) are still relevant, and a role for uterine histamine remains to be clearly defined.

Prostaglandins The proposition that uterine prostaglandins have an oblig¬ atory role in the development of endometrial vascular per¬ meability and subsequent decidual transformation is based on several lines of evidence (317,318). First, it has been shown that blocking the synthesis of prostaglandins with indomethacin during the first few days of pregnancy inhibits or delays implantation in mice (319-322), rats (323-325), hamsters (326), and rabbits (327). It was generally observed in those experiments that, following treatment with indo¬ methacin, the implantation sites appeared later and were smaller than in control animals and that embryonic devel¬ opment was retarded. Furthermore, the expected localized increases in vascular permeability associated with implan¬ tation are blocked or delayed by indomethacin in rats (324,328) and rabbits (329,330). In some cases, exogenous prosta¬ glandins partially overcame the effects of indomethacin (320,322,327). That this effect is on the uterus rather than on the embryos, or on the production of steroid hormones by the ovary, was shown by experiments with spayed an¬ imals given progesterone and estrogen replacement to achieve uteri with maximum sensitivity to decidualizing stimuli. Again, it was found that indomethacin blocked or greatly attenuated the artificially stimulated decidual response (331— 336). Second, the concentration of prostaglandins is observed to increase in decidualizing tissue in both normal pregnant animals and those with artificially stimulated deciduoma; again, this increase is blocked by indomethacin (324,328,334,337-341). Furthermore, exogenous prosta¬ glandins placed in the lumen of sensitized uteri are able to stimulate an increase in vascular permeability, even when endogenous prostaglandin synthesis is inhibited. There has been some disagreement about which of the prostaglandins

244

/

Chapter 7

is most effective. For example, Kennedy (328) found that prostaglandin E-2 (PGE-2) instilled into the lumen was ef¬ fective in increasing vascular permeability whereas pros¬ taglandin F-2a (PGF-2a) was not. On the other hand, con¬ stant infusion of PGF-2a was as effective as PGE-2 (336). Complete decidual transformation can be elicited by intra¬ luminal application of prostaglandins, with or without sup¬ pressing endogenous synthesis with indomethacin; PGF-2a instilled into the lumen is reported to be effective (342), and implants of PGE-2 and PGF-2a are both effective (343). More recently, it has been shown that there are specific receptors for PGE-2 in the stroma of rat endometrium (344) and that their concentration increases with progesterone, reaching a maximum on the fifth and sixth days of pseu¬ dopregnancy. There are no receptors for PGF-2a in the rat endometrium (345). It is suggested, therefore, that any ef¬ fect of PGF-2a on decidual tissues is a result of its con¬ version to PGE-2 or because PGF-2a cross-reacts with PGE2 receptors. This could provide an explanation for the effect of PGF-2a after constant infusion, when instillation as a bolus was less effective (328,336). However, the uterus is in a neutral state after several days of progesterone, and there was no change of PGE-2 receptor concentration upon addition of estrogen to develop the sensitive or receptive state (346). It has been reported that progesterone translo¬ cation to the nucleus is mediated by prostaglandins (347). Furthermore, production of prostaglandins increases in uter¬ ine tissue on the fifth day of pregnancy or pseudopregnancy (325). However, Kennedy (337) did not find such differ¬ ences in ovariectomized animals treated with ovarian steroid hormones, even though the expected increase in vascular permeability in response to intraluminal PGE-2 (or saline) was present. Thus, the acquisition of endometrial sensitivity does not appear to be directly correlated to the ability of the uterus to produce prostaglandins, and it appears that no simple relationship exists between the condition of endo¬ metrial receptivity and the level of prostaglandin production or receptors. Several observations have been made which suggest that changes in cyclic AMP mediate the decidualizing effects of prostaglandin at the cellular level. Thus, there is a rapid and dramatic increase in cyclic AMP following artificial stimulation of the decidual response (334,348-350), and this is inhibited by indomethacin (334,350). Furthermore, the instillation of cholera toxin causes increases in both vascular permeability and decidualization (349). Although the process of decidualization following cholera toxin may not be identical with that associated with PGE-2 (351), there are at least the typical changes in permeability and steroid receptors (352). On the other hand, intrauterine instillation of cyclic AMP or dibutyryl cyclic AMP does not induce decidualization (343,348,349,353), but will induce implan¬ tation if embryos are present (354,355). Thus, it is clear that prostaglandins have some obligatory role in implanta¬ tion, presumably involving the increase in vascular per¬ meability associated with decidualization, but the impor¬

tance of their role in the overall decidual transformation and the process of implantation remains unclear.

Embryonic Signals It has long been suspected that some form of embryonic signaling is necessary for the process of implantation and the “maternal recognition of pregnancy” (356,357). How¬ ever, the nature of the putative signals, and how they func¬ tion remains very controversial. There are several problems: First, the localized nature of the uterine response during the apposition phase of implantation implies that some type of embryonic signal acts at short range, whereas systemic changes associated with the maternal recognition of pregnancy, such as maintenance of the corpus luteum and modulation of the immune response, indicate that some are effective at longer -range. Therefore, it seems probable that in many cases there are more than one embryonic signal. Second, even when the purpose of embryonic signaling is the same in two spe¬ cies, the mode of action may be quite different. For ex¬ ample, in women and nonhuman primates it is the produc¬ tion of a chorionic gonadotropin that is responsible for “rescuing” the corpus luteum (358), and although there is controversy about whether that embryonic signal is actually synthesized by preimplantation blastocysts, it is clear that it operates as a luteotrophic factor (11). By contrast, in domestic animals such as the sheep, pig, and cow it is the production of an antiluteolysin by the embryo that is es¬ sential to neutralize the effect of uterine PGF-2a and thus prevent destruction of the corpus luteum. Many reports have appeared over the last 25 years that deal with the attempts to demonstrate that various potential signal substances are synthesized and released by preim¬ plantation embryos. At one time or another, carbon dioxide, steroids, histamine, prostaglandins, and proteins of embry¬ onic origin have all been proposed as signals. In addition to diffusible chemical factors, it has even been suggested that physical contact between the embryo and the endo¬ metrium provides a signaling mechanism. In reviewing the evidence for these various factors it should be kept in mind that it is unlikely that any one factor will be identified that could be considered as “the” signal for implantation in all animals; furthermore, in most cases there is no reason to suggest that any of the proposed factors are mutually ex¬ clusive.

Physical Stimuli Two observations have been used to support the hypoth¬ esis that it is physical contact between the embryo and the epithelium that is responsible for signaling at implantation. The first is the finding that embryo-sized beads of glass, paraffin, or agar produce decidual reactions in pseudopreg¬ nant rats (359,360). Although it is interesting, this hypoth¬ esis has not been supported by results of other experiments

Biology of Implantation

with various kinds of artificial or surrogate embryos. For example, unfertilized rat eggs, two-cell embryos, or mouse or sea-urchin eggs apparently had little or no ability to elicit the reaction in rats (360); McLaren (361) found that beads made of glass or an acrylic polymer did not produce deciduomas in pseudopregnant mice, and Blandau (359) was unable to obtain the reaction in pseudopregnant guinea pigs by using beads made of glass or paraffin. The second pro¬ posal is that because microvilli of the trophoblast interdigitate with those of the epithelium, pulsations of the blas¬ tocyst at this stage (362,363) might lead to distortion of the epithelium (317,364) and augment a physical signal. How¬ ever, the finding that localized edema will occur when em¬ bryos are still in the zona pellucida (102,262) makes the significance of this mechanism questionable. Although it may eventually be possible to demonstrate that such physical contact results in epithelial distortion and that it is important, the findings to date have not been convincing and none of the observations is clear-cut enough to assign a specific role to contact-mediated signals at implantation.

Carbon Dioxide The possibility that carbon dioxide (produced as a met¬ abolic by-product of developmentally active blastocysts) is important as a signal for implantation was originally pro¬ posed by Boving (365,366). It was hypothesized that carbon dioxide removed from the rabbit embryo as bicarbonate ion is converted to carbonic acid and an alkaline carbonate salt in the uterine epithelium, with the carbonic acid subse¬ quently being converted to carbon dioxide by carbonic anhydrase. It was envisioned that a resulting increase in pH could have local effects on the uterus. Hetherington (116,367) also suggested that embryonic carbon dioxide might be in¬ volved in eliciting the decidual reaction, since small bubbles of that gas, or air, were more effective in inducing a decidual response in pseudopregnant mice than was N2 or 02. Al¬ though there is no direct evidence to support this hypothesis, the observation that ethoxzolamide (an inhibitor of carbonic anhydrase) reduces the number of implantations in pregnant rabbits (366) is difficult to discount, and the question of a role for embryonic carbon dioxide as a unique embryonic signal at implantation remains unresolved.

Steroids The concept that the embryos synthesize steroid hormones which then play a role in implantation has evolved from the original observation by Huff and Eik-Ness (368) that 6-dayold rabbit blastocysts were not only capable of forming pregnenolone from [14C]acetate, but that they biotrans¬ formed” pregnenolone, 17ct-hydroxypregnenolone, proges¬ terone, and androstenedione to other phenolic compounds. The question of synthesis of estrogen by blastocysts and the putative involvement of such “embryonic estrogen in im¬

/

245

plantation (369) has been controversial. However, it should be noted that steroid metabolism by preimplantation em¬ bryonic tissue has been found in many of the animals that have been studied in detail (356,357,370,371). The obser¬ vations in various species are summarized as follows. Pig: It has been known for more than 10 years that 12to 14-day-old pig trophoblast is capable of synthesizing estrogens from labeled androstenedione, dehydroepiandrosterone, and testosterone and thus that the embryos have aromatase activity (370,372-374). The finding that pig blas¬ tocysts convert labeled progesterone and pregnenolone to estrogen, in the presence of a system for generating cofac¬ tors, demonstrates functional A5-3(3- and 17(3-hydroxysteroid dehydrogenase activities as well as those of the steroid C-17-20 lyase (375). Similar results have recently been re¬ ported by Fischer et al. (376), who demonstrated that es¬ trogen can be produced from labeled progesterone by pig embryos. Estradiol production was first observed at the large spherical blastocyst stage; estrone and estradiol were syn¬ thesized by tubular embryos, with amounts decreasing at the filimentous stage and increasing again between days 16 and 25. The enzymatic activity was demonstrated defini¬ tively in these studies by conversion of labeled substrates in vitro, as well as recovery and recrystallization (to constant specific activity) of the products. The results confirmed ear¬ lier histochemical findings on changing levels of A5-3(3- and 17P-hydroxysteroid dehydrogenase in the embryos between days 12 and 16 (377). Furthermore, the blastocysts have relatively high concentrations of estrogen and progesterone in utero, and the gradients between mother and embryo make it appear likely that the steroids are of embryonic rather than maternal origin (375,378). The maternal recognition of pregnancy in the pig occurs between day 10 and 12 (379) and involves an antiluteolytic effect. Bazer and Thatcher (380) have argued that estrogen of embryonic origin is involved as follows: (i) PGF-2a from the uterus is the luteolysin in the pig and is reduced in uteroovarian blood between days 12 and 20 of gestation (381,382), but its concentration in the uterine lumen is increased at that time (383); (ii) systemic estrogen duplicates this pattern of changes in PGF-2a (381,382); and (iii) it is estrogen from the embryo that is responsible for redirecting the secretion of endometrial prostaglandin from the bloodstream to the uterine lumen and thus spares the corpus luteum. Pig en¬ dometrium incubated with an embryo also has the capacity to convert progesterone to estrogen. There is no evidence for endometrial conversion of progesterone to estrogen by pseudopregnant animals and, therefore, the embryo must be responsible for altering the endometrial cells (376). A sec¬ ond possibility has been raised, namely, that estrogen from the embryo is sulfated in the endometrium (356,373,384) and, in the conjugated form, goes to the ovary, where it is luteotrophic (357). A third possibility has recently been raised with the report that pig blastocysts have the capacity for synthesis of catechol estrogens from estradiol (385). Because catechol estrogens have been implicated in regu-

246

/

Chapter 7

lation of prostaglandin synthesis, and because the transient increase in estrogen 2,4-dioxylase activity occurs at the time of maternal recognition of pregnancy, Mondschein et al. (385) suggested that estrogen synthesized by the embryo as a result of increased aromatase activity, between days 10 and 14 of pregnancy, is used in the formation of catechol estrogen, which acts as a signal in implantation. It is of interest in this regard that catechol estrogens have been reported to cause implantation in delayed-implanting mice (386) and to stimulate production of prostaglandins by preimplantation rabbit embryos and endometrium (387). The endometrium of pig also appears to be responsible for con¬ centrating steroids in the lumen that may act as substrates for embryos (388). Rabbit: The hypothesis that steroids of embryonic origin are important not only for development of the blastocyst but also locally to induce implantation (389,390) has been con¬ troversial. As applied to rabbits, this concept has been at¬ tacked on several grounds, including: lack of specificity of the histochemical assay (391); the supposition that high concentrations of steroids in rabbit blastocysts (392,393) are of maternal origin rather than from the embryo (394— 396); and the argument that the presence of enzymatic ca¬ pacity does not necessarily mean that it functions in vivo (397) . Nevertheless, definitive measurements were even¬ tually made of the conversion of dehydroepiandrosterone to androstenedione by 5-day-old rabbit embryos and of con¬ version of testosterone to estradiol by 7-day-old embryos (398) . Coupled with demonstrations of aromatase activity in cell-free lysates of embryos in the presence of an NADPHgenerating system (399) and in whole blastocysts in vitro (400), these observations appear to establish that the en¬ zymatic capacity for synthesis of estrogen exists in preim¬ plantation rabbit blastocysts. The question of a function for steroids associated with the embryos, whether of maternal or embryonic origin, is unresolved. The several observations that implicate estradiol as a factor in preimplantation development and implantation in the rabbit are: (i) incubation of blastocysts with the anti¬ estrogen CI-628 reduces their ability to implant when trans¬ ferred to pseudopregnant recipients, and the effect is re¬ versible (401); (ii) instillation of CI-628 into the uterine lumen reduces the number of implantations (402) and pre¬ vents the increases in acid phosphatase that are expected in luminal epithelium adjacent to the embryos (403); and (iii) estradiol binds to a soluble cytosolic protein in rabbit blas¬ tocysts, and this binding is blocked by CI-628 (401). Al¬ though these findings implicate estradiol in development and implantation in the rabbit, they do not prove that it is of embryonic origin. Indeed, the finding that an inhibitor of aromatase (4-hydroxy-4-androstene-3,17-dione) reduces blastocyst production of estradiol from testosterone in vitro, but does not interfere with either embryo development or implantation (400), is difficult to reconcile with that prem¬ ise. The further finding that rabbit endometrium can syn¬ thesize labeled estrogen from [3H]progesterone and

[3H]androstenedione and that the presence of the embryo influences this metabolic activity complicates this problem further (404). Rat, Mouse, and Hamster: Enzymatic capacity for steroid metabolism in preimplantation embryos of small laboratory rodents has also been inferred from the histochemical dem¬ onstration of A5-3f3- and 17(3-hydroxysteroid dehydrogenase activities in rats (405-408) and hamsters (409). Dickmann and his colleagues have published an extensive series of articles in which it has been proposed that estrogen of em¬ bryonic origin is important for the development of preim¬ plantation stage rat, mouse, and hamster embryos as well as for the initiation of implantation (390,410). The hypoth¬ esis is supported largely by the histochemical evidence for changes in enzyme activity at the morula and blastocyst stages and the observation that CI-628 blocks embryo de¬ velopment at the morula stage (411) and interferes with “implantation (91,412). Although this concept has now gen¬ erally been accepted with respect to embryos of pigs and rabbits, largely because it has been possible to demonstrate enzymatic conversion of precursors to estrogen with bio¬ chemical techniques in addition to the histochemistry, that has not been the case with the embryos of the smaller lab¬ oratory rodents. Indeed, attempts by several investigators to identify transformed products of pregnenolone, proges¬ terone, androstenedione, and dehydroepiandrosterone with preimplantation embryos of mouse and rat by radioimmu¬ noassay (413-415) or chromatographic methods have been unsuccessful (413-416). Although the various studies with CI-628 seem to point to estrogen of embryonic origin (91,403,411,412), a question has been raised as to whether the effects are due to nonspecific toxicity (411); these effects may be difficult to reconcile in light of the conflicting ob¬ servations that there is little estrogen receptor in the nucleus at implantation sites (270). Levels of enzymatic activity in hamsters do not change with development (417), and in¬ hibitors of aromatase (418) and steroidogenesis (419) do not block implantation, at least in hamsters. Other Species: Estrogen production by horse blastocysts in vitro has been reported (420), and in a comparative study with tissue from sheep, cows, roe deer, ferrets, cats, a plains viscacha, rabbits, and pigs, Gadsby et al. (374) reported observing significant aromatase activity and estrogen syn¬ thesis in pig trophoblast, whereas it was appreciably lower in all other species. In that study, labeled estrogens were recovered only from incubations of allantochorionic tissue of roe deer recovered shortly after implantation, as well as from pooled samples of tissue from early bovine embryos.

Histamine It has been reported that bits (421) and mice (422) synthesize histamine from decarboxylase). In rabbits

preimplantation embryos of rab¬ have the enzymatic capacity to histidine in vitro (i.e., histidine the activity peaks on the sixth

Biology of Implantation

day of pregnancy and intraluminal instillation of low doses of an inhibitor of histidine decarboxylase (a-methylhistidine dihydrochloride) on the fifth day of pregnancy delayed im¬ plantation and, at higher doses, interrupted implantation; simultaneous administration of histidine counteracted the inhibitor (423). Blastocyst formation in mice was also in¬ hibited with a-methylhistidine, and again this effect was overcome with histidine (422). These findings, along with the observation that histamine reduces the requirement for estrogen in inducing implantation in hypophysectomized progesterone-treated rats (424), prompted the suggestion that histamine synthesized by the embryo is important for development of the blastocyst and acts as a local signal to the endometrium at the time of implantation. Although the hypothesis that histamine produced by the embryo is in¬ volved in causing localized changes in the endometrium at implantation is appealing, it has not been substantiated and is difficult to reconcile with the report that histamine-re¬ leasing implants did not induce a significant decidual re¬ action in pseudopregnant rabbits (329). Furthermore, it is clear that localized decidual reactions will occur without an embryo being present.

Prostaglandins Several approaches have been taken in evaluating the ability of preimplantation embryos of different species to synthesize prostaglandins and in assessing their role in the process of implantation (425). Thus, it has been possible to demonstrate that some biological processes are suppressed in blastocysts in vitro by inhibitors of prostaglandin syn¬ thesis; it has been shown in vitro that the quantity of pros¬ taglandin within the embryo or released into the medium increases with time; and in some cases it has been possible to demonstrate the synthesis of labeled prostaglandins from radioactive arachidonic acid supplied either exogenously or from endogenous pools. The reported observations made with these various approaches can be summarized as fol¬ lows. Rabbit: The presence of prostaglandins in preimplanta¬ tion blastocysts was first reported by Dickmann and Spilman (426) . Prostaglandin of the F and E series was detected by radioimmunoassay in freshly recovered blastocysts on the sixth day of development. An increase in the content of prostaglandin F was also observed in rabbit blastocysts in¬ cubated in vitro for 24 hr (427), demonstrating that they do have the capacity for prostaglandin synthesis. Dey et al. (427) did not observe the release of prostaglandins into the medium during incubation of rabbit blastocysts. However, more recent studies have demonstrated both synthesis and release of prostaglandins E and F by 6- and 7-day-old rabbit embryos (428). Although it has not been possible to dem¬ onstrate the synthesis of labeled prostaglandins by rabbit blastocysts from exogenous arachidonic acid, it has been shown that when the endogenous phospholipid pools were

/

247

prelabeled in vitro with [3H]arachidonic acid and released by a calcium-specific ionophore, labeled prostaglandins were synthesized and released into the medium (425). From these observations it seems clear that rabbit blastocysts have the ability to synthesize and release prostaglandins to influence the endometrium locally. Furthermore, it appears that treat¬ ment with indomethacin early in pregnancy reduces the number of implantation sites in rabbits (327) and, therefore, that prostaglandins are involved in the process of implan¬ tation in this species. However, it remains unproven as to whether the local changes in endometrium (428) and the increase in concentration of prostaglandins at implantation sites (429) are caused by embryonic prostaglandins. Cow: Measureable amounts of immunoreactive prosta¬ glandins of E and F series were observed in cow blastocysts recovered on the thirteenth through the sixteenth day of development and incubated for up to 48 hr in vitro; the amounts increased in proportion to the ages of the embryos (430). Similarly, increasing amounts of radiolabeled pros¬ taglandins were recovered after incubation of 16- and 19day-old bovine embryos with radioactive precursors (431). Clearly, cow blastocysts have the capacity to synthesize prostaglandins, and again the suggestion that they (or other metabolites of arachidonic acid) might be important for em¬ bryonic development, act as local signals to the uterus, or be involved in maintenance of the corpus luteum (431) re¬ mains unproven. Rat and Mouse: It has not been possible to detect the synthesis of prostaglandins by preimplantation embryos of rat using radioimmunoassay methods, even after incubation of up to 150 embryos for 24 hr (318). Similarly, it has not been possible to demonstrate synthesis of labeled prosta¬ glandins by mouse blastocysts from either exogenous or endogenous [3H]arachidonic acid (425). However, in the case of the mouse at least, there is strong evidence that prostaglandins of embryonic origin are involved in expan¬ sion of the blastocyst, because several antagonists suppress the process of hatching in vitro (432-434). Although in¬ stillation of some of these prostaglandin antagonists into the uterine lumen also interfered with implantation, the degree of their effectiveness was not the same as that for suppres¬ sion of hatching (435), and it is not clear if they act at the level of the embryo or the endometrium. Because prosta¬ glandins of the E series are often involved in water transport across epithelia, Biggers and his colleagues have suggested that these prostaglandins are important for that function in the blastocyst as well and thus could be involved in im¬ plantation by virtue of maintaining the turgidity necessary for apposition and adhesion of the blastocyst and endo¬ metrium rather than as local signal factors (432,435). It must be recognized that these possibilities are not mutually exclusive. Sheep: Sheep blastocysts (at days 12 and 15 of devel¬ opment) were found to synthesize prostaglandins of the E and F series when incubated with labeled arachidonic acid (436), and the total amounts released into the medium in

248

/

Chapter 7

hr was 28 times the amount contained in the embryos at the time of recovery (437). Although the concentrations of prostaglandins E and F were shown to be high in both 14and 23-day-old blastocysts, and synthesis of prostaglandin F in the endometrium increased after day 14, indomethacin had no effect on implantation in this species (438). Taken together, these findings provide strong evidence for production of prostaglandins by preimplantation em¬ bryos of the rabbit, sheep, and cow. The evidence for the mouse is indirect and less compelling, and, as pointed out by Racowsky and Biggers (425), alternative methods will have to be used before the question can be settled with respect to the rat. When embryonic prostaglandins function as local signals to the uterus or are important to the embryo simply for maintaining normal cell function is unclear.

Proteins It has been known for almost 20 years that proteins of embryonic origin are important for the maternal recognition of pregnancy in sheep. Evidence is now beginning to ac¬ cumulate which supports the suggestion that such embryonic protein factors are involved in establishing pregnancy in a number of other species as well. Sheep: The concept that a protein signal of embryonic origin plays a role in maternal recognition of pregnancy in sheep dates back to the observations, by Moor and Rowson (439-442), that luteolysis is prevented if a conceptus is present between days 12 and 13, or if homogenates of 14to 15-day-old conceptuses are instilled into the uterine lu¬ men. The active principal in the homogenates was heat labile and presumed, therefore, to be a protein. In similar exper¬ iments, Martal et al. (443) confirmed this observation and demonstrated that the active material (which they called trophoblastin) from 14- to 16-day-old embryos was not only heat labile, but was ineffective after treatment with protease. That extracts from older embryos [i.e., 21 and 23 days old (443) or 25 days old (442)] were ineffective in prolonging the life of the corpus luteum led to the suggestion that the protein was synthesized by the embryos only during the period from day 13 to day 21. This protein factor is pre¬ sumably of trophoblast origin because transfer of tropho¬ blastic vesicles (from 11- to 13-day-old embryos) to the uteri of nonpregnant ewes (on day 12 of the cycle) prolonged luteal life (444). Although the signal has not been definitively identified, it has recently been shown in vitro that stage-specific pro¬ teins are synthesized and secreted by preimplantation sheep blastocysts. Thus, Godkin et al. (445) demonstrated with two-dimensional polyacrylamide gel electrophoresis that the major labeled product of 13-day-old sheep embryos incu¬ bated in vitro with [3H]leucine was a low-molecular-weight (17,000 dalton), acidic (PI 5.5) protein (initially designated protein X). Although several other proteins were synthe¬ sized and secreted by embryos between days 14 and 21,

protein X was predominant up to day 23, when it could no longer be detected. This protein is apparently synthesized in trophoblast, as shown by immunocytochemical methods (446), and has been redesignated “ovine trophoblast protein 1” (oTP-1). It binds to receptors in the endometrium with high affinity and apparently changes the pattern of protein synthesis in endometrium in vitro (446). This protein is the major translation product of trophoblastic mRNA, in a cellfree wheat-germ lysate system, and its production appears to peak in 13-day-old embryos (447). In addition, instillation of oTP-1 into the uterine lumen of cyclic ewes prolongs the life of the corpus luteum (448). However, this protein does not compete for luteinizing-hormone receptors on the corpus luteum nor does it stimulate progesterone synthesis (446), and thus it is apparently not the luteotrophic factor in con¬ ceptus homogenates reported by Godkin et al. (449) and Ellinwood et al. (450). For these reasons it has been sug¬ gested that oTP-1 is the embryonic protein factor involved in protecting against luteolysis (trophoblastin), presumably because its interaction with the uterine epithelium leads to altered release or metabolism of endometrial PGF-2a (447). In addition, Masters et al. (451) have reported that the major glucosamine-labeled product, purified by ion-exchange and gel-filtration chromatography of medium from 14- to 16day-old embryos, is a large glycoprotein (660,000 daltons), consisting of at least 50% carbohydrate (largely A-acetylglucosamine and galactose) and relatively resistant to pro¬ teolysis. A similar embryonic-secreted factor has been ob¬ served in the cow and pig, but no functional significance has been ascribed to this glycoprotein as yet. Cow: The cow is similar to the sheep and pig in that the presence of conceptus tissues in utero (prior to day 17) results in the maintenance of the corpus luteum (452-454). Furthermore, the infusion of homogenates of 17- and 18day-old embryos has been shown to delay luteal regression, although it is not known if the active principal is sensitive to heat or protease as it is in sheep (454). Preattachment bovine conceptuses have been shown to synthesize and se¬ crete a complex array of stage-specific proteins between days 16 and 24 of development (455). The individual pro¬ teins were separated by two-dimensional polyacrylamide gel electrophoresis and ion-exchange and gel-filtration chro¬ matography. The amount of radiolabel incorporated into secreted material increases from day 16 through day 22 and decreases by day 24. Several low-molecular-weight acidic proteins are secreted during this period which are similar to, but not identical with, those secreted by ovine tropho¬ blast. These factors are no longer evident by day 29 and thus are restricted to the period of maternal recognition of pregnancy in the cow. In addition, a large glycoprotein labeled with [3H]glucosamine was secreted by tissue from all stages including postimplantation (day 69) chorion. This may be the same factor isolated earlier by Masters et al. (451) using similar techniques. Pig: The time of maternal recognition of pregnancy in the pig is day 10 to day 12 and as with the sheep and the

Biology of Implantation / cow, a conceptus must be present in utero (prior to day 13) if the corpus luteum is to survive (379,456). It has been known for some time that the preattachment pig blastocyst is active in synthesizing and releasing proteins (457-459). Stage-specific proteins have been demonstrated in the pig in vitro by incubation of preattachment-stage embryos with radiolabeled precursors and by analysis of the conditioned medium using two-dimensional polyacrylamide gel electro¬ phoresis (460). The major labeled products between 10i and 12 days appear to be a group of low-molecular-weight acidic proteins similar to those reported for the sheep and the cow. However, between days 13 and 16 the major products are larger and more basic, and after day 18 the major secreted products are a group of serum proteins synthesized by the embryo rather than by the trophoblast. A large glycoprotein labeled with [3H]glucosamine and similar to that observed in sheep and cows was isolated with ion-exchange and gelfiltration chromatography from all stages (451). Mouse: The concept that proteins secreted by the preim¬ plantation mouse embryo might be involved in signaling at implantation dates back to the observation, by Fishel and Surani (461), that a labeled glycoprotein (approximately 87,000 daltons) could be recovered from the medium after incubating blastocysts with [3H]glucosamine. More re¬ cently, it has been demonstrated that a complex array of stage-specific proteins are synthesized and secreted when preimplantation mouse blastocysts are incubated with [35S]methionine (462). The proteins were isolated from con¬ ditioned medium and separated with two-dimensional poly¬ acrylamide gel electrophoresis. It was found that synthesis and secretion of labeled proteins increased between days 4 and 5 of pregnancy and, as with embryos of the sheep, pig, and cow, there were several low-molecular-weight acidic proteins released before implantation. Of special interest was the finding that some of the proteins secreted by the mouse embryos were decreased in amount as embryos en¬ tered the dormant phase associated with delayed implan¬ tation. Those proteins that decreased as embryos became dormant typically increased as the embryos were reacti¬ vated. Furthermore, the appearance of these secreted factors was correlated temporally with the appearance of the Pontamine Blue reaction in the uterus. Although these findings are highly suggestive of a signal role for secreted proteins in preimplantation mouse embryos, that function remains to be demonstrated.

Early Pregnancy Factor Several observations have been reported which indicate that other systemic signals of embryonic origin are involved in the maternal recognition of pregnancy. Of these, the socalled early pregnancy factor (EPF) has received the most attention. The existence of EPF was hypothesized after the observation that lymphocytes from pregnant mice had less activity in the rosette inhibition test with a standard anti¬

249

lymphocyte serum than did those from nonpregnant animals (463). Subsequently, it was found that the activity was a serum factor that enhanced the ability of rabbit anti-mouse serum to prevent rosette formation with normal red blood cells and spleen cells in the presence of complement (464,465). The amount of activity appeared to vary with the number of fetuses and to drop quickly once the embryos were re¬ moved (466). The factor was reported to suppress “adoptive transfer” of contact sensitivity to trinitrochlorobenzene, and it was suggested that it regulates cellular immunity in vivo (467). There are at least two types of EPF activity that appear at characteristic times in pregnancy (i.e., pre- and post¬ implantation). The early form is a large molecule [mice, 180,000 daltons by gel filtration (468); sheep, 250,000 dal¬ tons (469)], which can be separated into a nonactive protein fraction and an active factor (50,000 daltons) by ion-ex¬ change chromatography; recombining these fractions returns activity. It appears, then, that the early form of EPF is associated with a normal serum protein carrier. The active principal in the early form of EPF consists of two compo¬ nents, EPF-A and EPF-B. These components can be sep arated by differential precipitation with 40% NH4S04; nei¬ ther component alone has any effect, but activity is restored when they are recombined (469-471). In the mouse, EPFA is secreted by the oviduct in an inactive form and will not alter the rosette inhibition test until EPF-B is added; EPF-B is secreted by the ovary [in the presence of a pituitary factor later shown to be prolactin (472)] in response to a factor secreted by the fertilized or parthenogenically stim¬ ulated ovum (473-475). The early form of mouse EPF has been purified by immunoabsorption, electrofocusing, and gel filtration (476); the monomeric form has a molecular weight of 21,000 daltons and can be resolved into peptides of three sizes (i at 10,501 daltons is EPF-A; ii at 7,200 and iii at 3,400 daltons combine to form EPF-B). In mice, the late form of EPF appears to be produced by the embryo as the oviduct and ovary lose the capacity to synthesize com¬ ponents of the early form by about day 7 of pregnancy, but EPF activity can still be detected in the serum and urine in those animals as well as in animals ovariectomized on day 4 of pregnancy (468). Similar activity has been reported in sheep (470,475,477,478), humans (465,479-483), rats (484), pigs (485), and cows (475). It appears that EPF from mice, sheep, pigs, and humans have similar characteristics with respect to the effect on the rosette inhibition test and the appearance of different forms in each stage of pregnancy. Furthermore, there appears to be no species specificity; for example, human and pig ova produce a factor that will work in the mouse after intraperitoneal injection and extracts of fertilized mouse ova (but not unfertilized ova) elicit EPF activity when injected into the sheep oviduct (475). Whether EPF functions in regulation of the maternal im¬ mune system in pregnancy, or exists at all, has been ques¬ tioned by some investigators (486,487). On the other hand, because it (a) has been detected early in pregnancy in all species studied, (b) requires the presence of a viable embryo

250

/

Chapter 7

(or fetus), and (c) appears to last through at least the first half of pregnancy, some investigators have proposed that it be used as a diagnostic test for pregnancy (488-493).

Other Signals Several other putative signal-response mechanisms have been reported that may be related to early pregnancy factors. For example, two-cell hamster embryos are reported to re¬ lease an octapeptide that inhibits ovulation (494^496); fer¬ tilized horse ova are transported into the uterus while un¬ fertilized ova are retained in the oviduct (497,498); lactate levels in the mouse oviduct remain elevated longer in ani¬ mals with viable embryos than in pseudopregnant animals (499); and thrombocytopenia occurs in mice from the first day to the seventh day of pregnancy and does not occur in pseudopregnant mice (500). This latter response can be ob¬ served, however, within 3 hr of the transfer of fertilized embryos to pseudopregnant mice and thus seems to be ini¬ tiated by an embryonic signal. Although the importance of these various signal factors to the biology of implantation and the precise mechanism of their action in various animals remain obscure, the study of embryonic factors is of major importance and will be an important area for research in the next few years.

Influence of the Uterine Environment on Blastocyst Development In addition to observations which indicate that blastocysts can affect local and systemic changes in the mother during the peri-implantation period, it has become clear that de¬ velopment of the embryos is, in turn, influenced by the uterine environment. The uterine potential for regulating development of preimplantation embryos is dramatically il¬ lustrated by the phenomenon of delayed implantation. In that situation, development is arrested at the blastocyst stage for a period of several days, or even months, and resumes only in response to a change in the maternal endocrine status. During this phase of developmental quiescence the blastocysts typically have reduced levels of metabolic and synthetic activity, and cell division actually stops. In some species such a period of embryonic diapause occurs as an obligatory part of pregnancy; in others it may or may not occur depending on conditions in the maternal environment. In either case, following reactivation the “dormant” blas¬ tocysts resume development, and the subsequent implan¬ tation and fetal development are normal (501,502). It is generally accepted that the uterus is responsible for the embryonic quiescence associated with delayed implantation, because removing blastocysts to extrauterine sites either in vivo or in vitro leads to their metabolic reactivation (503). The presumed mechanism, as proposed 50 years ago by Brambell (504), is that the uterus regulates development in delayed implantation by either (a) restricting a critical “growth

factor” or (b) secreting an “inhibitory substance” into the lumen. Most studies directed at defining the mechanisms responsible for embryonic quiescence in delayed implan¬ tation have made use of the fact that the blastocysts become “reactivated” in vitro after being removed from the uterus and incubated for a few hours in various tissue culture me¬ dia. In this case, reactivation is characterized by (a) in¬ creases in metabolic activity and macromolecular synthesis, as occurs with reactivation of embryos in vivo (503,505518), and (b) the outgrowth of trophoblast cells, which has been likened to the initial changes associated with implan¬ tation in utero (519-524). The observations most often cited to support the concept that restriction of essential factors is a mechanism for de¬ layed implantation can be summarized as follows: Tropho¬ blast outgrowth does not occur in vitro in the absence of certain amino acids (521-524), serum factors (521,522), or glucose (523,525,526). It has been suggested from such observations that the uterus might impose developmental quiescence on the embryos by restricting one or more of these factors or even restricting concentrations of various ions (526-532) during delayed implantation. However, the level of amino acids in uterine fluid from delayed-implanting mice appears to be the same as that in normal animals (533), and although deletion of amino acids or serum from the culture medium prevents outgrowth, it apparently does not prevent metabolic activation (516). The suggestion that em¬ bryos do not develop beyond the blastocyst stage in vitro in the absence of glucose because they are energy deficient and thus that developmental arrest in vivo is due to the same cause (534) has not proven tenable. Dormant embryos ac¬ tually have a higher ATP/ADP ratio than reactivated em¬ bryos and, indeed, it appears that reduced utilization of glucose by dormant embryos (514,535) is due to allosteric inhibition of glycolysis because of the high energy state of the cells (512). Furthermore, it has been impossible to main¬ tain metabolic quiescence in vitro by restricting the con¬ centrations of various ions in the medium, and it now seems unlikely that this is a mechanism by which the uterus renders the embryos quiescent in vivo (513). Several investigators have reported the results of experiments in which embryos were incubated in uterine fluid and examined for changes in metabolic activity or shedding of the zona pellucida (507,508,517,518,536-538). Results indicate that there is a factor present in flushings of uteri from cyclic or pseu¬ dopregnant animals that reduces RNA synthesis by blas¬ tocysts in vitro. The factor is heat stable, dialyzable, and is neutralized in the uteri of pregnant, but not pseudopreg¬ nant, animals 6.5 hr after the injection of estrogen (538). The nature of this putative inhibitory factor has not been determined. It is not known if the blastotoxic factor reported by Psychoyos et al. (539) is the same one responsible for these changes, and although these results provide support for the interesting possibility that the uterus can inhibit em¬ bryonic growth, this proposal has not been proven conclu¬ sively and has not been universally accepted.

Biology of Implantation

SUMMARY It is obvious from the foregoing presentation that im¬ plantation of mammalian embryos is a complex phenome¬ non in which a variety of interactive processes occur be¬ tween the conceptus and the mother. It should also be clear that although various facets of the process have been de¬ scribed in much detail, relatively little is known at the mo¬ lecular or cellular level about the actual mechanisms re¬ sponsible for implantation in any one species, let alone in a comparative sense. However, in spite of our failure to understand totally this critical aspect of mammalian repro¬ duction, it is possible to develop a general overview of the process in hopes that it will allow the reader to focus on common and, thus, presumably important features. (a) The embryo and the uterus must be synchronized. From work in mice and rats, this seems to be related largely to the ability of the uterus to respond to an appropriate stimulus from a blastocyst, with changes in both the epi¬ thelium and stroma leading to attachment and the formation of a decidua. The period of uterine receptivity is limited, and the changes responsible for this condition are entrained by estrogen in progesterone-conditioned endometrium. The transition from nonsensitivity to receptivity and on to re¬ fractoriness is associated with many changes in the endo¬ metrium, including altered rates of synthesis of RNA and protein, cell proliferation, and various changes in cytologic characteristics. However, it is not clear, at the cellular or molecular level, what uterine receptivity is; at present, sen¬ sitivity is only an operational definition that describes an essential condition for implantation. (b) In response to a locally effective signal from the embryo, the sensitized endometrial stroma undergoes the process of decidual transformation. This typically involves cellular proliferation and differentiation, including the de¬ velopment of localized increases in vascular permeability, polyploid nuclei, dramatic cytological changes, and syn¬ thesis of unique species of RNA and protein. The so-called “decidua” that are formed typically provide a solid mass of cells into which the conceptus is embedded. The process responsible for formation of decidual tissue appears to re¬ quire an intact epithelium to conduct the embryonic signal to the stroma and may utilize histamine and/or the local release of prostaglandin E-2 to initiate the reaction. Al¬ though formation of the decidua is a conspicuous part of the process of implantation in many species, and it seems to represent the development of an entirely new organ at implantation, its actual function remains unknown. (c) The trophoblast of the embryo and the luminal epi¬ thelium of the uterus become adherent, with or without subsequent penetration of the endometrium. Adhesion of embryos to the uterine epithelium presumably involves changes in the glycoprotein molecules on one or both sur¬ faces. Changes in lectin binding may reflect the expression of complementary surface glycoproteins and, thus, might be related to the acquisition of adhesiveness. However, it

/

251

has not yet been shown that these changes are causally related to attachment and implantation. (d) In response to a variety of signal factors from preim¬ plantation embryos, there are local and systemic changes in the mother that can be considered to constitute the “maternal recognition of pregnancy.” It is generally assumed that im¬ plantation and subsequent pregnancy will not be successful unless such recognition takes place. REFERENCES 1. Psychoyos, A. (1973): Hormonal control of ovoimplantation. In: Vitamins and Hormones: Advances in Research and Applications, Vol. 31, edited by R. S. Harris, P. L. Munson, E. Diczfalusy, and J. Grover, pp. 201-256. Academic Press, New York. 2. Psychoyos, A. (1973): Endocrine control of egg implantation. In: Handbook of Physiology, Section 7: Endocrinology, Vol. II, Part 2, edited by R. O. Greep and E. B. Astwood, pp. 187-215. American Physiological Society, Washington, D.C. 3. Finn, C. A. (1977): The implantation reaction. In: Biology of the Uterus, edited by R. Wynn, pp. 245-308. Plenum Press, New York. 4. O’Grady, J. E., and Bell, S. C. (1977): The role of the endometrium in blastocyst implantation. In: Development in Mammals, Vol. 1, edited by M. H. Johnson, pp. 165-243. North-Holland, New York. 5. Yochim, J. M. (1975): Development of the progestational uterus: Metabolic aspects. Biol. Reprod., 12:106-133. 6. Glasser, S. R., and Clark, J. H. (1975): A determinant role for progesterone in the development of uterine sensitivity to decidualization and ovo-implantation. In: The Developmental Biology of Re¬ production, edited by S. L. Markert and J. Papaconstantinou, pp. 311-345. Academic Press, New York. 7. Glasser, S. R., and McCormack, S. A. (1981): Cellular and molec¬ ular aspects of decidualization and implantation. In: Proteins and Steroids in Early Pregnancy, edited by H. M. Beier and P. Karlson, pp. 245-310. Springer-Verlag, New York. 8. Enders, A. C. (1972): Mechanisms of implantation of the blastocyst. In: Biology of Reproduction: Basic and Clinical Studies, edited by J. T. Velardo and B. A. Kasprow, pp. 313-333. Symposium on Reproductive Biology, Sponsored by Third Pan American Congress of Anatomy. 9. Schlafke, S., and Enders, A. C. (1975): Cellular basis of interaction between trophoblast and uterus at implantation. Biol. Reprod., 12:41— 65. 10. Wimsatt, W. A. (1975): Some comparative aspects of implantation. Biol. Reprod., 12:1-40. 11. Heap, R. B., Flint, A. P. F., and Gadsby, J. E. (1979): Role of embryonic signals in the establishment of pregnancy. Br. Med. Bull., 35:129-135. 12. Enders, A. C., and Schlafke, S. (1967): A morphological analysis of the early implantation stages in the rat. Am. J. Anat., 120:185-226. 13. Hedlund, K., Nilsson, O., Reinius, S., and Aman, G. (1972): At¬ tachment reaction of the uterine luminal epithelium at implantation: Light and electron microscopy of the hamster, guinea-pig, rabbit and mink. J. Reprod. Fertil., 29:131-132. 14. Nilsson, B. O. (1966): Structural differentiation of luminal membrane in rat uterus during normal and experimental implantations. Z. Anat., 125:152-159. 15. Nilsson, B. O. (1970): Some ultrastructural aspects of ovo-implan¬ tation. In: Ovo-implantation. Human Gonadotropins and Prolactin, edited by P. O. Hubinens, F. Lercy, C. Robyn, and P. Leleux, pp. 52-72. S. Karger, New York. 16. Young, M. P., Whicher, J. T., and Potts, D. M. (1968); The ultra¬ structure of implantation in the golden hamster (cricetus auratus). J. Embryol. Exp. Morphol., 19:341-345. 17. Potts, D. M. (1966): The attachment phase of ovoimplantation. Am. J. Obstet. Gynecol., 96:1122-1128. 18. Potts, D. M. (1968): The ultrastructure of implantation in the mouse. J. Anat., 103:77-90. 19. Potts, M. (1969): The ultrastructure of egg-implantation. In: Ad¬ vances in Reproductive Physiology, edited by A. McLaren, pp. 241— 267. Logos, London.

252

/

Chapter 7

20. Psychoyos, A., and Mandon, P. (1971): Scanning electron micros¬ copy of the surface of the rat uterine epithelium during delayed implantation. J. Reprod. Fertil., 26:137—138. 21. Nilsson, O. (1967): Attachment of rat and mouse blastocysts onto uterine epithelium. Int. J. Fertil., 12:5-13. 22. Reinius, S. (1967): Ultrastructure of blastocyst attachment in mouse. Z. Zellforsch. Mikrosk. Anat., 77:257-266. 23. Potts, D. M., and Psychoyos, A. (1967): Evolution de 1’ultrastructure des relations ovoendometriales sous 1’influence de l’oestrogene, chez la ratte en retard experimental de nidation. C. R. Acad. Sci. Paris Ser. D, 264:370-373. 24. Potts, D. M., and Psychoyos, A. (1967): L’ultrastructure des rela¬ tions ovoendometriales au cours du retard experimental de nidation chez la souris. C. R. Acad. Sci. Paris Ser. D, 264:956-958. 25. Mayer, G., Nilsson, O., and Reinius, S. (1967): Cell membrane changes of uterine epithelium and trophoblast during blastocyst at¬ tachment in rat. Z. Anat. Entwick., 126:43-48. 26. Tachi, S., Tachi, C., and Lindner, H. R. (1970): Ultrastructural features of blastocyst attachment and trophoblastic invasion in the rat. J. Reprod. Fertil., 21:37-56. 27. Parkening, T. A. (1979): Apposition of uterine luminal epithelium during implantation in senescent golden hamster. J. Gerontol., 34:335344. 28. Martin, L., Finn, C. A., and Carter, J. (1970): Effects of proges¬ terone and oestradiol on the luminal epithelium of the mouse uterus. J. Reprod. Fertil., 21:461-469. 29. Hedlund, K., and Nilsson, O. (1971): Hormonal requirements for the uterine attachment reaction and blastocyst implantation in the mouse, hamster and guinea-pig. J. Reprod. Fertil., 26:267-269. 30. Pollard, R. M., and Finn, C. A. (1972): Ultrastructure of the uterine epithelium during the hormonal induction of sensitivity and insen¬ sitivity to a decidual stimulus in the mouse. J. Endocrinol., 55:293298. 31. McLaren, A. (1968): A study of blastocysts during delay and sub¬ sequent implantation in lactating mice. J. Endocrinol., 42:453-463. 32. Lundkvist, O., Nilsson, B. O., and Bergstrom, S. (1979): Studies on the trophoblast-epithelial complex during decidual induction in rats. Am. J. Anat., 154:211-230. 33. Warren, R., and Enders, A. C. (1964): An electron microscope study of the rat endometrium during delayed implantation. Anat. Rec., 148:177-195. 34. Ljungkvist, I. (1972): Attachment reaction of rat uterine luminal epithelium. IV. The cellular changes in the attachment reaction and its hormonal regulation. Fertil. Steril., 23:847-865. 35. Tachi, S., and Tachi, C. (1979): Ultrastructural studies on maternalembryonic cell interactions during experimentally induced implan¬ tation of rat blastocysts to the endometrium of the mouse. Dev. Biol., 68:203-223. 36. McLaren, A., and Nilsson, O. (1971): Electron microscopy of lu¬ minal epithelium separated by beads in the pseudopregnant mouse uterus. J. Reprod. Fertil., 26:379-381. 37. Enders, A. C., and Nelson, D. M. (1973): Pinocytotic activity of the uterus of the rat. Am. J. Anat., 138:277-300. 38. Leroy, F., Van Hoeck, J., and Bogaert, C. (1976): Hormonal control of pinocytosis in the uterine epithelium of the rat. J. Reprod. Fertil., 47:59-62. 39. Parr, M. R., and Parr, E. L. (1974): Uterine luminal epithelium: Protrusions mediate endocytosis, not apocrine secretion, in the rat. Biol. Reprod., ll;:220-233. 40. Lundkvist, O. (1979): Morphometric estimation of stromal edema during delayed implantation in the rat. Cell Tissue Res., 199:339348. 41. Yochim, J. M., and Saldarini, R. J. (1969): Glucose utilization by the myometrium during early pseudopregnancy in the rat. J. Reprod. Fertil., 20:481-489. 42. Murphy, C. R., Swift, J. G., Mukherjee, T. M., and Rogers, A. W. (1982): Changes in the fine structure of the apical plasma membrane of endometrial epithelial cells during implantation in the rat. J. Ceil Sci., 55:1-12. 43. Parr, M. (1982): Apical vesicles in rat uterine epithelium during early pregnancy: A morphometric study. Biol. Reprod., 26:915-924. 44. Pollard, R. M., and Finn, C. A. (1974): Influence of the trophoblast upon differentiation of the uterine epithelium during implantation in the mouse. J. Endocrinol., 62:669-674.

45. Enders, A. C., Hendrickx, A. G., and Schlafke, S. (1983): Implan¬ tation in the rhesus monkey: Initial penetration of endometrium. Am. J. Anat., 167:275-298. 46. Parr, M. B., and Parr, E. L. (1982): Relationship of apical domes in the rabbit Uterine epithelium during the peri-implantation period to endocytosis, apocrine secretion and fixation. J. Reprod. Fertil., 66:739-744. 47. Bjorkman, N. (1973): Fine structure of the fetal-maternal area of exchange in the epitheliochorial and endotheliochorial types of placentation. Acta Anat., 86(Suppl.), 61:1-22. 48. Boshier, D. P. (1969): A histological and histochemical examination of implantation and early placentome formation in sheep. J. Reprod. Fertil., 19:51-61. 49. Wathes, D. C., and Wooding, F. B. P. (1980): An electron micro¬ scopic study of implantation in the cow. Am. J. Anat., 159:285-306. 50. Enders, A. C., and Schlafke, S. (1969): Cytological aspects of trophoblast-uterine interaction in early implantation. Am. J. Anat., 125:130. 51. Trinkaus, J. P. (1984): Cell adhesion. III. Mechanisms. In: Cells Into Organs: The Forces That Shape the Embryo, pp. 120-178. Prentice-Hall, Englewood Cliffs, N.J. 52. Enders, A. C., and Schlafke, S. (1974): Surface coats of the mouse blastocyst and uterus during the preimplantation period. Anat. Rec., * 180:31-46. 53. Anderson, T. L., and Hoffman, L. H. (1984): Alterations in epithelial glycocalyx of rabbit uteri during early pseudopregnancy and preg¬ nancy, and following ovariectomy. Am. J. Anat., 171:321-334. 54. Hewitt, K., Beer, A. E., and Grinnell, F. (1979): Disappearance of anionic sites from the surface of the rat endometrial epithelium at the time of blastocyst implantation. Biol. Reprod., 21:691-707. 55. Chavez, D. J., and Anderson, T. L. (1985): The glycocalyx of the mouse uterine luminal epithelium during estrus, early pregnancy, the peri-implantation period and delayed implantation. I. Acquisition of ricinus communis I binding sites during pregnancy. Biol. Reprod., 32:1135-1142. 56. Enders, A. C., and Schlafke, S. (1972): Implantation in the ferret: Epithelial penetration. Am. J. Anat., 133:291-316. 57. Hakansson, S., and Sundkvist, K. G. (1975): Decreased antigenicity of mouse blastocysts after activation for implantation from experi¬ mental delay. Transplantation, 19:479-484. 58. Hakansson, S., Heyner, S., Sundqvist, K-G., and Bergstrom, S. (1975): The presence of paternal H-2 antigens on hybrid mouse blastocysts during experimental delay of implantation and the dis¬ appearance of these antigens after onset of implantation. Int. J. Fertil., 20:137-140. 59. Nilsson, O., Lindqvist, I., and Ronquist, G. (1973): Decreased sur¬ face charge of mouse blastocysts at implantation. Exp. Cell Res., 83:421-423. 60. Nilsson, O., Lindqvist, I., and Ronquist, G. (1975): Blastocyst sur¬ face charge and implantation in the mouse. Contraception, 11:441450. 61. Jenkinson, E. J., and Searle, R. F. (1977): Cell surface changes on the mouse blastocyst at implantation. Exp. Cell Res., 106:386-390. 62. Clemetson, C. A. B., Moshfeghi, M. M., and Mallikarjuneswara, V. R. (1970): Electrophoretic mobility of the rat blastocyst. Con¬ traception, 1:357-360. 63. Nilsson, B. O., and Hjerten, S. (1982): Electrophoretic quantification of the changes in the average net negative charge density of mouse blastocysts implanting in vivo and in vitro. Biol. Reprod., 27:485493. 64. Enders, A. C., and Schlafke, S. (1971): Penetration of the uterine epithelium during implantation in the rabbit. Am. J. Anat., 132:219— 240. 65. Bergstrom, S., and Nilsson, O. (1976): Blastocyst attachment and early invasion during oestradiol-induced implantation in the mouse. Anat. Embryol., 149:149-154. 66. Konwinski, M., Vorbrodt, A., Solter, D., and Koprowski, H. (1977): Ultrastructural study of concanavalin-A binding to the surface of preimplantation mouse embryos. J. Exp. Zool., 200:311-324. 67. Chavez, D. J., and Enders, A. C. (1981): Temporal changes in lectin binding of peri-implantation mouse blastocysts. Dev. Biol 87 267276. 68. Chavez, D. J., and Enders, A. C. (1982): Lectin binding of mouse blastocysts: Appearance of dolichos biflorus binding sites on the

Biology of Implantation

trophoblast during delayed implantation and their subsequent dis¬ appearance during implantation. Biol. Reprod., 26:545-552. 69. Sobel, J. S., and Nebel, L. (1976): Concanavalin A agglutinability of the developing mouse trophoblast. J. Reprod. Fertil., 47 399402. 70. Sobel, J. S., and Nebel, L. (1978): Changes in concanavalin A agglutinability during development of the inner cell mass and tro¬ phoblast of mouse blastocysts in vitro. J. Reprod. Fertil., 52: 239-248. 71. Nilsson, B. O., Naeslund, G., and Curman, B. (1980): Polar dif¬ ferences of delayed and implanting mouse blastocysts in binding of alcian blue and concanavalin A. J. Exp. Zool., 214:177-180. 72. Carollo, J. R., and Weitlauf, H. M. (1981): Regional changes in the binding of [3H] concanavalin A to mouse blastocysts at implantation: An autoradiographic study. J. Exp. Zool., 218:247-251. 73. Pinsker, M. C., and Mintz, B. (1973): Change in cell-surface gly¬ coproteins of mouse embryos before implantation. Proc. Natl. Acad. Sci. USA, 70:1645-1648. 74. Surani, M. A. H. (1979): Glycoprotein synthesis and inhibition of glycosylation by tunicamycin in pre-implantation mouse embryos: Compaction and trophoblast adhesion. Cell, 18:217-227. 75. Sherman, M. I., Shalgi, R., Rizzino, A., Sellens, M. H., Gay, S., and Gay, R. (1979): Changes in the surface of the mouse blastocyst at implantation. In: Maternal Recognition of Pregnancy, CIBA Foun¬ dation Symposium No. 64 (new series), pp. 33-52. ExcerptaMedica, New York. 76. Zetter, B. R., and Martin, G. R. (1978): Expression of a high mo¬ lecular weight cell surface glycoprotein (LETS protein) by preim¬ plantation mouse embryos and teratocarcinoma cells. Proc. Natl. Acad. Sci. USA, 75:2324-2328. 77. Gulamhusein, A. P., and Beck, F. (1973): Light and electron mi¬ croscopic observations at the pre- and early post-implantation stages in the ferret uterus. J. Anat., 115:159-174. 78. Enders, A. C., and Schlafke, S. (1965): The fine structure of the blastocyst: Some comparative studies. In: Preimplantation Stages of Pregnancy, CIBA Foundation Symposium, pp. 29-59. Little, Brown, Boston. 79. Parr, E. L. (1973): Shedding of the zona pellucida by guinea pig blastocysts: An ultrastructural study. Biol. Reprod., 8:531-544. 80. Allen, W. R., Hamilton, D. W., and Moor, R. M. (1973): The origin of equine endometrial cups. II. Invasion of the endometrium by trophoblast. Anat. Rec., 177:485-502. 81. Finn, C. A., and Lawn, A. M. (1968): Transfer of cellular material between the uterine epithelium and trophoblast during the early stages of implantation. J. Reprod. Fertil., 15:333-336. 82. Wilson, I. B. (1963): A new factor associated with the implantation of the mouse egg. J. Reprod. Fertil., 5:281-282. 83. Enders, A. C., and Schlafke, S. (1979): Comparative aspects of blastocyst-endometrial interactions at implantation. In: Maternal Recognition of Pregnancy, CIBA Foundation Symposium No. 64 (new series), pp. 3-32. Excerpta Medica, New York. 84. Finn, C. A., and Hinchliffe, J. R. (1965): Histological and histochemical analysis of the formation of implantation chambers in the mouse uterus. J. Reprod. Fertil., 9:301-309. 85. Finn, C. A., and Hinchliffe, J. R. (1964): Reaction of the mouse uterus during implantation and deciduoma formation as demonstrated by changes in the distribution of alkaline phosphatase. J. Reprod. Fertil., 8:331-338. 86. Hinchliffe, J. R., and El-Shershaby, A. M. (1975): Epithelial cell death in the oil-induced decidual reaction of the pseudopregnant mouse: An ultrastructural study. J. Reprod. Fertil., 45:463-468. 87. El-Shershaby, A. M., and Hinchliffe, J. R. (1975): Epithelial au¬ tolysis during implantation of the mouse blastocyst: An ultrastructural study. J. Embryol. Exp. Morphol., 33:1067-1080. 88. Finn, C. A., and Bredl, J. C. S. (1973): Studies on the development of the implantation reaction in the mouse uterus: influence of actinomycin D. J. Reprod. Fertil., 34:247-253. 89. Moulton, B. C., and Elangovan, S. (1981): Lysosomal mechanisms in blastocyst implantation and early decidualization. In: Cellular and Molecular Aspects of Implantation, edited by S. R. Glasser and D. W. Bullock, pp. 335-344. Plenum Press, New York. 90. Roy, S. K., Sengupta, J., and Manchanda, S. K. (1983): Histochemical study of (3-glucuronidase in the rat uterus during implan¬ tation and pseudopregnancy. J. Reprod. Fertil., 68:161-164.

/

253

91. Sengupta, J., Paria, B. C., and Manchanda, S. K. (1981): Effect of an oestrogen antagonist on implantation and uterine leucylnaphthylamidase activity in the ovariectomized hamster. J. Reprod. Fertil., 62:437-440. 92. Van Hoorn, G., and Denker, H. W. (1975): Effect of the blastocyst on a uterine amino acid arylamidase in the rabbit. J. Reprod. Fertil., 45:359-362. 93. Boshier, D. P. (1976): Effects of the rat blastocyst on neutral lipids and nonspecific esterases in the uterine luminal epithelium at the implantation site. J. Reprod. Fertil., 46:245-247. 94. Moulton, B. C. (1974): Ovum implantation and uterine lysosomal enzyme activity. Biol. Reprod., 10:543-548. 95. Finn, C. A. (1982): Cellular changes in the uterus during the estab¬ lishment of pregnancy in rodents. J. Reprod. Fertil. (Suppl.), 31:105111. 96. Cowell, T. P. (1969): Implantation and development of the mouse eggs transferred to the uteri of non-progestational mice. J. Reprod. Fertil., 19:239-245. 97. Steer, H. W. (1970): The trophoblastic knobs of the preimplanted rabbit blastocyst: A light and electron microscopic study. J. Anat., 107:315-325. 98. Steer, H. W. (1971): Implantation of the rabbit blastocyst: The in¬ vasive phase. J. Anat., 110:445-462. 99. Larsen, J. F. (1961): Electron microscopy of the implantation site in the rabbit. Am. J. Anat., 109:319-334. 100. Psychoyos, A. (1960): La reaction deciduale est precedee de mod¬ ifications precoces de la permeabilite capillarie de l’uterus. C. R: Soc. Biol., 154:1384-1387. 101. Psychoyos, A. (1961): Permeabilite capillarie et decidualisation uter¬ ine. C. R. Acad. Sci. Paris, 252:1515-1517. 102. McLaren, A. (1969): Can mouse blastocysts stimulate a uterine re¬ sponse before losing the zona pellucida? J. Reprod. Fertil., 19:199201. 103. Loeb, L. (1907): Wounds of the pregnant uterus. Proc. Soc. Exp. Biol. Med., 4:93-96. 104. Loeb, L. (1908): The production of deciduomata and the relation between the ovaries and formation of the decidua. JAMA, 50:18971901. 105. Loeb, L. (1908): The experimental production of the maternal part of the placenta of the rabbit. Proc. Soc. Exp. Biol. Med., 5:102-105. 106. Deanesly, R. (1971): The differentiation of the decidua at ovo-implantation in the guinea-pig contrasted with that of the traumatic deciduoma. J. Reprod. Fertil., 26:91-97. 107. Lundkvist, O., and Nilsson, B. O. (1982): Endometrial ultrastructure in the early uterine response to blastocysts and artificial deciduogenic stimuli in rats. Cell Tissue Res., 225:355-364. 108. Welch, A. O., and Enders, A. C. (1985): Light and electron micro¬ scopic examination of the mature decidual cells of the rat with em¬ phasis on the antimesometrial decidua and its degeneration. Am. J. Anat., 172:1-29. 109. DeFeo, V. J. (1967): Decidualization. In: Cellular Biology of the Uterus, edited by R. M. Wynn, pp. 191-290, Appleton-CenturyCrofts, New York. 110. Long, I. A., and Evans, H. M. (1922): The oestrus cycle in the rat and its associated phenomena. Mem. Univ. Calif., 6:1-148. 111. Parkes, A. S. (1929): The functions of the corpus luteum. II. The experimental production of placentoma in the mouse. Proc. R. Soc. Lond. {Biol.), 104:183-188. 112. Kraicer, P. F., and Shelesnyak, M. C. (1959): Determination de la periode de sensibilite maximale de l’endometre a la decidualis¬ ation au moyen de deciduomes provoques par un traitement empruntant la voie vasculaire. C. R. Acad. Sci. Paris, 248:3213— 3215. 113. Shelesnyak, M. C., and Kracier, P. F. (1960): Time-limits of uterine sensitivity to decidualization during progestation. Proc. 1 stint. Congr. Endocrinol., (Copenh.): 547. 114. DeFeo, V. J. (1963): Determination of the sensitive period for the induction of deciduomata in the rat by different inducing procedures. Endocrinology, 73:488-497. 115. DeFeo, V. J. (1963): Temporal aspect of uterine sensitivity in the pseudopregnant or pregnant rat. Endocrinology, 72:305-316. 116. Hetherington, C. M. (1968): The development of deciduomata in¬ duced by two nontraumatic methods in the mouse. J. Reprod. Fertil., 17:391-393.

254

/

Chapter 7

117. Harper, M. J. K. (1969): Deciduomal response of the golden hamster uterus. Anat. Rec., 163:563-574. 118. Finn, C. A., and Martin, L. (1972): Endocrine control of the timing of endometrial sensitivity to a decidual stimulus. Biol. Reprod., 7:8296. 119. McLaren, A., and Michie, D. (1956): Studies on the transfer of fertilized mouse eggs to uterine foster-mothers. I. Factors affecting the implantation and survival of native and transferred eggs. J. Exp. Biol., 33:394-416. 120. Psychoyos, A. (1965): Control de la nidation chez les mammiferes. Arch. Anat. Micros. Morphol. Exp., 54:85-104. 121. Psychoyos, A. (1966): Etude des relations de l’oeuf et de l’endometre au cours du retard de la nidation ou des premieres phases du processes de la nidation chez la ratte. Compt. Rend. Acad. Sci. Paris Ser. D, 263:1755-1758. 122. Psychoyos, A. (1966): Recent researches of egg-implantation. In: Egg Implantation, CIBA Foundation Study Group 23, pp. 4-28. Churchill. London. 123. Psychoyos, A. (1969): Hormonal factors governing decidualization. Excerpta Med. Found. Int. Congr. Serv., 184:935-938. 124. Psychoyos, A. (1969): Hormonal requirements for egg implantation. In: Advances in BioSciences. IV. Mechanisms Involved in Concep¬ tion, edited by G. Raspe, pp. 275-290. Pergamon Press, London. 125. Noyes, R. W., and Dickmann, Z. (1960): Relationship of ovular age to endometrial development. J. Reprod. Fertil., 1:186-196. 126. Dickmann, Z., and Noyes, R. W. (1960): The fate of ova transferred into the uterus of the rat. J. Reprod. Fertil., 1:197-212. 127. Psychoyos, A. (1967): The hormonal interplay controlling egg-im¬ plantation in the rat. In: Advances in Reproductive Physiology, edited by A. McLaren, pp. 257-277. Logos Press, London. 128. Humphrey, K. W. (1967): The induction of implantation in the mouse after ovariectomy. Steroids, 10:591-600. 129. Shelesnyak, M. C. (1962): Decidualization: The decidua and the deciduoma. Perspect. Biol. Med., 5:503-518. 130. Finn, C. A., and Martin, L. (1967): Patterns of cell division in the mouse uterus during early pregnancy. J. Endocrinol., 39:593-597. 131. Zhinkin, L. N., and Samoshkina, N. A. (1967): DNA synthesis and cell proliferation during formation of deciduomata in mice. J. Embryol. Exp. Morphol., 17:593-605. 132. Hall, K. (1969): Uterine mitosis, alkaline phosphatase and adenosine triphosphatase during development and regression of deciduomata in pseudopregnant mice. J. Endocrinol., 44:91-100. 133. Leroy, F., Galand, P., and Chretien, J. (1969): The mitogenic action of ovarian hormones on the uterine and vaginal epithelium during the oestrus cycle in the rat: An autoradiographic study. J. Endo¬ crinol., 45:441-447. 134. Chaudhury, R. R., and Sethi, A. (1970): Effects of an intra-uterine contraceptive device on mitosis in the rat uterus on different days of pregnancy. J. Reprod. Fertil., 22:33-40. 135. Marcus, G. J. (1974): Mitosis in the rat uterus during the estrous cycle, early pregnancy and early pseudopregnancy. Biol. Reprod., 10:447-452. 136. Tachi, C., Tachi, S., and Lindner, H. R. (1972): Modification by progesterone of oestradiol-induced cell proliferation, RNA synthesis and oestradiol distribution in the rat uterus. J. Reprod. Fertil., 31:5976. 137. Mehrotra, S. N., and Finn, C. A. (1974): Cell proliferation in the uterus of the guinea-pig. J. Reprod. Fertil., 37:405-409. 138. Marcus, G. J. (1974): Hormonal control of proliferation in the guineapig uterus. J. Endocrinol., 63:89-97. 139. Schmidt, I. G. (1943): Proliferation in the genital tract of the normal mature guinea pig treated with colchicine. Am. J. Anat., 73:59-80. 140. Krueger, W. A., and Maibenco, H. C. (1972): DNA replication and cell division in the hamster uterus. Anat. Rec., 173:229-234. 141. Herken, R. (1983): Cell kinetics of early gestation mouse uterus. Cell Tissue Kinet., 16:419-428. 142. Galassi, L. (1968): Autoradiographic study of the decidual cell re¬ action in the rat. Dev. Biol., 17:75-84. 143. Leroy, F., and Galand, P. (1969): Radioautographic evaluation of mitotic parameters in the endometrium during the uterine sensitivity period in pseudopregnant rat. Fertil. Steril., 20:980-922. 144. O’Grady, J. E., and Heald, P. J. (1976): Uterine nucleic acid and phospholipid mfetabolism in the early stages of rat pregnancy. J. Endocrinol., 68:33P-34P.

145. Lobel, B.L., Levy, E., and Shelesnyak, M. C. (1967): Studies on the mechanism of nidation. XXXIV. Dynamics of cellular interac¬ tions during progestation and implantation in the rat. Acta Endocri¬ nol. (Suppl.), 123:7-109. 146. Clark, B. F. (1973): The effect of oestrogen and progesterone on uterine cell division and epithelial morphology in spayed-hypophysectomized rats. J. Endocrinol., 56:341-342. 147. Allen, E., Smith, G. M., and Gardner, W. U. (1937): Accentuation of the growth effect of Theelin on genital tissues of the ovariectomized mouse by arrest of mitosis with colchicine. Am. J. Anat., 61:321-341. 148. Perrotta, C. A. (1962): Initiation of cell proliferation in the vaginal and uterine epithelia of the mouse. Am. J. Anat., 111:195-204. 149. Epifanova, O. I. (1966): Mitotic cycles in estrogen-treated mice: An autoradiographic study. Exp. Cell Res., 42:562-577. 150. Martin, L., and Finn, C. A. (1968): Hormonal regulation of cell division in epithelial and connective tissues of the mouse uterus. J. Endocrinol., 41:363-371. 151. Finn, C. A., and Martin, L. (1973): Endocrine control of gland proliferation in the mouse uterus. Biol. Reprod., 8:585-588. 152. Martin, L., Finn, C. A., and Trinder, G. (1973): Hypertrophy and hyperplasia in the mouse uterus after oestrogen treatment: An au¬ toradiographic study. J. Endocrinol., 56:133-144. f53. Clark, B. F. (1971): The effects of oestrogen and progesterone on uterine cell division and epithelial morphology in spayed, adrenalectomized rats. J. Endocrinol., 50:527-528. 154; Lee, A., and Dukelow, W. R. (1972): Synthesis of DNA and mitosis in rabbit uteri after oestrogen and progesterone injections and during early pregnancy. J. Reprod. Fertil., 31:473-476. 155. Koseki, Y., and Fujimoto, G. I. (1974): Progesterone effects con¬ trasted with 17-3 estradiol on DNA synthesis in epithelial nuclear proliferation in the castrate rabbit uterus. Biol. Reprod., 10:596-604. 156. Nilsson, O. (1958): Ultrastructure of mouse uterine surface epithe¬ lium under different estrogenic influences. 1. Spayed animals and oestrus animals. J. Ultrastruct. Res., 1:375-396. 157. Nilsson, O. (1958): Ultrastructure of mouse uterine surface epithe¬ lium under different estrogenic influences. 3. Late effect of estrogen administered to spayed animals. J. Ultrastruct. Res., 2:185-199. 158. Nilsson, O. (1958): Ultrastructure of mouse uterine surface epithe¬ lium under different estrogenic influences. 2. Early effect of estrogen administered to spayed animals. J. Ultrastruct. Res.. 2:73-95. 159. Das, R. M. (1972): The effects of oestrogen on the cell cycle in epithelial and connective tissues of the mouse uterus. J. Endocrinol., 55:21-30. 160. Das, R. M. (1972): The time-course of the mitotic response to oes¬ trogen in the epithelium and stroma of the mouse uterus. J. Endo¬ crinol., 55:203-204. 161. Martin, L., Das, R. M., and Finn, C. A. (1973): The inhibition by progesterone of uterine epithelial proliferation in the mouse. J. En¬ docrinol., 57:549-554. 162. Das, R. M., and Martin, L. (1973): Progesterone inhibition of mouse uterine epithelial proliferation. J. Endocrinol., 59:205-206. 163. Martin, L., and Finn, C. A. (1969): Duration of progesterone treat¬ ment required for a stromal response to oestradiol-170 in the uterus of the mouse. J. Endocrinol., 44:279-280. 164. Finn, C. A., and Martin, L. (1970): The role of the oestrogen secreted before oestrus in the preparation of the uterus for implantation in the mouse. J. Endocrinol., 47:431-438. 165. Finn, C. A., and Martin, L. (1974): The control of implantation. J. Reprod. Fertil., 39:195-206. 166. Finn, C. A., Martin, L., and Carter, J. (1969): A refractory period following oestrogenic stimulation of cell division in the mouse uterus. J. Endocrinol., 44:121-126. 167. Talley, D. J., Tobert, J. A., Armstrong, E. G. Jr., and Villee, C. A. (1977): Changes in estrogen receptor levels during deciduomata development in the pseudopregnant rat. Endocrinology, 101:15381544. 168. Armstrong, E. G. Jr., Tobert, J. A., Talley, D. J., and Villee, C. A. (1977): Changes in progesterone receptor levels during decid¬ uomata development in the pseudopregnant rat. Endocrinology 101:1545-1551. 169. Logeat, F., Sartor, P., Vu Hai, M. T., and Milgrom, E. (1980): Local effect of the blastocyst on estrogen and progesterone receptors in the rat endometrium. Science, 207:1083-1085.

Biology of Implantation 170. Vu Hai, M. T., Logeat, F., and Milgrom, E. (1978): Progesterone receptors in the rat uterus: Variations in cytosol and nuclei during the oestrous cycle and pregnancy. J. Endocrinol., 76:43-48. 171. Martel, D., and Psychoyos, A. (1978): Progesterone-induced oes¬ trogen receptors in the rat uterus. J. Endocrinol., 76:145-154. 172. Clark, J. H., Markaverich, B., Upchurch, S., Eriksson, H., Hardin, J. W., and Peck, E. J. (1980): Heterogeneity of estrogen binding sites: Relationship to estrogen receptors and estrogen response. Re¬ cent Prog. Horm. Res., 36:89-134. 173. Do, Y. S., and Leavitt, W. W. (1978): Characterization of a specific progesterone receptor in decidualized hamster uterus. Endocrinology 102:443-451. 174. Martel, D., and Psychoyos, A. (1980): Behavior of uterine steroid receptors at implantation. In: Progress in Reproductive Biology, Vol. 7, edited by F. Leroy, C. A. Finn, A. Psychoyos, and P. O. Hubinont, pp. 216-233. Karger, New York. 175. Nilsson, O. (1959): Ultrastructure of mouse uterine surface epithe¬ lium under different estrogenic influences. 4. Uterine secretion. J. Ultrastruct. Res., 2:331-341. 176. Nilsson, B. O. (1974): Changes of the luminal surface of the rat uterus at blastocyst implantation: Scanning electron microscopy and ruthenium red staining. Z. Anat. Entwick., 144:337-342. 177. Alden, R. H. (1947): Implantation of the rat egg. II. Alterations in osmiophilic epithelial lipids of the rat uterus under normal and ex¬ perimental conditions. Anat. Rec., 97:1-19. 178. Elftman, H. (1958): Estrogen control of the phospholipids of the uterus. Endocrinology, 62:410—415. 179. Elftman, H. (1963): Estrogen induced changes in the Golgi apparatus and lipid of the uterine epithelium of the rat in the normal cycle. Anat. Rec., 146:139-143. 180. Fuxe, K., and Nilsson, O. (1963): The effect of oestrogen on the histology of the uterine epithelium of the mouse. Exp. Cell Res., 32:109-117. 181. Boshier, D. P., and Holloway, H. (1973): Effects of ovarian steroid hormones on histochemically demonstrable lipids in the rat uterine epithelium. J. Endocrinol., 56:59-67. 182. Hall, K. (1973): Lactic dehydrogenase and other enzymes in the mouse uterus during the peri-implantation period of pregnancy. J. Reprod. Fertil., 34:79-91. 183. Hall, K. (1975): Lipids in the mouse uterus during early pregnancy. J. Endocrinol., 65:233-243. 184. Enders, A. C. (1961): Comparative studies on the endometrium of delayed implantation. Anat. Rec., 139:483-497. 185. Smith, M. S. R., and Wilson, I. B. (1971): Histochemical obser¬ vations on early implantation in the mouse. J. Embryol. Exp. Morphol., 25:165-174. 186. Wood, C., and Psychoyos, A. (1967): Activite de certaines enzymes hydrolytiques dans 1 ’endometre et la myometre au cours de la pseu¬ dogestation et de divers etats de receptivite uterine chez la ratte. C. R. Acad. Sci. Paris, Ser. D, 265:141-144. 187. Bergstrom, S. (1972): Delay of blastocyst implantation in the mouse by ovariectomy or lactation. A scanning electron microscope study. Fertil. Steril., 23:548-561. 188. Bergstrom, S., and Nilsson, O. (1972): Ultrastructural response of blastocysts and uterine epithelium to progesterone deprivation during delayed implantation in mice. J. Endocrinol., 55:217-218. 189. Christie, G. A. (1966): Implantation of the rat embryo: Glycogen and alkaline phosphatases. J. Reprod. Fertil., 12:279-294. 190. Abraham, R., Hendy, R., Dougherty, W. J., Fulfs, J. C., and Golberg, L. (1970): Participation of lysosomes in early implantation in the rabbit. Exp. Mol. Pathol., 13:329-345. 191. Elangovan, S., and Moulton, B. C. (1980): Blastocyst implantation in the rat and the immunohistochemical distribution and rate of syn¬ thesis of uterine lysosomal cathespin D. Biol. Reprod., 23:663-668. 192. Moulton, B. C., and Ingle, C.B. (1981): Uterine lysosomal cathespin D activity, rate of synthesis and immunohistochemical localization following initiation of decidualization in pseudopregnant rats. Biol. Reprod., 25:393-398. 193. Moulton, B. C. (1982): Progesterone and estrogen control of the response of rat uterine lysosomal cathespin D activity to a deciduogenic stimulus. Endocrinology, 110:1197-1202. 194. Finn, C. A., and Martin, L. (1971): Endocrine control of the pro¬ liferation and secretion of uterine glands in the mouse. Acta Endo¬ crinol. (Suppl.), 155:139.

/

255

195. Enders, A. C., and Given, R. L. (1977): The endometrium of delayed and early implantation. In: Biology of the Uterus, edited by R. M. Wynn, pp. 203-243. Plenum Press, New York. 196. Given, R. L., and Enders, A. C. (1980): Mouse uterine glands during the peri-implantation period: Fine structure. Am. J. Anat., 157:169179. 197. Given, R. L., and Enders, A. C. (1978): Mouse uterine glands during the delayed and induced implantation periods. Anat. Rec., 190:271— 284. 198. Hooker, C. W., and Forbes, T. R. (1947): A bio-assay for minute amounts of progesterone. Endocrinology, 41:158-169. 199. Tachi, C., Tachi, S., and Lindner, H. R. (1974): Effects of ovarian hormones upon nucleolar ultrastructure in endometrial stromal cells of the rat. Biol. Reprod., 10:404-413. 200. Fainstat, T. (1963): Extracellular studies of uterus. I. Disappearance of the discrete collagen bundles in endometrial stroma during various reproductive states in the rat. Am. J. Anat., 112:337-370. 201. Miller, B. G., and Emmens, C. W. (1969): The effects of oestradiol and progesterone on the incorporation of tritiated uridine into the genital tract of the mouse. J. Endocrinol., 43:427-436. 202. Miller, B. G., Owen, W. H., and Emmens, C. W. (1968): The incorporation of tritiated uridine in the uterus and vagina of the mouse during early pregnancy. J. Endocrinol., 41:189-195. 203. Miller, B: G., Owen, W. H., and Emmens, C. W. (1968): Uridine incorporation in the rat genital tract during early pregnancy. J. En¬ docrinol., 42:351-352. 204. Heald, P. J., and O’Grady, J. E. (1970): The uptake of [3H] uridine into the nucleic acids of the rat uterus during early pregnancy. Biochem. J., 117:65-71. 205. O’Grady, J. E., Heald, P. J. and O’Hare, A. (1970): Incorporation of [3H] uridine into the ribonucleic acid of rat uterus during pseu¬ dopregnancy and in the presence of I.C.I. 46474 [trans-l-(p-$dimethylaminoethoxyphenyl)-1,2-diphenylbut-1 -ene]. Biochem. J., 119:609-613. 206. Heald, P. J., O’Grady, J. E., O’Hare, A., and Vass, M. (1972): Changes in uterine RNA during early pregnancy in the rat. Biochim. Biophys. Acta, 262:66-74. 207. Heald, P. J., O’Grady, J. E., and Moffat, G. E. (1972): The in¬ corporation of [3H] uridine into nuclear RNA in the uterus of the rat during early pregnancy. Biochim. Biophys. Acta, 281:347-352. 208. O’Grady, J. E., Moffat, G. E., McMinn, L., Vass, M. A., O’Hare, A., and Heald, P. J. (1975): Uterine chromatin template activity during the early stages of pregnancy in the rat. Biochim. Biophys. Acta, 407:125-132. 209. Heald, P. J., O’Grady, J. E., O’Hare, A., and Vass, M. (1975): Nucleic acid metabolism of cells of the luminal epithelium and stroma of the rat uterus during early pregnancy. J. Reprod. Fertil., 45:129-138. 210. Kaye, A. M., Icekson, I., and Lindner, H. R. (1971): Stimula¬ tion by estrogens of omitinine and S-adenosylmethionine decar¬ boxylases in the immature rat uterus. Biochim. Biophys. Acta, 252: 150-159. 211. Saunderson, R., and Heald, P. J. (1974): Ornithine decarboxylase activity in the uterus of the rat during early pregnancy. J. Reprod. Fertil., 39:141-143. 212. Heald, P. J. (1979): Changes in ornithine decarboxylase during early implantation in the rat. Biol. Reprod., 20:1195-1199. 213. Reid, R. J., and Heald, P.J. (1970): Uptake of 3H-leucine into pro¬ teins of rat uterus during early pregnancy. Biochim. Biophys. Acta, 204:278-279. 214. Reid, R. J., and Heald, P. J. (1971): Protein metabolism of the rat uterus during the oestrous cycle, pregnancy and pseudopregnancy and as affected by an anti-implantation compound, ICI 46,474. J. Reprod. Fertil., 27:73-82. 215. Smith, J. A., Martin, L., King, R. J. B., and Vertes, M. (1970): Effects of oestradiol-17(3 and progesterone on total and nuclearprotein synthesis in epithelial and stromal tissues of the mouse uterus and of progesterone on the ability of these tissues to bind oestradiol17p. Biochem. J., 119:773-784. 216. Bell, S. C., Reynolds, S., and Heald, P. J. (1976): Presumptive induced protein synthesis in the rat uterus during early pregnancy. J. Endocrinol., 68:34p-35p. 217. Bell, S. C., Reynolds, S., and Heald, P. J. (1977): Uterine protein synthesis during the early stages of pregnancy in the rat. J. Reprod. Fertil., 49:177-181.

256

/

Chapter 7

218. Beall, J. R. (1972): Uterine lipid metabolism—A review of the lit¬ erature. Comp. Biochem. Physiol., 42B: 175-195. 219. Goswami, A., Kar, A. B., and Chowdhury, S. R. (1963): Uterine lipid metabolism in mice during the oestrous cycle: Effect of ovar¬ iectomy and replacement therapy. J. Reprod. Fertil., 6:287-295. 220. Aizawa, Y., and Mueller, G. C. (1961): The effect in vivo and in vitro of estrogens on lipid synthesis in the rat uterus. J. Biol. Chem., 236:381-3867 221. Davis, J. S., and Alden, R. H. (1959): Hormonal influence on lipid metabolism of rat uterus. Anat. Rec., 134:725-737. 222. Ray, S. C., and Morin, R. J. (1965): Lipid composition of the nongravid and gravid rabbit endometrium. Proc. Soc. Exp. Biol. Med., 120:849-853. 223. Morin, R. J., and Carrion, M. (1968): In vitro incorporation of acetate- 1-4C into the phospholipids of rabbit and human endometria. Lipids, 3:349-353. 224. Beall, J. R., and Werthessen, N. T. (1971): Lipid metabolism of the rat uterus after mating. J. Endocrinol., 51:637-644. 225. Yochim, J.M. (1971): Intrauterine oxygen tension during the preim¬ plantation period. In: Biology of the Blastocyst, edited by R. J. Blandau, pp. 363-382. University of Chicago Press, Chicago. 226. Battellino, L. J., Sabulsky, J., and Blanco, A. (1971): Lactate de¬ hydrogenase isoenzymes in rat uterus: Changes during pregnancy. J. Reprod. Fertil., 25:393-399. 227. Clark, S. W., and Yochim, J. M. (1971): Effect of ovarian steroids on lactic dehydrogenase activity in endometrium and myometrium of the rat uterus. Endocrinology, 89:358-365. 228. Clark, S. W., and Yochim, J. M. (1971): Lactic dehydrogenase in the rat uterus during progestation, its relation to intrauterine oxygen tension and the regulation of glycolysis. Biol. Reprod., 5:152-160. 229. Saldarini, R. J., and Yochim, J. M. (1967): Metabolism of the uterus of the rat during early pseudopregnancy and its regulation by estrogen and progesterone. Endocrinology, 80:453-466. 230. Surani, M. A. H., and Heald, P. J. (1971): The metabolism of glucose by rat uterus tissue in early pregnancy. Acta Endocrinol., 66:16-24. 231. Yochim, J. M., and Clark, S. W. (1971): Lactic dehydrogenase activity in the uterus of the rat during the estrous cycle and its relation to intrauterine oxygen tension. Biol. Reprod., 5:146-151. 232. Yochim, J.M., and Mitchell, J. A. (1968): Intrauterine oxygen ten¬ sion in the rat during progestation: Its possible relation to carbohy¬ drate metabolism and the regulation of nidation. Endocrinology, 83:706-713. 233. Yochim, J. M., and Pepe, G. J. (1971): Effect of ovarian steroids on nucleic acids, protein, and glucose-6-phosphate dehydrogenase activity in endometrium of the rat; a metabolic role for progesterone in “progestational differentiation.” Biol. Reprod., 5:172-182. 234. Mallonee, R. C., and Yochim, J. M. (1980): Uterus of the rat dur¬ ing progestation: Pyridine nucleotide activity and its relation to preimplantation changes in pentose cycle activity. Biol. Reprod., 23: 588-594. 235. Yochim, J. M., and Mallonee, R. C. (1980): Hormonal control of pyridine nucleotide activity in the uterus: A model for progestational differentiation. Biot. Reprod., 23:595-605. 236. Pepe, G. J., and Yochim, J. M. (1971): Pentose cycle activity in endometrium of the rat during progestation: Its regulation by intra¬ uterine oxygen and its relation to the “progestational” action of pro¬ gesterone. Endocrinology, 89:366-377. 237. Cummings, A. M., and Yochim, J. M. (1983): Nicotinamide adenine dinucleotide in rat uterus: Role of progesterone in the regulation of preimplantation differentiation. Endocrinology, 112:1407-1411. 238. Cummings, A.M., and Yochim, J. M. (1983): Nicotinamide adenine dinucleotide kinase in the rat uterus: Regulation by progesterone and decidual induction. Endocrinology, 112:1412-1419. 239. Yochim, J. M. (1984): Modulation of uterine sensitivity to decidual induction in the rat by nicotinamide: Challenge and extension of a model of progestational differentiation. Biol. Reprod., 30:637-645. 240. Krehbiel, R. H. (1937): Cytological studies of the decidual reaction in the rat during pregnancy and in the production of deciduomata. Physiol. Zool., 10:212-238. 241. Velardo, J. T., Dawson, A. B., Olsen, A. G., and Hisaw, F. L. (1953): Sequence of histological changes in the uterus and vagina of the rat during prolongation of pseudopregnancy associated with the presence of deciduomata. Am. J. Anat., 93:273-305.

242. Lobel, B. L., Tic, L., and Shelesnyak, M. C. (1965): Studies on the mechanisms of nidation. XVII. Histochemical analysis of decidualization in the rat. Part 2. Induction. Acta Endocrinol., 50:469-485. 243. Lobel, B. L., Tic, L., and Shelesnyak, M. C. (1965): Studies on the mechanism of nidation. XVII. Histochemical analysis of decidualization in the rat. Part 3. Formation of the deciduomata. Acta Endocrinol., 50:517-536. 244. Ledford, B. E., Rankin, J. C., Froble, V. L., Serra, M. J., Markwald, R. R., and Baggett, B. (1978): The decidual cell reaction in the mouse uterus: DNA synthesis and autoradiographic analysis of responsive cells. Biol. Reprod., 18:506-509. 245. Finn, C. A., and McLaren, A. (1967): A study of the early stages of implantation in mice. J. Reprod. Fertil., 13:259-267. 246. Sachs, L., and Shelesnyak, M. C. (1955): The development and suppression of polyploidy in the developing and suppressed deciduoma in the rat. J. Endocrinol., 12:146-151. 247. Dupont, H., Duluc, J. A., and Mayer, G. (1971): Evolution cytologique et genese de la polyploidie dans le deciduome experimental chez la ratte en gestation unilateral. C. R. Acad. Sci. Paris Ser. D, 272:2360. 248. Ansell, J. D., Barlow, P. W., and McLaren, A. (1974): Binucleate and polyploid cells in the decidua of the mouse. J. Embryol. Exp. Morphol., 31:223-227. 249. Jollie, W., and Benscome, S. A. (1965): Electron microscopic ob¬ servations on primary decidua formation in the rat. Am. J. Anat., 116:217-236. 250. Lundkvist, O., and Ljungkvist, I. (1977): Morphology of the rat endometrial stroma at the appearance of the pontamine blue reaction during implantation after an experimental delay. Cell Tissue Res. 184:453-466. 251. Parkening, T. A. (1976): An ultrastructural study of implantation in the golden hamster. III. Initial formation and differentiation of de¬ cidual cells. J. Anat., 122:485-498. 252. O’Shea, J. D., Kleinfeld, R. G., and Morrow, H. A. (1983): Ul¬ trastructure of decidualization in the pseudopregnant rat. Am J Anat., 166:271-298. 253. Abrahamsohn, P. (1983): Ultrastructural study of the mouse antimesometrial decidua. Anat. Embryol., 166:263-274. 254. Enders, A. C., Welsh, A. O., and Schlafke, S. (1985): Implantation in the rhesus monkey: Endometrial responses. Am. J. Anat., 173:147169. 255. Grinnell, F., Head, I. R., and Hoffpauir, J. (1982): Fibronectin and cell shape in vivo: Studies on the endometrium during pregnancy. J. Cell Biol., 94:597-606. 256. Finn, C. A., and Lawn, A. M. (1967): Specialized junctions between decidual cells in the uterus of the pregnant mouse. J. Ultrastruct. Res., 20:321-327. 257. Lawn, A. M., Wilson, E. W., and Finn, C. A. (1971): The ultra¬ structure of human decidual and predecidual cells. J. Reprod. Fertil., 26:85-90. 258. Kleinfeld, R., Morrow, H. A., and DeFeo, V. J. (1976): Intercellular junctions between decidual cells in the growing deciduoma of the pseudopregnant rat uterus. Biol. Reprod., 15:593-603. 259. Abrahamsohn, P., Lundkvist, O., and Nilsson, O. (1983): Ultra¬ structure of the endometrial blood vessels during implantation of the rat blastocyst. Cell Tissue Res., 229:269-280. 260. Milligan, S. R., and Mirembe, F. M. (1984): Time course of the changes in uterine vascular permeability associated with the devel¬ opment of the decidual cell reaction in ovariectomized steroid-treated rats. J. Reprod. Fertil., 70:1-6. 261. Hoos, P. C., and Hoffman, L. H. (1980): Temporal aspects of rabbit uterine vascular and decidual responses to blastocyst stimulation. Biol. Reprod., 23:453-459. 262. Das, R. M., and Martin, L. (1978): Uterine DNA synthesis and cell proliferation during early decidualization induced by oil in mice. J. Reprod. Fertil., 53:125-128. 263. Ledford, B. E., Rankin, J. C., Markwald, R. R., and Baggett, B. (1976): Biochemical and morphological changes following artificially stimulated decidualization in the mouse uterus. Biol. Reprod 15 529— 535. 264. Shelesnyak, M. C., and Tic, L. (1963): Studies on the mechanism of decidualization. IV. Synthetic processes in the decidualizing uterus. Acta Endocrinol., 42:465-472.

Biology of Implantation

265. Shelesnyak, M. C., and Tic, L. (1963): Studies on the mechanism of decidualization. V. Suppression of synthetic processes of the uterus (DNA, RNA, and protein) following inhibition of decidualization by an antioestrogen, ethanoxytriphetol (MER-25). Acta Endocrinol 43:462-463. 266. Leroy, F., Bogaert, C., vanHoeck, J., and Delcroix, C. (1974): Cytophotometric and autoradiographic evaluation of cell kinetics in decidual cell growth in rats. J. Reprod. Fertil., 38:441-449. 267. Moulton, B. C., and Blaha, G. C. (1978): Separation of deciduomal cells by velocity sedimentation at unit gravity. Biol. Reprod , 18141147. 268. Moulton, B. C. (1979): Effect of progesterone on DNA, RNA and protein synthesis of deciduoma cell fractions separated by velocity sedimentation. Biol. Reprod., 21:667-672. 269. Moulton, B. C., and Koenig, B. B. (1984): Uterine deoxyribonucleic acid synthesis during preimplantation in precursors of stromal cell differentiation during decidualization. Endocrinology, 115:1302-1307. 270. Martel, D., and Psychoyos, A. (1981): Estrogen receptors in the nidatory sites of the rat endometrium. Science, 211:1454-1455. 271. Ward, W. F., Frost, A. G., and Orsini, M. W. (1978): Estrogen binding by embryonic and interembryonic segments of the rat uterus prior to implantation. Biol. Reprod., 18:598-601. 272. Sartor, P. (1977): Exogenous hormone uptake and retention in the rat uterus at the time of ova-implantation. Acta Endocrinol., 84:804812. 273. Moulton, B. C., and Koenig, B. B. (1981): Estrogen receptor in deciduoma cells separated by velocity sedimentation. Endocrinology, 108:484-488. 274. McConnell, K. N., Sillar, R. G., Young, B. D., and Green, B. (1982): Ploidy and progesterone-receptor distribution in flow-sorted deciduomal nuclei. Mol. Cell. Endocrinol., 25:99-104. 275. Miller, B. G. (1973): Metabolism of RNA and pyrimidine nucleotides in the uterus during the early decidual cell reaction. J. Endocrinol., 59:275-283. 276. Serra, M. J., Ledford, B. E., Rankin, J. C., and Baggett, B. (1978): Changes in RNA polymerase activity in isolated mouse uterine nu¬ clei during the decidual cell reaction. B iochim. Biophys. Acta, 521:267273. 277. Barkai, U., and Kraicer, P. F. (1978): Definition of period of in¬ duction of deciduoma in the rat using ornithine decarboxylase as a marker of growth onset. Int. J. Fertil., 23:106-111. 278. Collawn, S. S., Rankin, J., Ledford, B. E., and Baggett, B. (1981): Ornithine decarboxylase activity in the artificially stimulated decidual cell reaction in the mouse uterus. Biol. Reprod., 24:528-533. 279. Tarachand, U., Sivabalan, R., and Eapen, J. (1980): Protein ana¬ bolism in endometrium and myometrium during the growth of in¬ duced deciduoma in rats. Experientia, 36:1154-1156. 280. Yoshinaga, K. (1972): Rabbit antiserum to rat deciduoma. Biol. Reprod., 6:51-57. 281. Yoshinaga, K. (1974): Interspecific cross-reactivity of deciduoma antiserum: Interaction between mouse deciduoma and anti-serum to rat deciduoma. Biol. Reprod., 11:50-55. 282. Joshi, S. G., Szarowski, D. H., and Bank, J. (1981): Deciduaassociated antigens in the baboon. Biol. Reprod., 25:591-598. 283. Sacco, A. G., and Mintz, B. (1975): Mouse uterine antigens in the implantation period of pregnancy. Biol. Reprod., 12:498-503. 284. Denari, J. H., and Rosner, J. M. (1978): Studies on biochemical char¬ acteristics of an early decidual protein. Int. J. Fertil., 23:123-127. 285. Umapathesivam, K., and Jones, W. R. (1978): An investigation of decidual specific proteins in the rat. Int. J. Fertil., 23:138-142. 286. Denari, J. H., Germino, N. I., and Rosner, J. M. (1976): Early synthesis of uterine proteins after a decidual stimulus in the pseu¬ dopregnant rat. Biol. Reprod., 15:1-8. 287. Bell, S. C. (1979): Synthesis of’decidualization-associated protein’ in tissues of the rat uterus and placenta during pregnancy. J. Reprod. Fertil., 56:255-262. 288. Bell, S. C. (1979): Protein synthesis during deciduoma morphogen¬ esis in the rat. Biol. Reprod., 20:811-821. 289. Bell, S. C. (1979): Immunochemical identity of decidualizationassociated proteins and a2 acute-phase macroglobulin in the pregnant rat. J. Reprod. Immunol., 1:193-206. 290. Bell, S. C., Hamer, J., and Heald, P. J. (1980): Induced protein and deciduoma formation in rat uterus. Biol. Reprod., 23:935-940.

/

257

291. Notides, A., and Gorski, J. (1966): Estrogen-induced synthesis of a specific uterine protein. Proc. Natl. A.cad. Sci. USA., 56:230-235. 292. Manak, R., Wertz, N., Slabaugh, M., Denari, H., Wang, J. T., and Gorski, J. (1980): Purification and characterization of the estrogeninduced protein (IP) of the rat uterus. Mol. Cell. Endocrinol., 17:119— 132. 293. Katzenellenbogen, B. S. (1975): Synthesis and inducibility of the uterine estrogen-induced protein, IP, during the rat estrous cycle: Clues to uterine estrogen sensitivity. Endocrinology, 96: 289-297. 294. Dupont-Mairess, N., and Galand, P. (1975): Estrogen action: In¬ duction of the synthesis of a specific protein (IP) in the myometrium, the stroma and the luminal epithelium of the rat uterus. Endocri¬ nology, 96:1587-1591. 295. Lejeune, B. Lecocq, R., Lamy, F., and Leroy, F. (1982): Changes in the pattern of endometrial protein synthesis during decidualization in the rat. J. Reprod. Fertil., 66:519-523. 296. MacDonald, R. G., Morency, K. O., and Leavitt, W. W. (1983): Progesterone modulation of specific protein synthesis in the decidualized hamster uterus. Biol. Reprod., 28:753-766. 297. Leavitt, W. W., MacDonald, R. G., and Shwaery, G. T. (1985): Characterization of deciduoma proteins in hamster uterus: Detection in decidual cell cultures. Biol. Reprod., 32:631-643. 298. Shelesnyak, M. C. (1952): Inhibition of decidual cell formation in the pseudopregnant rat by histamine antagonists. Am. J. Physiol., 170:522-527. 299. Shelesnyak, M. C. (1957): Some experimental studies on the mech¬ anism of ovo-implantation in the rat. Recent Prog. Horm. Res., 13:269-317. 300. Kraicer, P. F., and Shelesnyak, M. C. (1958): The induction of deciduomata in the pseudopregnant rat by systemic administration of histamine and histamine releasers. J. Endocrinol., 17:324-328. 301. Shelesnyak, M. C. (1959): Fall in uterine histamine associated with ovum implantation in pregnant rat. Proc. Soc. Exp. Biol. Med., 100:380-381. 302. Spaziani, E., and Szego, C. M. (1958): The influence of estradiol and cortisol on uterine histamine of the ovariectomized rat. Endo¬ crinology, 63:669-678. 303. Spaziani, E., and Szego, C. M. (1959): Further evidence for me¬ diation by histamine of estrogenic stimulation of the rat uterus. En¬ docrinology, 64:713-723. 304. Humphrey, K. W., and Martin, L. (1968): Attempted induction of deciduomata in mice with mast-cell, capillary permeability and tissue inflammatory factors. J. Endocrinol., 42:129-141. 305. Wrenn, T. R., Bitman, J., Cecil, H. C., and Gilliam, D. R. (1964): Uterine deciduomata: Role of histamine. J. Endocrinol., 28:149152. 306. Finn, C. A., and Keen, P. M. (1962): Influence of systemic anti¬ histamines on formation of deciduoma. J. Endocrinol., 24:381-382. 307. Goldstein, A., and Hazel, M. M. (1955): Failure of an antihistamine drug to prevent pregnancy in the mouse. Endocrinology, 56:215216. 308. Harper, M. J. K. (1965): Failure of various antihistaminic drugs to prevent implantation in rats. J. Reprod. Fertil., 9:359-361. 309. Finn, C. A., and Keen, P. M. (1962): Failure of histamine to induce deciduomata in the rat. Nature, 194:602-603. 310. Banik, U. K., and Ketchel, M. M. (1964): Inability of histamine to induce deciduomata in pregnant rats and pseudopregnant rats. J. Reprod. Fertil., 7:259-261. 311. Brandon, J. M., and Wallis, R. M. (1977): Effect of mepyramine, a histamine Hr, and burinamide, a histamine H2-receptor antagonist, on ovum implantation in the rat. J. Reprod. Fertil., 50:251-254. 312. Dey, S. K., Villanueva, C., and Abdou, N. 1. (1979): Histamine receptors on rabbit blastocyst and endometrial cell membranes. Na¬ ture, 278:648-649. 313. Brandon, J. M., and Raval, P. J. (1979): Interaction of estrogen and histamine during ovum implantation in the rat. Eur. J. Pharmacol 57:171-177. 314. Brandon, J. M. (1980): Some recent work on the role of histamine in ovum implantation. In: Blastocyst-Endometrium Relationships, Progress in Reproductive Biology, Vol 7, edited by F. Leroy, C. A. Finn, A. Psychoyos, and P. Hubinont, pp. 244-252. S. Karger, New York.

258

/

Chapter 7

315. Ferrando, G., and Nalbandov, A. V. (1968): Relative importance of histamine and estrogen on implantation in rats. Endocrinology, 83:933937. 316. Dey, S. K., Villanueva, C., Chien, S. M., and Crist, R. D. (1978): The role of histamine in implantation in the rabbit. J. Reprod. Fertil., 53:23-26. 317. Kennedy, T. G. (1983): Embryonic signals and the initiation of blastocyst implantation. Aust. J. Biol. Sci., 36:531-543. 318. Kennedy, T. G., and Armstrong, D. T. (1981): The role of pros¬ taglandins in endometrial vascular changes at implantation. In: Cel¬ lular and Molecular Aspects of Implantation, edited by S. R. Glasser and D. W. Bullock, pp. 349-363, Plenum Press, New York. 319. Lau, I. F., Saksena, S. K., and Chang, M. C. (1973): Pregnancy blockade by indomethacin, an inhibitor of prostaglandin synthesis: Its reversal by prostaglandins and progesterone in mice. Prostaglan¬ dins, 4:795-803. 320. Saksena, S. K., Lau, I. F., and Chang, M. C. (1976): Relationship between oestrogen, prostaglandin F2a and histamine in delayed im¬ plantation in the mouse. Acta Endocrinol., 81:801-807. 321. Lundkvist, O., and Nilsson, B. O. (1980): Ultrastructural changes of the trophoblast-epithelial complexes in mice subjected to implan¬ tation blocking treatment with indomethacin. Biol. Reprod., 22:719726. 322. Holmes, P. V., and Gordashko, B. J. (1980): Evidence of prosta¬ glandin involvement in blastocyst implantation. J. Embryol. Exp. Morphol., 55:109-122. 323. Gavin, M. A., Dominguez Femandez-Tejerina, J. C., Montanes de las Heras, M. F., and Vijil Maeso, E. (1974): Efectos de un inhibidor de la biosintesis de las prostaglandinas (indometacina) sobre la implantacion en la rata. Reproduccion, 1:177-186. 324. Kennedy, T. G. (1977): Evidence for a role for prostaglandins in the initiation of blastocyst implantation in the rat. Biol. Reprod., 16:286291. 325. Phillips, C. A., and Poyser, N. L. (1981): Studies on the involvement of prostaglandin in implantation in the rat. J. Reprod. Fertil., 62:7381. 326. Evans, C. A., and Kennedy, T. G. (1978): The importance of pros¬ taglandin synthesis for the initiation of blastocyst implantation in the hamster. J. Reprod. Fertil., 54:255-261. 327. Hoffman, L. H. (1978): Antifertility effects of indomethacin during early pregnancy in the rabbit. Biol. Reprod., 18:148-153. 328. Kennedy, T. G. (1979): Prostaglandins and increased endometrial vascular permeability resulting from the application of an artificial stimulus to the uterus of the rat sensitized for the decidual cell re¬ action. Biol. Reprod., 20:560-566. 329. Hoffman, L. H., DiPietro, D.L., and McKenna, T. J. (1978): Effects of indomethacin on uterine capillary permeability and blastocyst de¬ velopment in rabbits. Prostaglandins, 15:823-829. 330. Hoos, P. C., and Hoffman, L. H. (1983): The effect of histamine receptor antagonists and indomethacin on implantation in the rabbit. Biol. Reprod., 29:833-840. 331. Castracane, V. D., Saksena, S. K., and Shaikh, A. A. (1974): Effect of IUDs, prostaglandins and indomethacin on decidual cell reaction in the rat. Prostaglandins, 6:397-404. 332. Sananes, N., Baulieu, E. E., and le Goascogne, C. (1976): Pros¬ taglandin^) as inductive factor of decidualization in the rat uterus. Mol. Cell. Endocrinol., 6:153-160. 333. Tobert, J. A. (1976): A study of the possible role for prostaglandins in decidualization using a nonsurgical method for the instillation of fluids into the rat uterine lumen. J. Reprod. Fertil., 47:391-393. 334. Rankin, J. C., Ledford, B. E., Jonsson, H. T., and Baggett, B. (1979): Prostaglandins, indomethacin and the decidual cell reaction in the mouse uterus. Biol. Reprod., 20:399-404. 335. Miller, M. M., and O’Morchoe, C. C. C. (1982): Decidual cell reaction induced by prostaglandin F2a in the mature oophorectomized rat. Cell Tissue Res., 225:189-199. 336. Kennedy, T. G., and Lukash, L. A. (1982): Induction of decidual¬ ization in rats by the intrauterine infusion of prostaglandins. Biol. Reprod., 27:253-260. 337. Kennedy, T. G. (1980): Timing of uterine sensitivity for the decidual cell reaction: Role of prostaglandins. Biol. Reprod., 22:519-525. 338. Kennedy, T. G. (1980): Estrogen and uterine sensitization for the decidual cell reaction: Role of prostaglandins. Biol. Reprod., 23:955962.

339. Kennedy, T. G., and Zamecnik, J. (1978): The concentration of 6keto-prostaglandin F)a is markedly elevated at the site of blastocyst implantation in the rat. Prostaglandins, 16:599-605. 340. Jonsson, H. T., Rankin, J. C., Ledford, B. E., and Baggett, B. (1979): Uterine prostaglandin levels following stimulation of the decidual cell reaction: Effects of indomethacin and tranylcypromine. Prostaglandins, 18:847-857. 341. Hoffman, L. H., Davenport, G. R., and Brash, A. R. (1984): En¬ dometrial prostaglandins and phospholipase activity related to im¬ plantation in rabbits: Effects of dexamethasone. Biol. Reprod., 30:544555. 342. Miller, M. M., and O’Morchoe, C. C. C. (1982): Inhibition of artificially induced decidual cell reaction by indomethacin in the mature oopherectomized rat. Anat. Rec., 204:223-230. 343. Hoffman, L. H., Strong, G. B., Davenport, G. R., and Frolich, J. C. (1977): Deciduogenic effect of prostaglandins in the pseudo¬ pregnant rabbit. J. Reprod. Fertil., 50:231-237. 344. Kennedy, T. G., Martel, D., and Psychoyos, A. (1983): Endometrial prostaglandin E2 binding: Characterization in rats sensitized for de¬ cidual cell reaction and changes during pseudopregnancy. Biol. Re¬ prod., 29:556-564. 345. Martel, D., Kennedy, T. G., Monier, M. N., and Psychoyos, A. _ (1985): Failure to detect specific binding sites for prostaglandin F2a in membrane preparations from rat endometrium. J. Reprod. Fer¬ til., 75:265-274. 346. Kennedy, T. G., Martel, D., and Psychoyos A. (1983): Endometrial prostaglandin E2 binding during the estrous cycle and its hormonal control in ovariectomized rats. Biol. Reprod., 29:565-571. 347. Peleg, S., and Lindner, H. R. (1982): The effect of prostaglandins on progestin receptor translocation and on decidual cell reaction in vivo and in vitro. Endocrinology, 110:1647-1652. 348. Leroy, F., Vansande, J., Shetgen, G., and Brasseur, D. (1974): Cyclic AMP and triggering of the decidual reaction. J. Reprod. Fertil. 39:207-211. 349. Rankin, J. C., Ledford, B. E., and Baggett, B. (1977): Early involvement of cyclic nucleotides in the artificially stimulated decidual cell reaction in the mouse uterus. Biol. Reprod., 17: 549-554. 350. Kennedy, T. G. (1983): Prostaglandin E2, adenosine 3' : 5'-cyclic monophsphate and changes in endometrial vascular permeability in rat uterus sensitized for decidual cell reaction. Biol. Reprod., 29:10691076. 351. Johnston, M. E. A., and Kennedy, T. G. (1984): Estrogen and uterine sensitization for the decidual cell reaction in the rat: Role of pros¬ taglandin E2 and adenosine 3' : 5'-cyclic monophosphate. Biol. Re¬ prod., 31:959-966. 352. Alleua, J. J., Kenimer, J. G., Jordan, A. W., and Lamanna, C. (1983): Induction of estrogen and progesterone receptors and decid¬ ualization in the hamster uterus of cholera toxin. Endocrinology, 11:2095-2106. 353. Webb, F. T. G. (1975): The inability of dibutyryl adenosine 3',-5'monophosphate to induce the decidual reaction in intact pseudo¬ pregnant mice. J. Reprod. Fertil., 42:187-188. 354. Webb, F. T. G. (1975): Implantation in ovariectomized mice treated with dibutyryl adenosine 3',5'-monophosphate (dibutyryl cyclic AMP). J. Reprod. Fertil., 42:511-517. 355. Holmes, P. V., and Bergstrom, S. (1975): Induction of blastocyst implantation in mice by cyclic AMP. J. Reprod. Fertil., 43:329332. 356. Heap, R. B., Flint, A. P. F., Gadsby, J. E., and Rice, C. (1979): Hormones, the early embryo and the uterine environment. J. Reprod Fertil., 55:267-275. 357. Flint, A. P. F., Burton, R. D., Gadsby, J. E., Saunders, P. T. K., and Heap, R. B. (1979): Blastocyst oestrogen synthesis and the maternal recognition of pregnancy. In: Maternal Recognition of Preg¬ nancy, CIBA Foundation Symposium No. 64 (new series), pp. 209238. Excerpta Medica, New York. 358. Atkinson, L. E., Hotchkiss, J., Fritz, G. R., Surve, A. H., Neill, J. D., and Knobil, E. (1975): Circulating levels of steroids and chorionic gonadotropin during pregnancy in the rhesus monkey, with special attention to the rescue of the corpus luteum in early pregnancy. Biol. Reprod., 12:335-345. 359. Blandau, R. J. (1949): Embryo-endometrial interrelationship in the rat and guinea pig. Anat. Rec., 104:331-360.

Biology of Implantation

360. Alden, R. H., and Smith, M. J. (1959): Implantation of the rat egg. IV. Some effects of artificial ova on the uterus. J. Exp. Zool., 142:215— 226. 361. McLaren, A. (1968): Can beads stimulate a decidual response in the mouse uterus? J. Reprod. Fertil., 15:313-315. 362. Cole, R. J. (1967): Cinemicrographic observations on the trophoblast and zona pellucida of the mouse blastocyst. J. Embryol. Exp. Morphol., 17:481-490. 363. Bitton-Casimiri, V., Bran, J-L., and Psychoyos, A. (1970): Comportement in vitro des blastocysts du 5e jour de la gestation chez la ratte; etude microcinematographique. C. R. Acad. Sci. Paris Ser. D, 270:2979-2982. 364. Lejeune, B., Van Hoeck, J., and Leroy, F. (1981): Transmitter role of the luminal uterine epithelium in the induction of decidualization in rats. J. Reprod. Fertil., 61:235-240. 365. Boving, B. G. (1959): Implantation. Ann. N.Y. Acad. Sci., 75:700725. 366. Boving, B. G. (1963): Implantation mechanisms. In: Mechanisms Concerned with Conception, edited by C. G. Hartman, pp. 321— 396. Pergamon Press, New York. 367. Hetherington, C. M. (1968): Induction of deciduomata in the mouse by carbon dioxide. Nature, 219:863-864. 368. Huff, R. L., and Eik-Nes, K. B. (1966): Metabolism in vitro of acetate and certain steroids by six-day-old rabbit blastocysts. J. Re¬ prod. Fertil, 11:57-63. 369. Dickmann, Z., and Dey, S. K. (1973): Two theories: The preim¬ plantation embryo is a source of steroid hormones controlling (1) morala-blastocyst transformation, and (2) implantation. J. Reprod. Fertil., 35:615-617. 370. Heap, R. B., Flint, A. P. F., and Gadsby, J. E. (1981): Embryonic signals and maternal recognition. In: Cellular and Molecular Aspects of Implantation, edited by S. R. Glasser and D. W. Bullock, pp. 311-325. Plenum Press, New York. 371. Sauer, M. J. (1979): Hormone involvement in the establishment of pregnancy. J. Reprod. Fertil., 56:725-743. 372. Perry, J. S., Heap, R. B., and Amoroso, E. C. (1973): Steroid hormone production by pig blastocysts. Nature, 245:45-47. 373. Perry, J. S., Heap, R. B., Burton, R. D., and Gadsby, J. E. (1976): Endocrinology of the blastocyst and its role in the establishment of pregnancy. J. Reprod. Fertil. (Suppl.), 25:85-104. 374. Gadsby, J. E., Heap, R. B., and Burton, R. D. (1980): Oestrogen production by blastocyst and early embryonic tissue of various spe¬ cies. J. Reprod. Fertil., 60:409-417. 375. Gadsby, J. E., Burton, R. D., Heap, R. B., and Perry, J. S. (1976): Steroid metabolism and synthesis in early embryonic tissue of pig, sheep, and cow. J. Endocrinol., 7L45P-46P. 376. Fischer, H. E., Bazer, F. W., and Fields, M. J. (1985): Steroid metabolism by endometrial and conceptus tissues during early pregnancy and pseudopregnancy in gilts. J. Reprod. Fertil., 75:69-78. 377. Flood, D. F. (1974): Steroid-metabolizing enzymes in the early pig conceptus and in the related endometrium. J. Endocrinol., 63:413— 414. 378. Heap, R. B., Flint, A. P. F., Hartmann, P. E., Gadsby, J. E., Staples, L. D., Ackland, N., and Hamon, M. (1981): Oestrogen production in early pregnancy. J. Endocrinol., 89:77p-94p. 379. Dhindsa, D. S., and Dziuk, P. J. (1968): Effect of pregnancy in the pig after killing embryos or fetuses in one uterine hom in early gestation. J. Anim. Sci., 27:122-126. 380. Bazer, F. W., and Thatcher, W. W. (1977): Theory of maternal recognition of pregnancy in swine based on estrogen controlled en¬ docrine versus exocrine secretion of prostaglandin F2a by uterine endometrium. Prostaglandins, 14:397-401. 381. Frank, M., Bazer, F. W., Thatcher, W. W., and Wilcox, C. J. (1977): A study of prostaglandin F2 as the leuteolysin in swine. III. Effects of estradiol valerate on prostaglandin F, progestins, estrone, and estradiol concentrations in the utero-ovarian vein of nonpregnant gilts. Prostaglandins, 14:1183-1196. 382. Moeljono, M. P. E., Thatcher, W. W., Bazer, F. W., Frank, M., Owens, L. J., and Wilcox, C. J. (1977): A study of prostaglandin F2a as the leuteolysin in swine. II. Characterization and comparison of prostaglandin F, estrogens and progestin concentrations in uteroovarian vein plasma of nonpregnant and pregnant gilts. Prostaglan¬ dins, 14:543-555.

/

259

383. Zavy, M. T., Bazer, F. W., Thatcher, W. W., and Wilcox, C. J. (1980): A study of prostaglandin F2_a as the leuteolysin in swine. V. Comparison of prostaglandin F, progestins, estrone and estradiol in uterine flushings from pregnant and nonpregnant gilts. Prostaglan¬ dins, 20:837-851. 384. Heap, R. B., Perry, J. S., Gadsby, J. E., and Burton, K. D. (1975): Endocrine activities of the blastocyst and early embryonic tissue in the pig. Biochem. Soc. Trans., 3:1183-1188. 385. Mondschein, J. S., Hersey, R. M., Dey, S. K., Davis, D. L., and Weisz, J. (1985): Catechol estrogen formation by pig blastocyst during the preimplantation period: Biochemical characterization of estrogen-2/4-dioxylase and correlation with asromatase activity. En¬ docrinology, 117:2339-2346. 386. Hoversland, R. C., Dey, S. K., and Johnson, D. C. (1982): Catechol estradiol induced implantation in the mouse. Life Sci., 30:18011804. 387. Pakrasi, P. L., and Dey, S. K. (1983): Catechol estrogens stimulate synthesis of prostaglandin in the preimplantation rabbit blastocyst and endometrium. Biol. Reprod., 29:347-354. 388. Stone, B. A., and Seamark, R. F. (1985): Steroid hormones in uterine washings and in plasma of gilts between days 9 and 15 after oestrus and between days 9 and 15 after coitus. J. Reprod. Fertil., 75:209-

221. 389. Dickmann, Z., Dey, S. K., and Sengupta, J. (1975): Steroidogenesis in rabbit preimplantation embryos. Proc. Natl. Acad. Sci. USA, 72:298-300. 390. Dickmann, Z., Dey, S. K., and Sen Gupta, J. (1976): A new concept: Control of early pregnancy by steroid hormones originating in the preimplantation embryo. Vitam. Horm., 34:215-242. 391. Bleau, G. (1981): Failure to detect A5-3p-hydroxysteroid oxidoreductase activity in the preimplantation rabbit embryo. Steroids, 37:121— 132. 392. Seamark, R. F., and Lutwak-Mann, C. (1972): Progestins in rabbit blastocysts. J. Reprod. Fertil., 29:147-148. 393. Fuchs, A. R., and Beling, C. (1974): Evidence for early ovarian recognition of blastocysts in rabbits. Endocrinology, 95:1054-1058. 394. Borland, R. M., Erickson, G. F., and Ducibella, T. (1977): Accu¬ mulation of steroids in rabbit preimplantation blastocysts. J. Reprod. Fertil., 49:219-224. 395. Singh, M. M., and Booth, W. D. (1978): Studies on the metabo¬ lism of neutral steroids by preimplantation rabbit blastocysts in vitro and the origin of blastocyst oestrogen. J. Reprod. Fertil., 53:297-304. 396. Fujimoto, S., and Sundaram, K. (1978): The source of progesterone in rabbit blastocysts. J. Reprod. Fertil., 52:231-233. 397. Bullock, D. W. (1977): Steroids from the pre-implantation blasto¬ cyst. In: Development in Mammals, Vol. 2, edited by M. H. Johnson, pp. 199-208. North-Holland, New York. 398. George, F. W., and Wilson, J. D. (1978): Estrogen formation in the early rabbit embryo. Science, 199:200-201. 399. Hoversland, R. C., Dey, S. K., and Johnson, D. C. (1982): Aromatase activity in the rabbit blastocyst. J. Reprod. Fertil., 66:259263. 400. Wu, J-T., and Lin, G. M. (1982): Effect of aromatase inhibitor on oestrogen production in rabbit blastocysts. J. Reprod. Fertil., 66:655662. 401. Bhatt, B. M., and Bullock, D. W. (1974): Binding of oestradiol to rabbit blastocysts and its possible role in implantation. J. Reprod. Fertil., 39:65-70. 402. Dey, S. K., Dickmann, Z., and Sen Gupta, J. (1976): Evidence that the maintenance of early pregnancy in the rabbit requires “blastocyst estrogen.” Steroids, 28:481-485. 403. Sengupta, J., Roy, S. K., and Manchanda, S. K. (1979): Hormonal control of implantation: A possible role of lysosomal function in the embryo-uterus interaction. J. Steroid Biochem., 11:729-744. 404. Wise, T., and Heap, R. B. (1983): Effects of the embryo upon endometrial estrogen synthesis in the rabbit. Biol. Reprod., 28:10971106. 405. Dey, S. K., and Dickmann, Z. (1974): Estradiol-17(l-hydroxysteroid dehydrogenase activity in preimplantation rat embryos. Steroids, 24:5762. 406. Dey, S. K., and Dickmann, Z. (1974): A5-3(3-Hydroxysteroid de¬ hydrogenase activity in rat embryos on days 1 through 7 of pregnancy. Endocrinology, 95:321-322.

260

/

Chapter 7

407. Dickmann, Z., and Dey, S.K. (1974): Steroidogenesis in the preim¬ plantation rat embryo and its possible influence on morula-blastocyst transformation and implantation. J. Reprod. Fertil., 37:91-93. 408. Dickmann, Z., and Dey, S. K. (1974): Evidence that A5-3p-Hydroxysteroid dehydrogenase activity in rat blastocysts is autonomous. J. Endocrinol., 61:513-514. 409. Dickmann, Z., and Sengupta, J. (1974): A5-33-Hydroxysteroid de¬ hydrogenase activity in preimplantation hamster embryos. Dev. Biol., 40:196-198. 410. Dickmann, Z., Sengupta, J., and Dey, S. S. (1977): Does ‘blastocyst estrogen’ initiate implantation? Science, 195:687-688. 411. Sengupta, J., Dey, S. K., and Dickmann, Z. (1977): Is mouse preim¬ plantation embryogenesis controlled by estrogen originating in the preimplantation embryo? Anat. Rec., 187:709 (abstract). 412. Sengupta, J., Roy, S. K., and Manchanda, S. K. (1981): Effect of an anti-oestrogen on implantation of mouse blastocysts. J. Reprod. Fertil., 62:433-436. 413. Chew, N. J., and Sherman, M. I. (1975): Biochemistry of differ¬ entiation of mouse trophoblast: A\3p-Hydroxy steroid dehydrogen¬ ase. Biol. Reprod., 12:351-359. 414. Sherman, M. I., and Atienza, S. B. (1977): Production and metab¬ olism of progesterone and androstenedione by cultured mouse blas¬ tocysts. Biol. Reprod., 16:190-199. 415. Marcal, J. M., Chew, N. J., Salomon, D. S., and Sherman, M. I. (1975): A5,3-Hydroxysteroid dehydrogenase activities in rat tropho¬ blast and ovary during pregnancy. Endocrinology, 96:1270-1279. 416. Antila, E., Koskinen, J., Niemela, P., and Saure, A. (1977): Steroid metabolism by mouse preimplantation embryos in vitro. Experientia, 33:1374-1375. 417. Niimura, S., and Ishida, K. (1976): Histochemical studies of A53(3-, 20a- and 203-hydroxy steroid dehydrogenases and possible pro¬ gestagen production in hamster eggs. J. Reprod. Fertil., 48:275278. 418. Brodie, A. M. H., Wu, J. T., Marsh, D. A., and Brodie, H. J. (1978): Aromatase inhibitors. III. Studies on the antifertil¬ ity effect of 4-acetoxy-4-androstene-3,17-dione. Biol. Reprod., 18:365-370. 419. Evans, C. A., and Kennedy, T. G. (1980): Blastocyst implantation in ovariectomized, adrenalectomized hamsters treated with inhibitors of steroidogenesis during the pre-implantation period. Steroids, 36:4152. 420. Zavy, M. T., Mayer, R., Vernon, M. W., Bazer, F. W., and Sharp, D. C. (1979): An investigation of the uterine lumenal environment of non-pregnant and pregnant pony mares. J. Reprod. Fertil., (Suppl.), 27:403-411. 421. Dey, S. K., Johnson, D. C., and Santos, J. G. (1979): Is histamine production by the blastocyst required for implantation in the rabbit? Biol. Reprod., 21:1169-1173. 422. Dey, S. K., and Johnson, D. C. (1980): Histamine formation by mouse preimplantation embryos. J. Reprod. Fertil., 60:457-460. 423. Dey, S. K. (1981): Role of histamine in implantation: Inhibition of histidine decarboxylase induces delayed implantation in the rabbit. Biol. Reprod., 24:867-869. 424. Johnson, D. C., and Dey, S. K. (1980): Role of histamine in im¬ plantation: Dexamethasone inhibits estradiol-induced implantation in the rat. Biol. Reprod., 22:1136-1141. 425. Racowsky, C., and Biggers, J. D. (1983): Are blastocyst prosta¬ glandins produced endogenously? Biol. Reprod., 29:379-388. 426. Dickmann, Z., and Spilman, C. H. (1975): Prostaglandins in rabbit blastocysts. Science, 190:997-998. 427. Dey, S. K., Chien, S. M., Cox, C. L., and Crist, R. D. (1980): Prostaglandin synthesis in the rabbit blastocyst. Prostaglandins, 19:449453. 428. Harper, M. J., Norris, C. J., andRajkumar, K. (1983): Prostaglandin release by zygotes and endometria of pregnant rabbits. Biol. Reprod., 28:350-362. 429. Pakrasi, P. L., and Dey, S. K. (1982): Blastocyst is the source of prostaglandins in the implantation site in the rabbit. Prostaglandins, 24:73-77. 430. Shemesh, M., Milaguir, F., Ayalon, N., and Hansel, W. (1979): Steroidogenesis and prostaglandin synthesis by cultured bovine blas¬ tocysts. J. Reprod. Fertil., 56:181-185. 431. Lewis, G. S., Thatcher, W. W., Bazer, F. W., and Curl, J. S. (1982): Metabolism of arachidonic acid in vitro by bovine blastocysts and endometrium. Biol. Reprod., 27:431-439.

432. Biggers, J. D., Leonov, B. V., Baskar, J. F,, and Fried, J. (1978): Inhibition of hatching of mouse blastocysts in vitro by prostaglandin antagonists. Biol. Reprod., 19:519-533. 433. Baskar, J. F., Torchiana, D. F., Biggers, J. D., Corey, E. J., An¬ dersen, N. H , and Subramanian, N. (1981): Inhibition of hatching of mouse blastocysts in vitro by various prostaglandin antagonists. J. Reprod. Fertil., 63:359-363. 434. Hurst, P. R., and MacFarlane, D. W. (1981): Further effects of nonsteroidal anti-inflammatory compounds on blastocyst hatching in vitro and implantation rates in the mouse. Biol. Reprod., 25:777784. 435. Biggers, J. D., Baskar, J. F., and Torchiana, D. F. (1981): Reduction of fertility of mice by the intrauterine injection of prostaglandin antagonists. J. Reprod. Fertil., 63:365-372. 436. Marcus, G. J. (1981): Prostaglandin formation by the sheep embryo and endometrium as indication of maternal recognition of pregnancy. Biol. Reprod., 25:56-64. 437. Hyland, J. H., Manns, J. G., and Humphrey, W. D. (1982): Pros¬ taglandin production by ovine embryos and endometrium in vitro. J. Reprod. Fertil., 65:299-304. 438. Lacroix, M. C., and Kann, G. (1982): Comparative studies of pros¬ taglandins F2a and E2 in late cyclic and early pregnant sheep: in vitro synthesis by endometrium and conceptuses. Effects of in vivo in* domethacin treatment on establishment of pregnancy. Prostaglan¬ dins, 23:507-526. 439. Moor, R. M., andRowson, L. E. A. (1964): Influence of the embryo and uterus on luteal function in the sheep. Nature 201:522-523. 440. Moor, R. M., and Rowson, L. E. A. (1966): The corpus luteum of the sheep: Functional relationship between the embryo and the corpus luteum. J. Endocrinol., 34:233-239. 441. Moor, R. M., and Rowson, L. E. A. (1966): The corpus luteum of the sheep: Effect of the removal of embryos on luteal function. J. Endocrinol., 34:497-502. 442. Rowson, L. E. A., and Moor, R. M. (1967): The influence of embryonic tissue homogenate infused into the uterus, on the life span of the corpus luteum in the sheep. J. Reprod. Fertil., 13:511516. 443. Martal, J., Lacroix, M.-C., Loudes, C., Saunier, M., and Winterberger-Torres, S. (1979): Trophoblastin, an antiluteolytic protein present in early pregnancy in sheep. J. Reprod. Fertil., 56:63-73. 444. Heyman, Y., Camous, S., Fevre, J., Meziou, W., and Martal, J. (1984): Maintenance of the corpus luteum after uterine transfer of trophoblastic vesicles to cyclic cows and ewes. J. Reprod. Fertil., 70:533-540. 445. Godkin, J. D., Bazer, F. W., Moffatt, J., Sessions, F., and Roberts, R. M. (1982): Purification and properties of a major, low molecular weight protein released by the trophoblast of sheep blastocysts at day 13-21. J. Reprod. Fertil., 65:141-150. 446. Godkin, J. D., Bazer, F. W., and Roberts, R. M. (1984): Ovine trophoblast protein 1, an early secreted blastocyst protein, binds specifically to uterine endometrium and affects protein synthesis. Endocrinology, 114:120-130. 447. Hansen, P. J., Anthony, R. V., Bazer, F. W., Baumbach, G. A., and Roberts, R. M. (1985): In vitro synthesis and secretion of ovine trophoblastic protein-1 during the period of maternal recognition of pregnancy. Endocrinology, 117:1424-1430. 448. Godkin, J. D., Bazer, F. W., Thatcher, W. W., and Roberts, R. M. (1984): Proteins released by cultured day 15-16 conceptuses prolong luteal maintenance when introduced into the uterine lumen of cyclic ewes. J. Reprod. Fertil., 71:57-64. 449. Godkin, J. D., Cote, C., and Duby, R. T. (1978): Embryonic stim¬ ulation of ovine and bovine corpora lutea. J. Reprod. Fertil., 54:375378. 450. Ellinwood, W. E., Nett, T. M., and Niswender, G. D. (1979): Maintenance of the corpus luteum of early pregnancy in the ewe. I. Luteotropic properties of embryonic homogenates. Biol. Reprod 21:281-288. 451. Masters, R. A., Roberts, R. M., Lewis, G. S., Thatcher, W. W., Bazer, F. W., and Godkin, J. D. (1982): High molecular weight glycoproteins released by expanding, pre-attachment sheep, pig and cow blastocysts in culture. J. Reprod. Fertil., 66:571-583. 452. Betteridge, K. J., Eaglesome, M. D., Randall, G. C. B., Mitchell, D., and Sugden, E. A. (1978): Maternal progesterone levels as evi¬ dence of luteotrophic or antiluteolytic effects of embryos transferred to heifers 12-17 days after estrus. Theriogenology, 9:86.

Biology of Implantation

453. Sreeran, J. M. (1978): Non-surgical embryo transfer in the cow. Theriogenology, 9:69-83. 454. Northey, D. L., and French, L. R. (1980): Effect of embryo removal and intrauterine infusion of embryonic homogenates on the lifespan of the bovine corpus luteum. J. Anim. Sci., 50:298-302. 455. Bartol, F. F., Roberts, R. M., Bazer, F. W., Lewis, G. S., Godkin, J. D., and Thatcher, W. W. (1984): Characterization of proteins produced in vitro by periattachment bovine conceptus. Biol. Reprod., 32:681-693. 456. Bazer, F. W., Geisert, R. E., Thatcher, W. W., and Roberts, R. M. (1982): The establishment and maintenance of pregnancy. In: Control of Pig Reproduction, edited by J. A. ColeandG. R. Foxcroft, pp. 227-252. Butterworths, London. 457. Wyatt, C. (1976): Endometrial components involved in protein syn¬ thesis by 16-day pig blastocyst tissue in culture. J. Physiol. (Lond.), 260:73P-74P. 458. Saunders, P. T. K., Ziecik, A. J., and Flint, A. P. F. (1980): Gonadotrophin-like substance in pig placenta and embryonic mem¬ branes. J. Endocrinol., 85:25P. 459. Rice, C., Ackland, N., and Heap, R. B. (1981): Blastocystendometrial interaction and protein synthesis during preim¬ plantation development in the pig studied in vitro. Placenta, 2:129-142. 460. Godkin, J. D., Bazer, F. W., Lewis, G. S., Geisert, R. D., and Roberts, R. M. (1982): Synthesis and release of polypeptides by pig conceptuses during the period of blastocyst elongation and attach¬ ment. Biol. Reprod., 27:977-987. 461. Fishel, S. B., and Surani, M. A. H. (1980): Evidence for the synthesis and release of a glycoprotein by mouse blastocysts. J. Reprod. Fertil., 59:181-185. 462. Nieder, G. L., Weitlauf, H. M., and Hartman, M. (1986): Synthesis and secretion of stage specific proteins by peri-implantation mouse embryos. Biol. Reprod. 36:687-699. 463. Morton, H., Hegh, V., and Clunie, G. J. A. (1974): Immuno¬ suppression detected in pregnant mice by rosette inhibition test. Na¬ ture, 249:459-460. 464. Morton, H., Hegh, V., and Clunie, G. J. A. (1976): Studies of the rosette inhibition test in pregnant mice: Evidence of immunosuppres¬ sion? Proc. R. Soc. Lond. (Biol.) 193:413-419. 465. Morton, H., Rolfe, B., Clunie, G. J. A., Anderson, M. J., and Morrison, J. (1977): An early pregnancy factor detected in human serum by the rosette inhibition test. Lancet, i:394-397. 466. Nancarrow, C. D., Evison, B. M., Scaramuzzi, R. J., and Turnbull, K. E. (1979): Detection of induced death of embryos in sheep by the rosette inhibition test. J. Reprod. Fertil., 57:385389. 467. Noonan, F. P., Halliday, W. J., Morton, H., and Clunie, G. J. A. (1979): Early pregnancy factor is immunosuppressive. Nature, 278:649650. 468. Clarke, F. M., Morton, H., and Clunie, G. J. A. (1978): Detection and separation of two serum factors responsible for depression of lymphocyte activity in pregnancy. Clin. Exp. Immunol., 32:318— 323. 469. Clarke, F. M., Morton, H., Rolfe, B. E., and Clunie, G. J. A. (1980): Partial characterization of early pregnancy factor in the sheep. J. Reprod. Immunol., 2:151-162. 470. Wilson, S., McCarthy, R., and Clarke, F. (1983): In search of early pregnancy factor: Isolation of active polypeptides from pregnant ewe’s sera. J. Reprod. Immunol., 5:275-286. 471. Clarke, F. M., and Wilson, S. (1982): Biochemistry of early preg¬ nancy factor. In: Pregnancy Proteins, edited by J. G. Grudzinskas, B. Teisner, and M. Seppala, pp. 407-412. Academic Press, New York. 472. Morton, H., Rolfe, B., and Cavanagh, A. (1982): Early pregnancy factor: Biology and clinical significance. In: Pregnancy Proteins, edited by J. G. Grudzinskas, B. Teisner, and M. Seppala, pp. 391— 405. Academic Press, New York. 473. Morton, H., Rolfe, B. E., McNeill, L., Clarke, P., Clarke, F. M., and Clunie, G. J. A. (1980): Early pregnancy factor: Tissues in¬ volved in its production in the mouse. J. Reprod. Immunol., 2: 73-82. 474. Cavanagh, A. C., Morton, H., Rolfe, B. E., and Gidley-Baird, A. (1982): Ovum factor: A first signal of pregnancy? Am. J. Reprod. Immunol., 2:97-101.

/

261

475. Nancarrow, C. D., Wallace, A. L. C., and Grewal, A. S. (1981): The early pregnancy factor of sheep and cattle. J. Reprod. Fertil. (Suppl.), 30:191-199. 476. Cavanagh, A. C. (1984): Production in vitro of mouse early preg¬ nancy factor and purification to homogeneity. J. Reprod. Fertil., 71:581-592. All. Morton, H., and Clunie, G. J. A. (1979): A test for early pregnancy in sheep. Res. Vet. Sci., 26:261-262. 478. Morton, H., Nancarrow, C. D., Scaramuzzi, R. J., Evison, B. M., and Clunie, G. I. A. (1979): Detection of early pregnancy factor in sheep by rosette inhibition test. J. Reprod. Fertil., 56:75-80. 479. Morton, H., Tinnenberg, H. R., Rolfe, B., Wolf, M., and Mettler, L. (1982): Rosette inhibition test: A multicentre investigation of early pregnancy factor in humans. J. Reprod. Immunol., 4:251-261. 480. Smart, Y. C., Roberts, T. K., Clancy, R. L., and Cripps, A. W. (1981): Early pregnancy factor: Its role in mammalian reproduction research review. Fertil. Steril., 35:397-402. 481. Koh, L. Y., and Jones, W. R. (1982): The rosette inhibition test in early pregnancy diagnosis. Clin. Reprod. Fertil., 1:229-233. 482. Tinnenberg, H. R., Staves, R. P., and Semm, K. (1984): Improve¬ ment of the rosette inhibition assay for the detection of early preg¬ nancy factor in humans using the monoclonal antibody, anti-humanlyt-3. Am. J. Reprod. Immunol., 5:151-156. 483. Rolfe, B. E., Morton, H., Cavanagh, A. C., and Gardiner, R. A. (1983): Detection of an early pregnancy factor-like substance in sera of patients with testicular germ cell tumors. Am. J. Reprod. Im¬ munol., 3:97-100. 484. Koch, E., Morton, H., and Ellendorff, F. (1983): Early preg¬ nancy factor: Biology and practical application. Br. Vet. J., 139: 52-58. 485. Morton, H., Morton, D. J., and Ellendorff, F. (1983): The appear¬ ance and characteristics of early pregnancy factor in the pig. J. Reprod. Fertil., 69:437-446. 486. Cooper, D. W., and Aitken, R. J. (1981): Failure to detect altered rosette inhibition titres in human pregnancy serum. J. Reprod. Fertil., 61:241-245. 487. Whyte, A., and Heap, R. B. (1983): Early pregnancy factor. Nature, 304:121-122. 488. Smart, Y. C., Fraser, I. S., Clancy, R. L., Roberts, T. K., and Crippis, A. W. (1982): Early pregnancy factor as a monitor for fertilization in women wearing intrauterine devices. Fertil. Steril., 37:201-204. 489. Smart, Y. C., Roberts, T. K., Fraser, I. S., Cripps, A. W., and Clancy, R. L. (1982): Validation of rosette inhibition test for detec¬ tion of early pregnancy in women. Fertil. Steril., 37:779-785. 490. Shaw, F. D., and Morton, H. (1980): The immunological approach to pregnancy diagnosis: A review. Vet. Rec., 106:268-270. 491. Rolfe, B., Cavanagh, A., Forde, C., Bastin, C. C., and Morton, H. (1984): Modified rosette inhibition test with mouse lymphocytes for detection of early pregnancy factor in human pregnancy serum. J. Immunol. Methods, 70:1-11. 492. Rolfe, B. E., Morton, H., and Clarke, F. M. (1983): Early pregnancy factor is an immuno-suppressive contaminant of commercial prep¬ arations of human chorionic gonadotrophin. Clin. Exp. Immunol., 51:45-52. 493. Rolfe, B. E. (1982): Detection of fetal wastage. Fertil. Steril., 37:655660. 494. Kent, H. A. Jr. (1973): A polypeptide from oviductal contents which influences ovarian function. Biol. Reprod., 8:38-42. 495. Kent, H. A. Jr. (1975): Contraceptive polypeptide from hamster embryos: Sequence of amino acids in the compound. Biol. Reprod., 12:504-507. 496. Kent, H. A. Jr. (1975): The two to four-cell embryos as source tissue of the tetrapeptide preventing ovulations in the hamster. Am. J. Anat., 144:509-512. 497. Van Niekerk, C. H., and Gemeke, W. H. (1966): Persistence and pathologic cleavage of tubal ova in the mare. Ondesstepoort J. Vet. Res., 31:195-232. 498. Betteridge, K. J., and Mitchell, D. (1972): Retention of ova by the fallopian tube in mares. J. Reprod. Fertil., 31:515. 499. Nieder, G. L., and Corder, C. (1983): Pyruvate and lactate levels in oviducts of cycling, pregnant, and pseudopregnant mice. Biol. Reprod., 28:566-574. 500. O’Neill, C. (1985): Thrombocytopenia is an initial maternal response to fertilization in mice. J. Reprod. Fertil., 73:559-566.

262 / Chapter 7 501. Renfree, M. B., and Calaby, J. H. (1981): Background to delayed implantation and embryonic diapause. J. Reprod. Fertil. (Suppl.), 29:1-9. 502. Aitken, R. J. (1977): Embryonic diapause. In: Development in Mam¬ mals, Vol. 1, edited by M. H. Johnson, pp. 307-359. North-Holland, New York. 503. McLaren, A. (1973): Blastocyst activation. In: The Regulation of Mammalian Reproduction, edited by S. J. Segal, R. Crozier, P. A. Corfman, and P. G. Condliffe, pp. 321-328. Charles C Thomas, Springfield, Ill. 504. Brambell, F. W. R. (1937): The influence of lactation on implantation of the mammalian embryo. Am. J. Obstet. Gynecol., 33:942-953. 505. Weitlauf, H. M. (1973): In vitro uptake and incorporation of amino acids by blastocysts from intact and ovariectomized mice. J. Exp. Zool., 183:303-308. 506. Weitlauf, H. M. (1974): Effect of actinomycin D on protein synthesis by delayed implanting mouse embryos in vitro. J. Exp. Zool., 189:197-

202. 507. Weitlauf, H. M. (1976): Effect of uterine flushings on RNA synthesis by ‘implanting’ and ‘delayed implanting’ mouse embryos in vitro. Biol. Reprod., 14:566-571. 508. Weitlauf, H. M. (1978): Factors in mouse uterine fluid that inhibit the incorporation of [3H] uridine by blastocysts in vitro. J. Reprod. Fertil., 52:321-325. 509. Weitlauf, H. M. (1982); A comparison of the rates of accumulation of nonpolyadenylated and polyadenylated RNA in normal and de¬ layed implanting mouse embryos. Dev. Biol., 93:266-271. 510. Weitlauf, H. M. (1985): Changes in the rate of translation with reactivation of delayed implanting mouse embryos. J. Exp. Zool., 236:309-312. 511. Given, R. L., and Weitlauf, H. M. (1982): Resumption of DNA synthesis in delayed implanting mouse blastocysts during activation in vitro. J. Exp. Zool., 224:111-114. 512. Nieder, G. L., and Weitlauf, H. M. (1984): Regulation of glycolysis in the mouse blastocyst during delayed implantation. J. Exp. Zool., 231:121-129. 513. Nieder, G. L., and Weitlauf, H. M. (1985): Effects of metabolic substrates and ionic environment on in-vitro activation of delayed implanting mouse blastocysts. J. Reprod. Fertil., 73:151-157. 514. Torbit, C. A., and Weitlauf, H. M. (1975): Production of carbon dioxide in vitro by blastocysts from intact and ovariectomized mice. J. Reprod. Fertil., 42:45-50. 515. Weitlauf, H. M., and Kiessling, A. A. (1980): Comparison of overall rates of RNA synthesis in implanting and delayed implanting mouse blastocysts in vitro. Dev. Biol., 77:116-129. 516. Weitlauf, H. M., and Kiessling, A. A. (1981): Activation of‘delayed implanting’ mouse embryos in vitro. J. Reprod. Fertil., (Suppl.), 29:191-202. 517. Psychoyos, A., and Bitton-Casimiri, V. (1969): Captation in vitro d’un priecurseur d’acide ribonucleique (ARN)-(uridine 5-3H) par le bastocyste de rat: Differences entre blastocysts normaux et blastocysts en diapause. C. R. Soc. Biol. (Paris), 268:188-190. 518. Psychoyos, A., Bitton-Casimiri, V., and Brun, J. L. (1975) Repres¬ sion and activation of the mammalian blastocyst. In: Regulation and Differentiated Function in Eukaryote Cells, edited by G. P. Talwar, pp. 509-514. Raven Press, New York. 519. Mintz, B. (1964): Formation of genetically mosaic mouse embryos and early development of “lethal (/12/rl2)-normal” mosaics. J. Exp. Zool., 157:273-292. 520. Cole, R. J., and Paul, J. (1965): Properties of cultured preimplan¬ tation mouse and rabbit embryos and cell strains derived from them. In: Preimplantation Stages of Pregnancy, CIBA Foundation Sym¬ posium, pp. 82-111. Little, Brown, Boston.

521. Gwatkin, R. B. L. (1966): Defined media and development of mam¬ malian eggs in vitro. Ann. N.Y. Acad. Sci., 139:79-90. 522. Gwatkin, R. B. L. (1966): Amino acid requirements for attachment and outgrowth of the mouse blastocyst in vitro. J. Cell Physiol., 68:335-344. * 523. Naeslund, G. (1979): The effect of glucose-, arginine-, and leucinedeprivation on blastocyst outgrowth in vitro. Upsala J. Med. Sci., 84:9-20. 524. Naeslund, G., Lundkvist, O., and Nilsson, B. O. (1980): Trans¬ mission electron microscopy of mouse blastocysts activated and growth arrested in vivo and in vitro. Anat. Embryol., 159:33-48. 525. Wordinger, R. J., and Brinster, R.L. (1976): Influence of reduced glucose levels on the in vitro hatching, attachment and trophoblast outgrowth of the mouse blastocyst. Dev. Biol., 53:294-296. 526. Van Blerkom, J., Chavez, D. J., and Bell, H. (1979): Molecu¬ lar and cellular aspects of facultative delayed implantation in the mouse. In: Maternal Recognition of Pregnancy, CIBA Foundation Symposium No. 64 (new series), pp. 141-172. Excerpta Medica, New York. 527. Aitken, R. J. (1974): Delayed implantation in roe deer (Capreolus Capreolus). J. Reprod. Fertil., 39:225-233. 528. Aitken, R. J. (1974): Calcium and zinc in the endometrium and uterine flushings of the roe deer (Carreolus Carreolus) during de¬ layed implantation. J. Reprod. Fertil., 40:333-340. 529. Van Winkle, L. J. (1977): Low Na+ concentration: A factor con¬ tributing to diminished uptake and incorporation of amino acids by diapausing mouse blastocysts? J. Exp. Zool., 202:275-281. 530. Van Winkle, L. J. (1981): Activation of amino acid accumulation in delayed implantation mouse blastocysts. J. Exp. Zool., 218:239246. 531. Van Winkle, L. J., Campione, A. L., and Webster, D. I. (1983): Sodium ion concentrations in uterine flushings from “implanting” and “delayed implanting” mice. J. Exp. Zool., 226:321-324. 532. Surani, M. A. H. (1977): Cellular and molecular approaches to blastocyst uterine interactions at implantation. In: Development in Mammals, Vol. 1, edited by M. H. Johnson, pp. 245-305. NorthHolland, New York. 533. Gwatkin, R. B. L. (1969): Nutritional requirements for post-blas¬ tocyst development in the mouse. Int. J. Fertil., 14:101-105. 534. Nilsson, B. O., Magnusson, C., Widehn, S., and Hillensjo, T. (1982): Correlation between blastocyst oxygen consumption and trophoblast cytochrome oxidase reaction at initiation of implantation of delayed mouse blastocysts. J. Embryol. Exp. Morphol., 71:7582. 535. Menke, T. M., and McLaren, A. (1970): Carbon dioxide production by mouse blastocysts during lactational delay of implantation or after ovariectomy. J. Endocrinol., 47:287-294. 536. Aitken, R. J. (1977): The culture of mouse blastocysts in the presence of uterine flushings collected during normal pregnancy, delayed im¬ plantation, and pro-oestrus. J. Embryol. Exp. Morphol., 41:295300. 537. O’Neill, C., and Quinn, P. (1981): Interaction of uterine flushings with mouse blastocysts in vitro as assessed by the incorporation of [3H] uridine. J. Reprod. Fertil., 62:257-262. 538. O’Neill, C., and Quinn, P. (1983): Inhibitory influence of uterine secretions on mouse blastocysts decreases at the time of blastocyst activation. J. Reprod. Fertil., 68:269-274. 539. Psychoyos, A., and Casimiri, V. (1981): Uterine blastotoxic fac¬ tors. In: Cellular and Molecular Aspects of Implantation, edited by S. R. Glasser and D. W. Bullock, pp. 327-334. Plenum Press, New York.

THE REPRODUCTIVE SYSTEMS

The Female

It,

The Physiology of Reproduction edited by E. Knobil and J. Neill et al. Raven Press, Ltd., New York © 1988.

CHAPTER 8

Embryology of Mammalian Gonads and Ducts Anne Grete Byskov and Poul Erik H0yer Early Gonadal Formation, 265 Primordial Germ Cells, 265 • Mesonephros, 267 • Other Cell Types, 267 Gonadal Sex Determination, 269 Differentiation of the Ovary, 272 Different Patterns of Early Ovarian Differentiation, 272 • Meiotic Prophase, 273 • Formation of Follicles, 278 • Early Steroid-Producing Cells, 280 Differentiation of the Testis, 281 Testicular Cords, 281 • Prespermatogonia, 283 • Sertoli Cells, 284 • Leydig Cells, 286

Control of Meiosis, 288 Female Germ Cells, 289 • Male Germ Cells, 291

Gonadal formation in mammals takes place early in fetal life. Although the genetic sex is determined at conception, sexual differences between fetuses can first be recognized at the time when the gonads become sex differentiated. The gonads evolve when germ cells and different somatic cells migrate and settle in the gonadal ridges to interact in a finely regulated manner. The following differentiation of the em¬ bryonic gonads into an ovary or a testis is crucial for priming the fetus in the female or male direction. Inadequate sexual differentiation of the gonad may alter production of sex hormones and other substances necessary for growth and differentiation of the sex ducts, external genitalia, and other secondary sex characteristics, as well as sex priming of the

Primordial Germ Cells

Early Steroid Hormone Production by the Gonads, 291 Ovary, 291 • Testis, 291 Formation and Differentiation of Genital Ducts, 293 Undifferentiated Ducts, 293 • Female Differentiation of Genital Ducts, 294 • Male Differentiation of Genital Ducts, 294 Acknowledgments, 295 References, 295

For decades it was believed that the PGC originated in the so-called germinal epithelium, i.e., the coelomic epi¬ thelium lining the gonad. However, as early as 1880, Nussbaum (1) proposed an extragonadal origin of the PGC in frog and trout. Subsequently, PGC have also been identified in extragonadal sites in many mammalian species [rat (2,3), mouse (4-6), human (7,8), rabbit (9)]. The migratory pathway and, in particular, the precise origin of the PGC are difficult to establish since they are not easy to distinguish from surrounding cells. Different experiments using cultures of mouse embryonic fragments indicate that the stem cells of the PGC reside in the epiblast of the inner cell mass of the blastocyst (10,11). Studies of X-chromosome inactivation during mouse embryogenesis suggest that somatic cell lines are allocated prior to the germ cell line (12). However, labeling of four- to 16-cell mouse embryos with retroviruses and analyzing the distribution of proviruses in the different tissues developing from such embryos suggest that a germ cell line is set aside very early, maybe in the four- to eight-cell stage, and before allocation of somatic tissues (13). The sex of migrating primordial germ cells may be rec¬ ognized by their sex chromatin status. Generally, female

brain.

EARLY GONADAL FORMATION The gonads develop along the ventral cranial part of the mesonephros. Two simultaneously occurring events char¬ acterize the initial gonadal formation: (a) migration of pri¬ mordial germ cells (PGC) into the coelomic epithelium and the underlying mesenchymal tissue covering the mesone¬ phros and (b) release and migration of mesonephric cells into the same area.

265

266

/ Chapter 8

cells contain a chromatin body, the Barr body, which rep¬ resents the inactive X chromosome. Lyon (14) proposed that a dose compensation mechanism is achieved by Xchromosome inactivation in order to avoid the aneuploidy effect by the presence of more than one X chromosome. In somatic cells, inactivation occurs during embryonic life (15— 17). In female germ cells, one X chromosome is also in¬ activated during migration (18), but reactivation has been noticed by the time they reach the ovarian anlage (19-21). In oogonia of human and mouse, only one X chromosome is active (17,22), but reactivation occurs when germ cells enter leptonema (23,24). The mechanisms by which the PGC are translocated from extragonadal sites to the gonadal ridges are poorly under¬ stood. The mobility of PGC suggests that these cells may migrate actively (for review, see ref. 25). The guidance of oriented migration toward the gonads is also uncertain. It appears that fibronectin is present along the migratory path¬ way (26). The locomotion of PGC in vitro is also enhanced by fibronectin in the substrate (27).

A more passive translocation of PGC may occur during the morphogenic rearrangements of the developing tissues (11,28,29). Finally, it is possible that the PGC are attracted by chemotactic substances produced by the gonad (7) as visualized by time-lapse films of chick PGC in culture (30). This attractant does not seem to be class-specific because mouse germ cells settle in chick gonads when mouse hindgut containing germ cells are transplanted into the coelomic cavity of chick embryos (31). A useful tool for tracing and counting PGC is the cytochemical demonstration of their relatively high activity of alkaline phosphatase (32). Ultrastructural studies have shown that the reaction product is mainly localized to the plasmalemma. It is believed that this enzyme is involved in the transfer of metabolites across the cell surface (4). The al¬ kaline phosphatase-stained PGC have been counted from the time they are seen in the primitive streak until they reach The gonad and then go past the allantois, the hindgut, and the mesentery. In the 8-day-old mouse embryo, about 10 to 100 PGC can be identified (5,6,33). During migration

FIG. 1. Oogonium of a 9-week-old fetal human ovary. The plasmalemma forms a finger-like projection (ar¬ rowhead) with bundles of microfilaments (B). (A) x 5,800; (B) x 30,000.

Embryology of Mammalian Gonads and Ducts /

267

their number rapidly increases and by day 13 the gonads contain around 10,000 germ cells (34). In the 5-week-old human embryo the number of migrating germ cells is about 700 to 1,300 (7), and by week 8 the germ cell number of the developing gonad is 600,000 (35). When the PGC arrive at the coelomic epithelium covering the gonadal ridges they seem to be “trapped” by processes from the epithelial cells (36). Soon thereafter, PGC are present in the underlying tissue as well. The PGC of fetal human and pig gonads occasionally exhibit unique straight finger-like projections with closely packed parallel micro¬ filaments (Fig. 1), the function of which is unknown.

Mesonephros The mesonephros is the second of the three consecutive nephroic structures (pro-, meso-, and metanephros), which develop consecutively during fetal life of all mammals. All three kidneys arise in the nephrogenic cord, which forms from the segmented intermediate mesoderm early in em¬ bryonic life. The pronephros develops first from the most cranial segment, the mesonephros develops somewhat later from the intermediate segment, and finally the metanephros, the permanent kidney, arises from the most caudally placed one. The pronephros never functions in mammals, but the pronephric duct serves as an inductor for the formation of the mesonephros and the metanephros (37). The nephrons of mesonephros develop from the nephrogenic cord in a cranial caudal direction and successively form a connection with the pronephric duct, now called the Wolffian duct. In some species, e.g., pig, sheep, rabbit, and human, the mesonephros is a functioning kidney with well-developed glomeruli and tubuli (Fig. 2). However, in other species, e.g., guinea pig and mouse, the mesonephric tissue consists only of tubuli, which in some cases may develop Bowman’s capsules but without functional glomeruli (Figs. 3 and 4) (38). Although Waldeyer (39) and Balfour (40) proposed more than a century ago that the central cell mass of the gonad originated in the mesonephros (Fig. 5), this idea was vir¬ tually neglected until Witschi came to the same conclusion while studying amphibian (41) and mammalian gonads (42). Subsequently, numerous studies have lent support to this idea. For example, in the bovine fetus a broad stream of cells exhibiting strong alkaline phosphatase activity was observed to connect glomerular tuft and the developing gonad (43). Similar cell streams that connect the gonads with the mesonephric tissue have been described in other species, supporting the idea that the mesonephros and the gonads are closely interacting [human (44), sheep (45,46), rabbit (47,48), mouse (49-52)] (Figs. 2, 6, and 7). Different experiments have shown that the mesonephros influences gonadal development of function. When fetal undifferentiated mouse ovaries are stripped from mesone¬ phric tissues, ovarian differentiation and meiosis are pre-

FIG. 2. Ovary-mesonephric complex of an 11-week-old hu¬ man fetus. The ovary (o) is attached to the cranial part of the mesonephros (arrowheads). Caudally, mesonephric tubules (me), the Wolffian duct (w), and the Mullerian duct (m) are seen. x200. vented or inhibited when transplanted subcutaneously into nude mice (53). Also, the steroid synthesis by cultured fetal rabbit gonads is influenced by the mesonephros (54,55) (see section entitled “Differentiation of the Testis”). The mesonephros seems to be crucial not only for gonadal development, but also for the formation of the fetal adrenal cortex (56).

Other Cell Types The future gonadal area consists of a loose mesenchymal tissue covered by the coelomic epithelium and supported by the developing mesonephric tissue. Mesenchymal cells can be recognized throughout gonadal development of both sexes. At very early stages, the de¬ veloping gonad is invaded by capillaries (57). A limited

268

/ Chapter 8

FIG. 3. Ovary-mesonephric complex of a 12j-day-old mouse fetus. The ovary appears as a rather dense homogeneous cell mass, (m) Mullerian duct; (me) mesonephric tubules; (w) Wolffian duct, x 400.

FIG. 4. Testis-mesonephric complex of a 12^-day-old mouse fetus. Beneath the surface epithelium a tunica albuginea (ar¬ rowheads) is developing. Testicular cords have begun to form in the cranial part of the testis, (m) Mullerian duct; (w) Wolffian duct. x425.

FIG. 5. Drawing showing that cell streams con¬ nect the parietal layer of Bowman’s capsule with the somatic cells of the developing gonad. This drawing, made by F. M. Balfour in 1878 (see ref. 40), is probably one of the first illustrations in¬ dicating that the mesonephros contributes cells to the gonad, (ov) Ovary; (ge) germinal epithe¬ lium; (t) tubuliferous tissue, derived from Mal¬ pighian bodies; (mg) Malpighian body.

Embryology of Mammalian Gonads and Ducts /

269

cylindrical appearance (58,59). Primordial germ cells, prob¬ ably in the process of migration, are often contained in the epithelium (60,61). During gonadal sex differentiation, the coelomic epithe¬ lium develops differently in the two sexes. In the male the epithelium is soon delineated by an intact basal lamina, whereby the developing testicular cords become separated from the epithelium. Simultaneously, the epithelium and the outermost testicular cords become separated by a mes¬ enchymal tissue, the developing tunica albuginea. The ep¬ ithelium initially becomes cuboidal; then later when the testis rapidly increases its volume, it becomes flattened. In some cases, germ cells recognized by their alkaline phos¬ phatase activity are trapped within the epithelium (62) (Fig. 8). In the female, the basal lamina of the coelomic epithe¬ lium becomes completed much later in development. As a consequence the epithelium remains, at some locations, in contact with the underlying germ cells (Fig. 9). Germ cells may even be seen within the epithelium a long time after gonadal sex differentiation begins. The morphology of the coelomic epithelium of the ovarian surface varies consid¬ erably with age. A conspicuous cell type with some resemblance to un¬ differentiated blood cells, particularly lymphocytes, has been found in differentiating gonads of both sexes from different species (A. G. Byskov and P. E. H^yer, unpublished ob¬ servations: human, rabbit, mouse). The cells are rounded and contain many ribosomes, sparse endoplasmic reticulum, and a relatively small, often spherical, nucleus with dense peripheral chromatin. They occur one by one or form ag¬ gregates situated close to the germ cells (Fig. 10). In the mouse, such cells disappear after the first week post-birth. The function of these cells remains to be determined.

GONADAL SEX DETERMINATION

FIG. 6. Part of ovary and mesonephros of a 42-day-old pig fetus. Dense cell masses (arrowheads) of mesonephric or¬ igin connect the germ cell cords of the ovary with the parietal layers of Bowman’s capsules, x 200.

amount of information is available on early innervation of the gonads. However, it is likely that an ingrowth of nerves follows the invasion of blood vessels. The epithelium that covers the gonad has previously been termed the germinal epithelium. This term is very unfor¬ tunate because the germ cells do not derive from the epi¬ thelium but rather pass through it (see subsection entitled “Primordial Germ Cells”). Before gonadal sex differentia¬ tion takes place, no complete separation between the epi¬ thelium proper and the underlying tissue exists, since the basal lamina is not yet intact. The epithelial layer consists of pleomorphic proliferating cells, which at places have a

Gonadal sex in mammals is normally determined by the genetic sex (63). From experiments with castrated rabbit fetuses, Jost (64,65) concluded that the genital structures basically are “programmed” for femaleness and that devel¬ opment in the male direction opposes the female program. Therefore, the gonadal primordium is considered to develop into an ovary unless a male organizer counteracts this trend and imposes testicular differentiation (66). In mammals the presence of a Y chromosome is normally associated with development of a testis. Individuals with sex chromosome constitutions XY, XXY, or even XXXXY develop as phenotypical males, and those with XO, XX, or XXX develop as females (67,68). It thus seems that the Y chromosome carries testis-determining sequences (Tds). This gene (or genes) may act by controlling the production of a substance that induces testicular differentiation. Neither tes¬ tis-determining genes nor substances that induce testicular differentiation have yet been identified. Testis determination in mammals is effected through the

270

/ Chapter 8

FIG. 7. Cranially placed mesonephric tubule (me) and mesonephric-derived cell mass (me) of a 9week-old female human fetus. An opening of the tubule is shown by an arrow, x 1,820.

Y chromosome (69), and the gene sequences in action were visualized in chromosome studies of sex-reversed (Sxr) mice (70), in which XX-Sxr develop as phenotypically normal males (71). The X chromosome of these males contains a body that, during meiosis (72), has been translocated from the Y chromosome (73). Some years ago it was proposed that a testis-determining substance might be identical to the male-specific H-Y an¬ tigen (74), also known as the classical histocompatible Y antigen (75). This hypothesis is attractive because, until recently, it was found that almost all mammals possessing a testis were H-Y positive independent of their karyotype (76). The detection of H-Y antigen in these studies is based on serological tests in which it is assumed that antisera raised against male cells will recognize H-Y antigen (serologically detectable male, SDM) (77). However, recently McLaren (78) discovered mice that develop testes but lack H-Y an¬ tigenicity, as determined by tests using T-lymphocyte-mediated histocompatibility response as the originally defined H-Y antigen. These specific male mice are variants of Sxr

XX mice, which are phenotypical males with testes (70) Previously, phenotypical males with testes, although sterile, have been found to be SDM antigen negative [man (79), mouse (80)]. Although theoretical models may explain why H-Y negative males develop testes (81), it still seems un¬ likely that the transplantation H-Y antigen is responsible for testicular formation. Since the transplantation H-Y antigen may not be the same as SDM antigen (77,82), it is possible that SDM antigen rather than H-Y antigen might interfere with testicular differentiation. Results of other experiments lack support with regard to the concept that SDM antigen is the testis inducer, having its primary effect by aggregating fetal male germ cells and Sertoli cells into testicular cords. Epididymal fluid has been reported to be rich in SDM antigen and to exert a reaggregational effect on dispersed rat ovarian cells (83). However, spent culture media from bull and human epididymis (84) or from human testes (85) have the opposite effect on fetal mouse testes, in vitro, by preventing testicular cord for¬ mation and inducing meiosis. Also, the addition of serum

Embryology of Mammalian Gonads and Ducts /

271

FIG. 8. Alkaline phosphatase activity in a cryosection of a 12-week-old human fetal testis. Germ cells of testicular cords and of the surface epithelium (ar¬ rowheads) as well as plasmalemma of Leydig cells (arrows) exhibit activity. x311.

to cultures of fetal rat testes prevents testicular cord for¬ mation (86). As indicated above, gonadal sex determination is depen¬ dent on the sex chromosome constitution. Chromosomal errors, particularly those affecting the X and Y, which can arise in germ cells during meiosis or in the early mitotic divisions during embryogenesis, may interfere with gonadal differentiation (for review, see ref. 67). However, several chromosomal errors, e.g., XO (Turner’s syndrome), XXY (Klinefelter’s syndrome), and Tfm (testicular feminization), do not seem to affect the primary sex determination of the gonad, but rather later stages of gonadal differentiation. An individual in which the gonad develops with both ovarian and testicular tissues is a hermaphrodite, or sex mosaic, and has been described in many species (78), in¬ cluding humans (87). They could develop from fused XX and XY embryos [i.e., chimeras, which have also been made experimentally (88,89)] or as a result of Y-chromosome nondisjunction (90). The gonads of chimeras do not differ¬ entiate into an ovary, an ovotestis, or a testis according to the relative proportion of male and female clones (91,92).

A male phenotype develops in more than 40% of the cases if the XY cells comprise more than one-third of the cells in the gonads (93). A dominant influence of the male tissue over the female tissue is also demonstrated by the inhibition of growth of the Mullerian duct in fetal mouse hermaph¬ rodites, in which only 15% of testicular tissue caused normal male inhibition of Mullerian duct growth (94). The gonadal sex of mammals is highly stable, and only in the primitive young marsupials can it be changed exper¬ imentally. In the newborn male opossum (95,96) and wal¬ labies (97), the testicular anlage can develop into an ovo¬ testis or an ovary by giving low doses of estradiol propionate. Turner and Asakawa (98) claimed that when embryonic mouse ovaries and testes were grafted closely together under the kidney capsule, the ovary developed testicular tubules. It was proposed that the testis secreted substances that in¬ duced differentiation of testicular structures in the ovarian anlage. However, these results could not be reproduced by Ozdzenski et al. (99) or by Burgoyne et al. (100), who suggest instead that the ovarian “testicular cords” are struc¬ tures of mesonephric origin. Neither in co-cultures between

272

/ Chapter 8

FIG. 9. Part of ovarian cortex of a 9-week-old human fetus showing connections between the down-grow¬ ing surface epithelium and an oogonium (arrow¬ head). x 1,820.

embryonic mouse ovaries and testis (101) nor between fetal and neonatal rabbit testes and ovaries (102) did any testicular structures develop in the ovaries.

DIFFERENTIATION OF THE OVARY A functional ovary depends on three major events taking place during early stages of gonadogenesis: the initiation of meiosis, the formation of follicles, and the differentiation of steroid-producing cells.

Different Patterns of Early Ovarian Differentiation Mesonephric-derived cells populate the gonad a long time before morphological sex differentiation takes place. The ovarian-mesonephric connection is retained during ovarian differentiation, although the mesonephric tissue gradually regresses (Fig. 11). In most species the central part of the

differentiating ovary becomes occupied by the invading me¬ sonephric cells (i.e., the intraovarian rete), which push the germ cells toward the periphery. An ovarian cortex richly populated with germ cells and a medulla consisting mainly of mesonephric cells are thereby formed. Two major patterns of ovarian differentiation can be rec¬ ognized depending on whether the germ cells of the ovary undergo “immediate” meiosis, without previous steroid pro¬ duction, or “delayed” meiosis (Fig. 12), with steroid se¬ creted before meiosis begins (102). In species with “im¬ mediate” meiosis (e.g., mouse, rat, hamster), the germ cells of the ovary enter the first meiotic prophase simultaneously with or shortly after gonadal sex can be recognized mor¬ phologically. These ovaries produce little or no steroids de novo until follicles are formed. In species with “delayed” meiosis (e.g., pig, sheep, dog, cow), the beginning of meiosis in the female is delayed up to 45 days (cow) with respect to testicular differentiation, i.e., delay period. In contrast to species with immediate meiosis, such ovaries produce

Embryology of Mammalian Gonads and Ducts /

273

FIG. 10. Aggregate of lymphocyte-like cells in a new¬ born mouse ovary. At some places, such cells are in contact with oocytes (arrowhead), x 4,560.

various amounts of steroids during the delay period (see subsection entitled “Ovary”). In species with immediate meiosis, the ovary appears compact when meiosis begins at an early stage of sex dif¬ ferentiation. The germ cells are distributed uniformly or in clusters throughout the entire ovarian tissue (e.g., mouse, hamster) or in a basically well-defined cortical area (e.g., human). In ovaries of species with delayed meiosis, the germ cells become enclosed in germ cell cords during the delay period. In some species [pig (103); cat, mink, ferret (104); sheep (43); cow (105)] the germ cell cords are lined with a basal lamina and clearly defined from the surrounding loose mes¬ enchyme (Fig. 13). The cords are irregularly shaped and are tightly packed with somatic cells and germ cells. By the end of the delay period the cell cords begin to break up in the central part of the ovary close to the intraovarian me¬ sonephric cell cords. This process is related to the beginning of meiosis (see subsection entitled “Meiotic Prophase”).

In other species (e.g., the rabbit) the germ cell cords are more closely packed in the ovarian cortex and are only clearly recognizable in the inner part, where they connect with the intraovarian mesonephric cell mass. The development of the human fetal ovary represents a transitory example between immediate and delayed meiosis. Although there is a delay period of 2 to 3 weeks, no or very little steroids are produced de novo (see subsection entitled “Ovary”). Intraovarian mesonephric cell cords occupy the medulla before meiosis starts, and the cortically placed germ cells are not confined to cords, but are rather gathered into large clusters (Fig. 14).

Meiotic Prophase The two meiotic divisions are unique for germ cells. During the first meiotic division, maternal and paternal genes are exchanged before the pairs of chromosomes are divided

274

/ Chapter 8

Mullerian duct Gonadal

Mesonephros

FIG. 11. Transformation of the genital duct system during the period when the gonads pass from the undifferentiated state to recognizable testes and ovaries. [From A. G. Byskov, Chap¬

into the two daughter cells, each containing In chromo¬ somes and 2c DNA. The second meiotic division occurs without being preceded by DNA synthesis. This division results in formation of the haploid germ cells with a In set of chromosomes and lc DNA (for review see ref. 106). Although meiosis represents similar events in germ cells of the two sexes, the time schedule and the resulting number of haploid germ cells differ greatly between ovary and testis. In the female germ cells, meiosis is initiated at early stages of development, often during fetal life. The first germ cells to begin meiosis are always localized at the inner part of the cortex. However, meiosis is arrested in late prophase of the first meiotic phase, and the divisions are delayed and do not take place until much later in the mature animal (around ovulation time). In the male germ cells, meiosis begins at puberty and proceeds without delay (Fig. 15). Meiosis in each female germ cell results in a single egg and two, or eventually three, abortive cells, the polar bodies, whereas four sperms are produced by each male germ cell (Fig. 15).

ter 1, in: Germ Cells and Fertilization, edited by C. R. Austin and R. V. Short, Cambridge University Press, Cambridge (1982).]

Proliferation and Premeiotic DNA Synthesis of Oogonia The oogonia continue to divide mitotically until they enter meiosis. Fluctuations in the total number of germ cells in fetal and neonatal ovaries of different mammalian species are seen in Fig. 16 (107). The rate of mitosis during the time preceding meiosis varies between species. In species with delayed meiosis, the mitotic activity is low during most of the delay period, but it increases rapidly shortly before meiosis starts [pig (108)]. In species in which meiosis starts shortly after gonadal sex differentiation, the period with low mitotic activity in the oogonial population is nonexistent (mouse) or very short (rat), and meiosis is introduced by a series of mitotic divisions shortly after gonadal sex differ¬ entiation (109). Often, groups of germ cells divide syn¬ chronously. When meiosis begins, similar groups exhibit synchrony while passing through transitory stages of meiosis (110,111). Germ cells of such groups are often connected by intercellular bridges (112). It has been proposed that a single-stem cell gives rise to such germ cell groups

Embryology of Mammalian Gonads and Ducts / 275 WITSCHI

BURNS

FIG. 12. Schematic drawings of models of ovarian differentia¬ tion from Witschi (39,40), Burns (94), and Byskov (197). Witschi and Burns suggest that somatic cells of ovary are mainly derived from surface epithelium, whereas Byskov proposes mesonephric origin. Model by Witschi includes degeneration of medulla and proliferation of cortex. Burns believes that sec¬ ondary proliferation of surface epithelium forms secondary sex cords, which contribute the bulk of ovarian somatic cells. Bys-

BYSKOV

kov’s model involves two types of transitory stages of ovarian differentiation, one in which meiosis starts almost simultane¬ ously with gonadal sex differentiation (immediate meiosis) and another in which meiosis is more or less delayed with respect to sex differentiation (delayed meiosis). In both cases, me¬ sonephric-derived cells are main contributors to ovarian cell mass. [Reprinted from A. G. Byskov, Differentiation of mam¬ malian embryonic gonad, Physiol. Rev., 66:71-117 (1986).]

FIG. 13. Germ cell cords of a 42-day-old pig ovary. The cords are tightly packed with somatic cells and germ cells (arrowheads), x 1,000.

FIG. 14. Part of ovary from a 21-week-old human fetus. Small follicles (arrowheads) are present in the inner part of the cortex. The middle part contains oocytes in different stages of meiosis, whereas the peripheral layer still contains oogonia. x 360.

MALE

FEMALE

Mitosis

Mitosis

Leptotene

Zygotene

Pachytene

Diplotene resting stage (4n DNA) Mitosis

^

Resting stage (2n DNA)

Leptotene

Zygotene

Adult life

Vjg>

Adult life

V

1st meiotic division

Pachytene 1st meiotic division

2nd meiotic^ division '

^ 2nd meiotic division ^ ^

^

(§) @ @ @

FIG. 15. Life cycles of male and female germ cells. Germ cells of both sexes divide mitotically until, or shortly after, gonadal sex differentiation. The female germ cells all enter meiosis at early stages of development, whereas the male germ cells keep a resting stem cell population which can divide mitotically and from which meiotic cells continue to emerge throughout life. The male germ cells rest with 2c DNA (2c = 2n), whereas the female germ cells rest in the diplotene stage with 4c DNA (4c = 4n). [From A. G. Byskov, Chapter 1, in: Germ Cells and Fertilization, ed¬ ited by C. R. Austin and R. V. Short, Cambridge University Press, Cam¬ bridge (1982).]

Embryology of Mammalian Gonads and Ducts / 277 300

200

x 50

S.

5) c

FIG. 4. Reductive pathways of progestin and androgen me¬ tabolism in the ovary. Notes: 1. The 20a-hydroxysteroid de¬ hydrogenase occurs in ovaries of most species (shown), while in bovines a different enzyme has 203 stereospecificity (not shown). 2. 5a-Reduction occurs similarly for C2r and C19-steroids, but it is not known whether it is the same 4-ene-5areductase enzyme acting in each instance. 53-Reduction, an important reaction in the peripheral metabolism of C2r and

C19-steroids, is an additional pathway present in the hamster ovary. 3. Conversion of 5a-reduced steroids via the 3a-hydroxysteroid dehydrogenase (shown) is well documented. An alternative pathway involving a 33-hydroxysteroid dehydro¬ genase (not shown) has been demonstrated under limited con¬ ditions. The expected 33-hydroxy products are similar to those shown. The 33-epimer of androsterone is epiandrosterone.

Testosterone

Scr-dihydrotestosterone

5 cr-androstane -3a,17|3-diol

340 / Chapter 10 steroidogenic process in the follicle. We will not consider the molecular mechanisms of gonadotropin action, which are beyond the scope of this chapter, nor will we review the extensive literature that has developed to support the concept that gonadotropins—follicle-stimulating hormone (FSH) and luteinizing hormone (LH)—regulate steroido¬ genesis and other aspects of follicular maturation by stim¬ ulating the production of cyclic AMP (cAMP), the so-called second messenger. These topics have been previously re¬ viewed (51,52). Suffice it to say, cAMP is now well es¬ tablished as the principal, if not the only intracellular me¬ diator of FSH and LH/human chorionic gonadotropin (hCG) actions on the follicular cell types. FSH and LH satisfy virtually all of the criteria orginally set out by Sutherland and colleagues (53) and subsequently extended by Kuo and Greengard (54) to indicate that a hormone mediates its ac¬ tions via cAMP and cAMP-dependent phosphorylation of proteins. Calcium has a permissive role in the steroidogenic process, as demonstrated for FSH-stimulated steroidogen¬ esis in rat granulosa cells (55). Steroidogenic Cells and Their Origins In all mammalian species the principal cell types involved in follicular steroidogenesis are of two basic types: (a) LHresponsive secretory cells, comprising the theca interna cells of the follicular envelope and the interstitial cells of the ovarian stroma, and (b) FSH-responsive cells, consisting exclusively of granulosa cells, which only later in follicular maturation also acquire the ability to respond to LH. These two basic cell types fulfill distinct roles in the steroidogenic process by virtue of their different regulatory hormones and their dissimilar expression of steroidogenic enzymes. Granulosa and theca/interstitial cells have distinct embryological origins. Granulosa cells appear to be derived mainly, although perhaps not exclusively, from certain cells within the intraovarian rete ovarii which closely resemble granulosa cells in terms of their organelles (56) and micro¬ filaments (57-60). The intraovarian rete ovarii in the prefollicular ovary consists of cell cords and tubules and of mesenchymal cells in the ovarian medulla. Early follicular development occurs centrally in the innermost part of the cortex within the rete cords as rete cells move between and attach to oocytes, differentiate into granulosa cells, and then organize follicles (56,61). Once follicles form and the granulosa cells become fully enclosed by the follicular basement membrane, it is clear that these cells alone proliferate to produce the membrana granulosa layer. However, this population of granulosa cells does not remain uniform, but as the antral follicle develops, these cells become organized into morphologically distin¬ guishable regions with specialized functions. The granulosa cells in the layer immediately adjacent to the oocyte (i.e., corona radiata cells) establish intimate contact with the oo¬ cyte up until the preovulatory stage by means of cellular processes traversing the zona pellucida and forming gap junctional complexes with the oolema. These cells serve as

“nurse” cells, providing nutrients for oocyte growth and also presumably exchanging regulatory factors with the oocyte, thereby relaying signals required for the coordinated mat¬ uration of the follicle and the oocyte. The cumulus granulosa cells comprise the cellular mass that surrounds the oocyte (i.e., the cumulus oophorus) and attaches the oocyte to the follicle wall. The cumulus cells physically support the oo¬ cyte within the follicle and may contribute to its nutritional and regulatory needs. Following ovulation the oocytecumulus complex facilitates pick-up by the oviduct and may contribute to the final maturation of the oocyte, as well as the capacitation of spermatozoa that must penetrate through the extracellular material formed by the cumulus cells. The majority of granulosa cells form the so-called mural or pari¬ etal granulosa cells lining the follicular cavity. Those mural cells adjacent to the follicular basement membrane are the first to differentiate steroidogenic responsiveness to LH based on the acquisition of LH binding sites (62,63), the expres¬ sion of A5-3(3-hydroxysteroid dehydrogenase activity (64), and the level of cytochrome P-450 (65). In addition, mural granulosa cells at this stage lose the differentiation antigen that is uniformly present on granulosa cells of earlier fol¬ licular stages (66). Other in vitro evidence suggests that subpopulations of granulosa cells may exist with respect to differential sensitivity to FSH and vasoactive intestinal pep¬ tide (VIP) (67). The theca cells appear to differentiate from mesenchymal cells in the ovarian stroma (68,69). Since the theca layer is not present in primary follicles but differentiates as follicles grow and mature, it is evident that theca cells arise contin¬ ually throughout reproductive life; the mesenchymal pro¬ genitor cells are perhaps pluripotent stem cells that also contribute to cells of loose connective tissue. It has been suggested that in the immature mouse the theca layer might also be formed from certain intraovarian rete cells that are initially contiguous with cells forming the follicular gran¬ ulosa cells but become separated from association with the oocyte as the follicle is enclosed (70,71). There is similar evidence in the rabbit (72). However, these cells in the mouse appear to give rise principally to the primary inter¬ stitial cells (70). Primary interstitial cells in fetal ovaries of the human appear to have distinct steroidogenic activities (see section on the prefollicular ovary, below). Secondary interstitial cells found in the adult ovary are derived from theca cells of atretic follicles (73-77). Whereas the oocyte and surrounding granulosa cells of an atretic follicle degenerate and are eliminated from the ovary, the theca cells in the follicular envelope survive as small islands of steroidogenic cells in the ovarian stroma. On this basis, certain similarities in function of theca and secondary in¬ terstitial cells of adult ovaries might reasonably be expected. However, theca and interstitial cells probably do not have identical biosynthetic properties since the interstitial cells, in contrast with the theca cells, are less likely to be influ¬ enced by paracrine regulatory substances secreted by the membrana granulosa; instead, they may receive direct sym¬ pathetic innervation. Secondary interstitial cells also differ

Follicular Steroidogenesis and Its Control / from theca cells in that their androgen biosynthetic activity not only persists throughout reproductive life (78-80) but continues in aged ovaries (81,82). In later sections of this chapter we consider the evidence for steroidogenic functions of interstitial cells, theca cells, and granulosa cells in the early stages of ovarian organo¬ genesis and at various stages of follicular development in the adult ovary. However, the characteristic steroidogenic functions of theca and secondary interstitial and granulosa cells in adult ovaries and mature follicles, typical of the follicular phase, are now described.

Pathways and Their Control by Gonadotropins (FSH and LH) Theca and Interstitial Cells The most abundant steroid products of mature theca cells of all species are C 19-compounds, including the 4-ene-3f3hydroxysteroids and 5a-reduced androgens, which are pro¬ duced from the catabolism of cholesterol by pathways pre¬ viously described. As discussed earlier, since the theca in¬ terna becomes highly vascularized as the follicle matures, it is reasonable that the internalization of blood-borne li¬ poprotein provides the major source of cholesterol for ste¬ roidogenesis by theca cells in vivo. Steroidogenesis in the absence of lipoprotein cholesterol in vitro is limited (83). The preferred enzymatic route for the conversion of cho¬ lesterol to androgens in the theca of human (84) and bovine (85) ovary is via the 5-ene-3(3-hydroxysteroid pathway. DHEA produced via this pathway is then metabolized to androstenedione. It is apparent that rat ovarian interstitial cell cultures (i.e., consisting of interstitial cells and follicular cell types) produce both 5-ene-3(3-hydroxy and 4-en-3-oxo intermediates (86). LH action via specific receptors present on the theca cells at all follicular stages (87-93), and consequent production of cAMP (81,94-96), provides the principal stimulus for these steroidogenic activities. Studies with cultured ovarian interstitial cells isolated from hypophysectomized rats in¬ dicate that the cells constitutively express A5-3(3-hydroxysteroid dehydrogenase and have functional LH receptors but are not steroidogenically active unless induced to dif¬ ferentiate with either LH or prostaglandin E2 (PGE2) (97). These results with ovarian interstitial cell cultures might reflect similar control mechanisms occurring in thecal tissue, which also responds to LH or PGE2 with increased androgen secretion (96,98,99). The steroidogenic action of LH on theca cells apparently increases the activities of 17a-hydroxylase:C-17,20-lyase in ovaries or follicles of rat (100,101,102) and hamster (103). These enzyme activities are rate-limiting and appear to be the site at which LH stimulates C19-steroid production by theca cells, as follicles progress from small antral stages to early preovulatory follicles in the rat (102), and where ovarian androgen production is substantially restricted in late preovulatory follicles (104) (also see the section on

341

preovulatory follicles, below). Recent studies have em¬ ployed immunoblot analysis to measure the specific contents (i.e., amount of enzyme protein per microgram of tissue protein) of this and other steroidogenic enzymes and their electron donors in follicles dissected from bovine ovaries of mature animals (105). These studies have demonstrated that 17a-hydroxylase P-450 in follicles increased fivefold between medium-sized (9-11 mm) and large (14-18 mm) follicles, indicating that an increase in enzyme protein oc¬ curs in the follicular cells (granulosa and theca) as follicles mature. Significantly, this enzyme protein was undetectable in bovine corpora lutea throughout the luteal phase, con¬ sistent with the loss of the enzyme activity as follicles lu¬ teinize in response to the LH surge. (This aspect is discussed later with respect to late preovulatory follicles after the LH surge.) These changes showed specificity for the Ha-hy¬ droxylase P-450 since its electron donor, NADPH-cytochrome P-450 reductase, has a similar specific content dur¬ ing follicle development and in corpora lutea. Although progesterone is apparently not limiting as an intermediate for thecal androgen production in small antral follicles, progesterone accumulation in isolated small antral and preovulatory rat follicles is stimulated in vitro by hCG, indicating that activation of the LH receptor also stimulates a step in the conversion of cholesterol to progesterone (102). Further studies tended to rule out the possibility that this action was due to decreased progesterone metabolism. Iso¬ lated theca cells from small antral and preovulatory rat fol¬ licles also produced increased progesterone when stimulated with cAMP, this stimulation being much greater with theca from preovulatory follicles (102). Therefore, it is evident from these studies that progesterone production by theca cells is also hormonally regulated and varies with the stage of follicular development. This effect of LH is probably the result of increased activity of C27 side-chain cleavage and is consistent with increased pregnenolone production in LHstimulated rat ovarian interstitial cell cultures (86). Ultrastructural immunocytochemical visualization of P-450Scc in ovaries of immature rats indicates that this mitochondrial enzyme is initially found in only a few theca cells, but the number of theca cells containing this enzyme increases after pregnant mare serum gonadotropin (PMSG) treatment (30). Significantly, these studies showed a strong reaction for P450SCc in interstitial cells even before PMSG treatment, with no apparent change as a result of hormonal stimulation. Androstenedione is the principal aromatizable C19-steroid produced by isolated theca interna tissue or cells, with lesser amounts of testosterone occurring, as described in the rat (94), hamster (103,106), pig (107), sheep (194), cow (85,108,109), and human (110,111). The greater abundance of androstenedione than testosterone is due to a deficiency in 17(3-hydroxysteroid dehydrogenase, a fact that has signifi¬ cance in the steroidogenic cooperation of theca and granulosa cells, as discussed later. Rat ovarian interstitial preparations in culture are also active in 5a-reduction of Ci9-steroids; the major LH-stimulated product is androsterone (86), which may be produced by sequential action of 5a-reductase and 3a-hy-

342

/

Chapter 10

droxysteroid dehydrogenase (Fig. 4). The next most abun¬ dant 5a-reduced metabolite is 5a-androstane-3a,17(3-diol, which can be derived from testosterone according to the same reactions described above for androstenedione or, alterna¬ tively, might result from conversion of androsterone to 5aandrostane-3a,17(3-diol in a reversible reaction catalyzed by the apparently weak thecal activity of 17(3-hydroxysteroid dehydrogenase. It has been demonstrated that 3a-hydroxy-5a-pregnan-20-one is converted to 5a-androstane3a,17(3-diol by supernatants of ovarian homogenates from immature rats, suggesting that this pregnane may be an earlier intermediate in a pathway to the androstanediol (112). Activity of 5a-reductase in thecal tissue from hamster preo¬ vulatory follicles may not be regulated by LH (103) but is subject to LH control in thecal/interstitial cells in prepubertal rat ovaries (100,101). There is no change in thecal 5areductase activity in the adult rat during pregnancy as fol¬ licles develop from small antral to preovulatory stages (113). Activity of ovarian 5a-reductase, which is greatest before the onset of puberty (14,40,45), is greatly decreased in ovaries of adult rats (114-116) and in immature rats in which puberty has been advanced with PMSG (42). The physio¬ logical significance of 5a-reduced androgens in the im¬ mature rat is indicated by the effect of exogenous 5a-androstane-3(3,17(3-diol, but not its 3a-epimer, to advance the onset of puberty (117) and by there being sufficient con¬ centrations of the 3(3-epimer in blood to account for these changes at puberty (37). FSH has also been shown to induce an epimerization reaction in the immature rat ovary that converts 5a-androstane-3a,17|3-diol to the biologically ac¬ tive 3(3-epimer (118). Alternate routes of progesterone catabolism, in addition to those involved in the production of androgens, also occur in theca cells. Transiently increased production of 17ahydroxyprogesterone in the human at midcycle (119,120) may primarily arise in the theca cells before similar path¬ ways are fully active in the granulosa cells. The corpus luteum becomes the principal source of this steroid in the luteal phase of the human (121). In rat interstitial cells, the principal C2] metabolite is 20a-dihydroprogesterone (86). The production of this metabolite is increased by LH treat¬ ment, but the amount produced is still less than that of intermediates and products of the 4-en-3-oxosteroid path¬ way. Theca cells may also aromatize androgens and contribute directly to ovarian estrogen secretion in varying degrees depending on species and follicular stage, e.g., in human (122-124), monkey (125), mare (126,127), cow (85), ham¬ ster (106), sheep (128), and pig (107). However, results of most studies indicate that aromatase activity (i.e., conver¬ sion of Ci9-steroids to estrogens) in the theca cells either is not significantly greater than that in the granulosa cells (e.g., as in pig and sheep) or is considerably less than in the granulosa cells. In studies with cultures of follicular tissue from immature pigs, theca initially has considerably less aromatase activity than granulosa cells expressed on a per follicle basis, but when follicular growth and maturation

was induced with PMSG treatment, aromatase activity in theca cells increased substantially while that in granulosa cells declined slightly (107). Studies with sheep follicle cell types also indicate substantially increased estrogen produc¬ tion by theca cells as follicles mature (128). Therefore, in mature follicles of pig and sheep, both theca and granulosa cells appear to have similar abilities in vitro to produce estrogens, either from endogenous androgen (theca cells) or supplied substrate (granulosa cells). Most studies with human theca cells in culture agree that the quantity of estrogen secretion is significant but small (129), regardless of whether the theca is obtained from small or large follicles or at various stages of the menstrual cycle (110). For the preovulatory human follicle it was estimated that 99.9% of the aromatase activity resides in the membrana granulosa (130,131), and there is an excellent positive cor¬ relation between granulosa aromatase activity and the con¬ centration of estradiol in follicular fluid (130). These results strongly indicate that thecal cell production of estrogen in the human is relatively minor in comparison to the aro¬ matizing capabilities of granulosa cells from large antral follicles. Results of other studies with human follicle cells in culture (111,124) are difficult to interpret in terms of relative theca and granulosa cell aromatase activities since data are often expressed as estradiol production per culture without corrections for cell number. There is little direct information to distinguish the ste¬ roidogenic functions of secondary interstitial cells from those of the theca interna cells, and the biosynthetic pathways are generally assumed to be similar. However, as already noted, the immunocytochemical staining for P-450scc in interstitial cells of immature rat ovary is greater than in theca cells; also, it does not depend on exogenous gonadotropin treat¬ ment (30). Whether this difference reflects similar differ¬ ences in enzymic activity is not known. In vitro studies using dispersed ovarian preparations from hypophysectomized rats (designated as ovarian interstitial cells but containing all ovarian cell types) have investi¬ gated the regulation of the LH-responsive cell types (86,97). The biosynthetic activities of these mixed cell cultures are similar to those already described for theca cells. Specific isolation of secondary interstitial cells from rat ovarian stroma has not been feasible without significant contamination by theca and follicle cell types but is more readily accomplished in humans and in larger animals. Nevertheless, true inter¬ stitial cells (i.e., glandular cells) in the ovarian stroma of the human constitute only about 1% of the ovarian volume during the menstrual cycle, with this value increasing to 4— 6% in late pregnancy (132). Human stromal cells obtained on cycle day 18 were incubated with [3H]pregnenolone and [14C]progesterone and were shown to use both 4-en-3-oxo and 5-ene-3^-hydroxy pathways, with the products being androstenedione (the major product), testosterone, proges¬ terone, 17a-hydroxyprogesterone (133), and, under some circumstances, estrone and estradiol (134). Production of these steroids was increased by treatment with hCG. In other studies, cultures of human stromal cells synthesized,

Follicular Steroidogenesis and Its Control / androstenedione, 17a-hydroxyprogesterone, progesterone, and estradiol but in lesser amounts than by theca cells (123,135).

Granulosa Cells Whereas the steroidogenic pathways in the theca and in¬ terstitial cells function primarily in the de novo production of androgens, the pathways in the granulosa cells are or¬ ganized principally for the metabolism of C19-steroids (i.e., androgens) to estrogens and for the de novo synthesis of progesterone and its C2i metabolites. As will be discussed in following sections, cooperation of the theca and granulosa cell compartments appears to be crucial to the control by gonadotropins of follicular steroid hormone secretion in all species studied. This cooperation takes the form of exchange of steroid pathway intermediates, direct steroidal effects on enzyme activities, and paracrine regulation. Evidence from both in vivo and in vitro studies indicates that the granulosa cells of large antral and preovulatory follicles are the principal, although not exclusive, site in all species of ovarian aromatase activity and estrogen biosyn¬ thesis. Regulation of androgen aromatization in the gran¬ ulosa cells of all species studied appears to be by the action of FSH (136), which in rat granulosa cell cultures stimulates aromatase enzyme activity as measured in a cell-free assay (137) and requires RNA and protein synthesis for expression of this action (138). Recent studies with rat granulosa cells suggest that aromatase cytochrome P-450, as detected in radiolabeled immunoisolates, is induced by FSH or dibutyryl cAMP (139). In addition, FSH or dibutyryl cAMP has been reported to stimulate two to three times the amount of the NADPH-cytochrome P-450 reductase detected in cul¬ tured rat granulosa cells (in immunoisolates and by immunoblot analysis), but this component is apparently in excess of the specific aromatase cytochrome P-450 (140). In addition to this action of FSH to induce aromatase enzyme in granulosa cells, estrogen biosynthesis requires the co¬ operation of the theca cells in supplying the androgen sub¬ strates for the aromatization reaction. This important aspect is discussed in detail in a following section. The production of progesterone and its metabolites (i.e., progestins) is one of the major biosynthetic activities of granulosa cells in large antral and preovulatory follicles. Progesterone biosynthesis occurs in granulosa cells initially in response to FSH stimulation, but this action is later aug¬ mented by LH after its receptors have differentiated. In culture, FSH stimulates progesterone biosynthesis in un¬ differentiated granulosa cells from immature rats (141-145) and in granulosa cells from various other species, including human (124,129,146), simian (147), porcine (24), and avian (148). A similar effect of FSH on progesterone production has also been shown for cumulus granulosa cells of rat (149). In contrast, LH alone does not stimulate progesterone bio¬ synthesis in cultured granulosa cells from hypophysectomized, estrogen-treated rats but is an effective stimulus in

343

vitro following either a 24-hr in vivo treatment with FSH to induce LH receptors (142) or after 2 days of FSH priming in vitro (150). Many studies in various species substantiate the stimulation of progesterone production by LH in mature granulosa cells from antral follicles. In all species studied the greatest stimulation of progesterone biosynthesis in vivo follows the LH surge, often after a transient decrease in production, as granulosa cells undergo differentiation (i.e., luteinization) to form granulosa-lutein cells. The progesterone biosynthetic pathway (see Fig. 2 and the first three sections on biosynthetic pathways, above) in granulosa cells is typical of all steroidogenic cells, involving conversion of cholesterol to pregnenolone and then to pro¬ gesterone. In antral and preovulatory follicular stages, in¬ tracellular cholesterol is probably almost entirely derived from de novo synthesis and perhaps to a small extent from endogenous lipid stores, since cholesterol associated with lipoproteins in blood (28,151) cannot penetrate the avascular granulosa cell layer. Only after ovulation is extracellular lipoprotein likely to be a major source of cholesterol as a precursor for biosynthesis in granulosa-lutein cells (23,152). The apparent rate-limiting reaction for progesterone bio¬ synthesis, regulated by FSH action on granulosa cells, is cholesterol side-chain cleavage (144,153,154). The effects of hormonal stimulation on this enzyme activity have been suggested to involve an increase in the association of cho¬ lesterol with the P-450scc via a low-molecular-weight ac¬ tivator peptide and to increase the supply of intramitochondrial cholesterol by a cytoskeleton-mediated process, as appears to be the situation in the adrenal cortex (32). There is similar evidence in the ovary for stimulatory effects of LH/hCG on cholesterol transport and activation of the cho¬ lesterol side-chain cleavage enzyme (155-157). In rat gran¬ ulosa cells, FSH stimulates pregnenolone production in the presence of cyanoketone, which inhibits metabolism to pro¬ gesterone; 25-hydroxycholesterol, which readily enters the mitochondria, further enhances this effect of FSH, sug¬ gesting that FSH may also increase cholesterol side-chain cleavage activity in this cell type (153). Concomitant in¬ creases in mitochondrial activity of side-chain cleavage en¬ zyme in ovaries of rat (158) and pig (154) further suggest that synthesis of this enzyme may be increased. However, immunoblot analysis of C27 side-chain cleavage cytochrome P-450 and its electron donor, adrenodoxin, in dissected bo¬ vine follicles has not shown increases in specific contents of either of these enzyme proteins as follicles mature from medium to large sizes (105). These studies report that both P-450SCc and adrenodoxin increase only in corpora lutea at the early-mid luteal phase, which suggests that induction at this stage is primarily the result of earlier LH stimulation. Additional actions of LH, in luteal tissue of various spe¬ cies, include enhancement of cholesterol availability by (a) stimulating 3-hydroxy-3-methylglutaryl coenzyme A reduc¬ tase, which is rate-limiting for cholesterol biosynthesis (159); (b) acutely stimulating cholesterol esterase, thereby increas¬ ing the availability of free cholesterol from intracellular stores of fatty acid esters (160-162); and (c) increasing the

344

/

Chapter 10

number of lipoprotein receptors (163), thereby enhancing uptake of cholesterol in the form of lipoprotein. FSH (87), and subsequently LH (164-167), greatly stim¬ ulate rat granulosa cell activity of A5-3(3-hydroxysteroid dehydrogenase:A5‘4-isomerase. It has been suggested that A5 4-isomerase activity may be present in excess in adrenal and testis tissue (168,169), but the issue of whether this enzyme activity and the A5-3 (3-hydroxysteroid dehydrogen¬ ase are separate entities or a single enzyme remains unre¬ solved. Following gonadotropin stimulation, activity of A5-3(3-hydroxysteroid dehydrogenase is apparently not limiting for progesterone synthesis in cultured granulosa cells from immature rats, which do not substantially accumulate pregnenolone during FSH-stimulated syn¬ thesis of progesterone and 20a-dihydroprogesterone (170). This may not be true in the pig, in which cultured granulosa cells accumulate greater quantities of pregnenolone than progesterone (171). The metabolism of progesterone to 20a-dihydroprogesterone in granulosa cells (see section on 20-hydroxysteroid dehydrogenase, above) is influenced by FSH stimulation of the 20a-hydroxysteroid dehydrogenase enzyme, as dem¬ onstrated by in vivo experiments with hypophysectomized, estrogen-treated rats (172) and with cultured rat granulosa cells (173). In contrast, LH is apparently not required for maximal activity of this enzyme in ovaries of proestrous rats (174), and hCG is not an effective stimulus in FSHprimed rat granulosa cells in culture (164). However, var¬ ious studies in the rat have reported inhibitory or stimulatory actions of LH/hCG on 20a-hydroxysteroid dehydrogenase, concomitant with luteotropic or luteolytic actions, respec¬ tively (46,175-177). The conversion of progesterone to 20adihydroprogesterone in the rat corpus luteum (178,179) is important in regulating progestational activity during preg¬ nancy (180) and parturition (181,182). The same pathway to 20a-dihydroprogesterone occurs in mouse (183) and hu¬ man (184); however, in bovine corpus luteum, the principal metabolite is 20(3-dihydroprogesterone (184). Other C2i metabolites of progesterone result from the sequential action of 5a-reductase and 3a-hydroxysteroid dehydrogenase, the products being 5a-pregnane-3,20-dione and 3a-hydroxy-5a-pregnan-20-one, respectively (Fig. 4). The extent of formation of these products depends on the 5a-reductase activity, which is greatest in immature rats (115,185). Similar sequential catabolism of 20a-dihydroprogesterone to 20a-hydroxy-5a-pregnan-3-one and then 5a-pregnane-3a,20a-diol is expected (Fig. 4); the latter metabolite has been found to be a significant product in long-term cultures of rat granulosa cells (186). A fully re¬ versible conversion of 5a-pregnane-3,20-dione to 3(3-hydroxy-5a-pregnan-20-one is also expected to be catalyzed by the 3(3-hydroxysteroid dehydrogenase (see section on 5areduced pathways, above), but little is known about the regulation involved. In contrast with theca cells, granulosa cells of all species studied, with the apparent exception of the bovine (85), do not have significant activities of C2i side-chain cleavage

enzymes and synthesize little or no C19-steroids from either pregnenolone or progesterone (85,94,99,187-192). Granulosa cells possess considerable activity of the 17(3hydroxysteroid dehydrogenase enzyme (103,193-196), which acts on both C19-steroids, androstenedione and testosterone, as well as aromatization products, estrone and estradiol (Fig. 2). Although androstenedione is the major ovarian androgen in most species, the 17(3-hydroxysteroid dehydrogenase re¬ action favors the production of estradiol as the major ovarian estrogen. The presence of 5a-reductase and 3a-hydroxysteroid dehydrogenase in granulosa cells also permits thecaderived androgens to be converted to several ring A-reduced metabolites (Fig. 4) according to the same reactions occur¬ ring for progesterone metabolism described above. FSH does not influence the production of 5a-reduced metabolites of testosterone (i.e., androsterone and 5a-androstane-3a, 17(3diol) in rat granulosa cell cultures (141). Furthermore, FSH was ineffective in maintaining elevated 5a-reductase activ¬ ity in prepubertal rats after hypophysectomy (197). From a physiological viewpoint, aromatization of thecaderived androgens is perhaps the most important pathway of androgen metabolism in the granulosa cell. This key concept of cooperation of the follicle cell types in the reg¬ ulation of estrogen biosynthesis is considered next. Other important actions of theca-derived androgens on estrogen biosynthesis via paracrine mechanisms are considered later (see section on estrogens, below). Cooperation of Theca and Granulosa Cells The functional basis of the modem concept of cellular cooperation in follicular estrogen biosynthesis has already been described: Theca cells are stimulated by LH to produce aromatizable androgens, and granulosa cells respond to FSH with increased aromatase activity but without de novo pro¬ duction of C19-steroid substrates. We will briefly discuss the historical development of the current “two-cell-type, two-gonadotropin” theory of steroidogenic regulation and then consider the direct in vitro experimental evidence in animals and humans that supports this concept and other aspects of cooperation in follicular steroid metabolism. The early literature implicated the follicle in estrogen biosynthesis (198), with some authors favoring the theca interna cells as the principal site of synthesis (199-201) and others suggesting the follicular epithelium (i.e., membrana granulosa) as the primary source (202). The first substantial evidence of the cooperation of ovarian cell types in estrogen biosynthesis was provided by Falck in 1959 (203) in nowclassic experiments in which ovarian cell types in the rat were transplanted either alone or in combination into the anterior eye chamber together with estrogen-sensitive vag¬ inal epithelium as a biological indicator of estrogen pro¬ duction. These studies established that estrogen biosynthesis required the cooperation of granulosa or lutein cells with theca or interstitial cells. Falck mistakenly interpreted these findings as indicating a permissive influence of granulosa or lutein cells (perhaps mediated by progesterone) on es-

Follicular Steroidogenesis and Its Control / trogen secretion by theca or interstitial cells in the rat. Al¬ though this proposal was incorrect for the rat, where thecal biosynthesis does not contribute significantly to estrogen production, in at least certain species C2i-steroids from the granulosa cells may indeed be utilized by theca cells for androgen production, thereby contributing to estrogen pro¬ duction by both cell types. A two-cell theory of the sort suggested by Falck was subsequently proposed for the mare (187), with the addi¬ tional proposal that granulosa cells have only weak Hahydroxylase and little C-17,20-lyase activities. Further stud¬ ies with isolated ovarian cell types in the pig demonstrated that granulosa cells converted pregnenolone to progestins, but not to androgens, while readily interconverting exoge¬ nous testosterone and androstenedione (i.e., by the 173hydroxy steroid dehydrogenase) and aromatizing these an¬ drogens (193). It was later proposed that C19-steroid pre¬ cursors in the pig are produced by the theca and are then transferred to the granulosa for conversion to testosterone (188). In this way, estrogen was presumed to be synthesized by granulosa as well as theca cells, although aromatase activity in pig theca was believed to be higher (193). The next major step in the development of the modem concept of regulation of estrogen biosynthesis was the de¬ termination of the cellular sites of FSH and LH action. Much earlier work by Greep and coworkers (204) in hypophysectomized immature rats provided the first hints that FSH and LH act upon different cell types to promote estrogen for¬ mation. Subsequently, Hollander and Hollander (205) dem¬ onstrated that FSH action in vivo or in vitro stimulated [14C]testosterone conversion to estradiol by canine ovarian slices. However, up until the 1970s, LH was still considered to be the key, if not the only, steroidogenic hormone in the follicle for a variety of reasons. The action of LH via cAMP was becoming established as an important control mecha¬ nism, LH was known to stimulate both androgen and es¬ trogen production by luteal and stromal ovarian preparations (184), and it was later shown to stimulate acute estrogen production by the isolated rabbit follicle (206). Furthermore, use of relatively impure FSH cast doubts on the active hor¬ mone in earlier studies. The crucial role of FSH in con¬ trolling follicular estrogen biosynthesis in the rat was then established with the findings that explanted ovaries from hypophysectomized immature rats produced estrogen in re¬ sponse to FSH but not to LH and that they required the addition of testosterone as substrate (207). Similar findings with isolated rat granulosa cells in culture determined that FSH acted directly on the granulosa cells to stimulate aromatization (136), thereby extending earlier work demon¬ strating a similar action of FSH on testicular Sertoli cells (208). The results of these in vitro studies led to the formulation of a modem two cell-type, two gonadotropin theory in which theca interna cells (and perhaps also interstitial cells) are stimulated by LH to produce androgens, which in turn tra¬ verse the follicular basement membrane to be utilized for estrogen biosynthesis in an FSH-stimulated reaction within

345

the granulosa cells. Current evidence indicates that this con¬ cept is valid for antral and preovulatory follicles in the rat and most other species, despite additional and sometimes significant production of estrogens by theca cells in certain species. Evidence in various species of the exclusive lo¬ calization of FSH binding sites on granulosa cells and of LH binding sites initially on theca/interstitial and only later on granulosa cells supports the two-cell-type, two-gonadotropin concept described above. The in vitro evidence di¬ rectly supporting the concept of metabolic cooperation in estrogen biosynthesis by the two cell types is of three kinds. First, there is evidence that androgens produced in the theca layer exit from these cells in order to participate in estrogen biosynthesis, the inference being that extracellular andro¬ gens diffuse into the granulosa cell layer. Second, the evi¬ dence indicates that addition of aromatizable androgens to granulosa cells is essential for significant estrogen produc¬ tion, and addition of pregnenolone or other C2i precursors to thecal cells tends to increase androgen production. Third, recombination of isolated theca tissue and granulosa cells by coincubation demonstrates synergism in estrogen syn¬ thesis, provided that the two cell types either are stimulated with the appropriate gonadotropins in vitro or are derived from mature steroidogenically active follicles. This sup¬ porting evidence is discussed below. First, in vivo evidence that FSH and LH act at separate sites and by separate mechanisms in stimulating estrogen secretion has been obtained in studies with hypophysecto¬ mized rats. Treatment of hypophysectomized rats with LH has been demonstrated to enhance ovarian androgen (tes¬ tosterone and dihydrotestosterone) content; however, con¬ comitant administration of FSH was required to elevate es¬ tradiol levels. Substitution of LH with aromatizable androgen (testosterone or androstenedione) led to similar increases in estradiol production, provided that the rats had also been treated with FSH (209). Thus, these findings are consistent with a stimulatory action of LH on production of androgens that were then used as substrate for conversion to estradiol in the presence of FSH, thus providing in vivo support for the two-cell, two-gonadotrophin mechanism for control of estrogen biosynthesis in the rat. This cell-cooperation hypothesis requires that androgens of thecal origin must diffuse across the basement membrane separating the granulosa and thecal layers of the follicle in order for androgens to gain access to the aromatase enzyme in the granulosa cells. There is ample evidence that such diffusion does, in fact, occur. Concentrations of aromatiz¬ able androgens (i.e., androstenedione and testosterone) in follicular fluid are consistent with their serving as efficient substrates for aromatization by the granulosa cells. Fur¬ thermore, even higher concentrations of androgens might be achieved at the level of mural granulosa cells as the result of an androgen gradient, presumed to exist from the theca cells across the follicular basement membrane to the mural cells and finally to the follicular fluid. One study, using an experimental ovarian model in sheep, has provided indirect evidence that extracellular androgen produced in the theca

346 / Chapter 10 layer must cross into the follicle to allow estrogen biosyn¬ thesis by the granulosa cells. In this study, ovaries of ewes were autotransplanted to the carotid-jugular circulation, and endogenous LH pulses were observed to result in episodic secretion of estradiol into the jugular vein (210). Following infusion into the ovarian artery of a high-titer antiserum against androgen, the normally episodic secretion of estra¬ diol was inhibited, suggesting that passage of androgen across the follicular membrane was prevented by binding to anti¬ bodies. There is also indirect evidence in the pig that sug¬ gests thecal androgen transference to the granulosa cells (107). Various studies in pig (107), sheep (128), and human (99,124,135,191,211) provide the evidence that granulosa cells must be supplied with an extracellular source of aromatizable androgen substrate in order to synthesize estro¬ gens. Evidence of substantial theca-granulosa cell synergism in estrogen production in vitro has been obtained in co¬ culture studies with follicle cells of sheep (194), hamster (103), and human (122,124,135). Also, evidence of stromagranulosa synergism in estrogen production in the human was found in one study (124) but not in another (135). Similar studies in sheep have shown that follicle wall prep¬ arations (theca and granulosa) were far more effective in the production of estrogen than either cell type alone (128). In the rat, evidence of exclusive production of androgens by theca cells (212) and of aromatization in granulosa cells (141), combined with information that preovulatory estro¬ gen production by the follicle is regulated by LH-stimulated thecal androgen production (102), provides convincing sup¬ port for the original concept. Apart from estrogen biosynthesis, there are other forms of follicle cell cooperation in steroid production and me¬ tabolism. Theca-granulosa cell synergism has been found for the production of thecal androgen in the sheep (128) and human (124,135). Other studies in the mare (189,213), ham¬ ster (103), rat (214), and pig (215) provide evidence that C2i-steroid precursors, which may be produced in the gran¬ ulosa layer, can be metabolized to androgens by the theca cells. In other studies in the rat, comparisons of production of immunoreactive androgen (perhaps DHT) by intact pre¬ ovulatory follicles or isolated cell types suggest that coor¬ dinated activities of both granulosa and theca cells are re¬ quired for this biosynthetic function (102). Furthermore, theca-granulosa (124,135), as well as stroma-granulosa (135), synergism is also involved in progesterone synthesis in the human.

Intraovarian Regulation by Follicular Steroids Estrogens Early experiments demonstrated that estrogens had a stim¬ ulatory effect on the ovary (216-223). In experiments in which estrogen was administered to hypophysectomized im¬ mature rats, ovarian weight was maintained and ovaries

became more responsive to gonadotropic stimulation. Fur¬ thermore, the ovarian weight response to gonadotropins was inhibited by antiserum to estradiol (224). Treatment of hy¬ pophysectomized rats with gonadotropins increased the number of atretic follicles per ovary, and administration of estrogen partially reversed this effect (225). Specific uptake and retention of [3H]estradiol in vivo by ovaries of immature rats (226,227), incorporation of estra¬ diol into granulosa cells (228), and binding of estradiol to nuclear fractions of rat granulosa (229) and luteal cells (230) indicate the presence of estrogen binding sites in the ovary. Saidudduin and Zassenhaus (231) have characterized estro¬ gen binding components from ovaries of immature rats. Their results suggested that ovarian estrogen binding sites are similar to specific, high-affinity estrogen receptors in uterine tissue. There is evidence suggesting that estrogen acts within the ovary to inhibit androgen production, the estrogen presum¬ ably originating from granulosa cells and androgen from theca. Treatment of intact immature rats with estradiol sup¬ pressed ovarian testosterone and 5a-dihydrotestosterone production. Concomitant administration of gonadotropins failed to overcome the inhibition by estrogen, indicating that the effect was not mediated by decreased circulating gonadotropin (232). To provide further evidence for a direct intraovarian action of estrogen, Silastic implants of estradiol were embedded under the ovarian bursa unilaterally. Under these conditions, LH-stimulated androgen content of the ipsilateral ovary was considerably lower than that of the contralateral ovary (232). An inhibitory effect of estradiol on LH-induced androgen content of ovaries from immature hypophysectomized rats was reported as further evidence that a pituitary factor was not involved (232). Results of in vitro experiments have demonstrated similar effects of estrogen on ovarian androgen synthesis. Whole ovaries from estrogen-treated immature, intact, or hypophy¬ sectomized rats responded to LH-stimulation in vitro with decreased androgen production when compared to that of ovaries obtained from rats that were not treated with estro¬ gen. Dibutyryl cAMP did not increase testosterone produc¬ tion by cultured ovaries of estrogen-pretreated rats (233). Estradiol treatment of immature rats in vivo also suppressed androgen secretion by isolated thecal tissue in vitro (234). In culture experiments with isolated porcine thecal tissue (234) or dispersed theca cell preparations (235), addition of estrogens directly to the culture medium inhibited LH-stim¬ ulated androgen production in a dose-dependent manner, establishing that the inhibitory action of estrogens is directly on theca cells. All indications are that estrogen inhibits ovarian androgen synthesis at a site distal to cAMP production, probably at an enzymatic step(s) in the steroidogenic pathway between androgens and their C2i precursors. In accordance with this hypothesis, estrogen pretreatment of ovaries from intact im¬ mature rats in vivo has been shown to inhibit conversion of radioactively labeled progesterone to androgens (testoster¬ one, androstenedione, and androsterone) in vitro. On the

Follicular Steroidogenesis and Its Control / other hand, incorporation into 3a-hydroxy-5a-pregnan-20one is enhanced, suggesting that estrogen may act by in¬ hibiting the 17a-hydroxylase:C-17,20-lyase enzyme system or by diverting C2i substrates into a pathway resulting in the formation of 5a-reduced pregnane compounds (233). In another study, treatment of immature rats with estradiol suppressed the stimulation by hCG of androstenedione, tes¬ tosterone, 17a-hydroxyprogesterone, and 17a-hydroxypregnenolone production by dispersed ovarian cells in cul¬ ture. Pregnenolone production was unchanged, while progesterone production was markedly enhanced. Estradiol had no effect on hCG binding capacity, hCG-stimulated cAMP synthesis, or the viability of ovarian steroidogenic cells. It was concluded that exogenous estradiol blocked ovarian androgen formation by reducing the activity of the 17a-hydroxylase enzyme (236). Further in vitro evidence with rat ovarian interstitial cell cultures indicates that es¬ tradiol directly causes rapid inhibition of 17a-hydroxylase and C-17,20-lyase enzyme activities (237). There is evidence that estrogens are capable of regulating metabolism of androgens by a direct action on 5a-reductase. Eckstein and Nimrod (238) have shown an inhibitory effect of estradiol on 5a-reductase activity in microsomal prepa¬ rations from immature rat ovaries. Since the minimal ef¬ fective concentration of estradiol required to inhibit enzyme activity was in the range measurable in follicular fluid, the authors suggested that estradiol may be physiologically sig¬ nificant in the regulation of androgen metabolism. Administration of LH to intact immature rats has been shown to affect ovarian progesterone metabolism in a man¬ ner identical to that of estrogen. Measured in vitro, ovarian androgen production was reduced and 3a-hydroxy-5a-pregnan-20-one secretion stimulated (233). Exposure of ovaries isolated from prepubertal rats to LH alters progesterone metabolism, favoring formation of 5a-reduced pregnane compounds while decreasing androgen and 5a-reduced an¬ drogen biosynthesis (239). In cultured preovulatory follicles from PMSG-treated immature rats, LH inhibited C-17,20lyase activity. Addition of inhibitors of steroid synthesis prevented the inhibitory action of LH on the conversion of 17a-hydroxyprogesterone to androgens but did not affect basal lyase activity. These experiments suggested that the inhibitory action of LH on androgen synthesis may be me¬ diated by the action of another ovarian steroid. That this steroid is estrogen is supported by experiments showing that the aromatase inhibitor, 4-acetoxy-androstane-3,17-dione, blocks the negative effect of LH on androgen synthesis by rat preovulatory follicles in vitro (240). After the ovulatory surge of LH, androgen levels in ovar¬ ian tissue, follicular fluid, and ovarian venous blood and serum initially rise and then fall precipitously several hours later (241-243). This apparent inhibitory effect of LH on androgen production raises the possibility that this inhibition represents a physiological role of estrogens in the intrafollicular control of androgen biosynthesis. In support of this possibility, Smith et al. (244) and Kalra and Kalra (245) have measured serum hormone concentrations during the

347

rat estrous cycle and have shown that, shortly after the LH surge on the day of proestrus, estradiol peaks and then rapidly declines. That the rise is dependent on the LH surge has been demonstrated in proestrous hamsters by blocking LH secretion with injections of phenobarbital (246). It may be that the surge of LH initially stimulates theca cells to produce androgens, which are aromatized to estrogens by granulosa cells. The estrogens then inhibit the thecal 17ahydroxylase.C-17,20-lyase enzyme system, thereby limit¬ ing further synthesis of androgens and their subsequent use as substrates for aromatization. The ability of low doses of estradiol to enhance progesterone production by isolated bovine theca cells (247) may be a reflection of precursor accumulation following the inhibitory action of estrogens. This action may contribute to the transition of the follicle from primarily an estrogen-secreting to a progesterone-se¬ creting structure, initiated by the LH surge. The role of estrogens in regulation of the corpus luteum is beyond the scope of this chapter and will be discussed elsewhere in the volume. Another intrafollicular regulatory action of estrogens that has been clearly established is their ability to enhance ovar¬ ian estrogen production through direct actions on granulosa cells. Clomiphene citrate, a weak estrogen, increased es¬ tradiol and estrone synthesis from radiolabeled androstene¬ dione by superfused canine ovaries in vivo (248). Clomi¬ phene citrate (249), estradiol, estrone, hexestrol, moxestrol, ethinyl estradiol, chlorotrianisene, mestranol (250), and triphenylethylene antiestrogens (251) have also been reported to have similar effects on FSH-induced estrogen synthesis by cultured granulosa cells isolated from ovaries of im¬ mature hypophysectomized, DES-primed rats. FSH-in¬ duced aromatase activity in rat granulosa cells was enhanced by in vitro addition of DES (252). Good correlation was found between receptor binding affinity and biological po¬ tency of both natural and synthetic estrogens or antiestro¬ gens, and the stimulatory effect could not be accounted for by increased granulosa cell viability or protein mass (250). In support of the hypothesis that estrogens are physio¬ logical regulators of granulosa cell aromatase activity, the minimal effective dose of estradiol required to elicit a re¬ sponse (3.7 x 10-10 m) is well within the range of estradiol in antral fluid of preovulatory follicles (250). Thus, estro¬ gens may function within the ovary or in individual follicles as end-product amplifiers to enhance FSH-induced aroma¬ tase. Estrogens have been shown to decrease progesterone se¬ cretion by porcine (253-256), bovine (247), rat (257), and human (258,259) granulosa cells and by large follicles from bovine ovaries (260). The inhibitory action of estrogens on porcine granulosa cells was both time-and dose-dependent and was demonstrable in short-term, but not long-term, cultures at estradiol concentrations similar to those found in vivo (261). Estrogen-inhibition of progesterone produc¬ tion was not dependent on cell density in culture or due to a cytotoxic effect, degree of follicular maturation, or ac¬ celerated metabolism of 20a-dihydroprogesterone. Instead,

348

/

Chapter 10

the action of estrogen appeared to limit the conversion of pregnenolone to progesterone, resulting in enhanced preg¬ nenolone accumulation in culture. Increased pregnenolone production in the presence of estrogen was also the result of enhanced cholesterol side-chain cleavage activity (154,261) and mitochondrial content of P-450 (154). The inhibitory action of estrogen on progesterone pro¬ duction has been corroborated in vivo. Administration of estradiol for 3 days decreased LH-induced ovarian proges¬ terone content in hypophysectomized immature rats but not in intact animals. Also, in ovaries of estradiol-treated hy¬ pophysectomized rats, dibutyryl cAMP, but not LH, re¬ stored in vitro progesterone production to values comparable to those of ovaries from control animals. This result suggests that estrogen inhibits progesterone synthesis by acting at a step early in the stimulative cascade, possibly prior to cAMP generation (262). There are also reports of stimulatory effects of estrogen on progesterone secretion in porcine (24,263) and rat (257,264,265) granulosa cells. Unlike the inhibitory action of estrogen, the stimulatory effect on porcine granulosa cells in culture was only demonstrable in longer-term incubations and was found to be dependent on the density of granulosa cells in culture and the maturational status of the follicle from which cells were isolated (261). Granulosa cells from small, but not larger, follicles responded to estrogen with increased progesterone secretion, an effect that was found to be due to increased activity of A5-3(3-hydroxysteroid dehydrogenase:A5_4-isomerase. In addition, with estrogen treatment, pregnenolone accumulation was increased, as were cholesterol side-chain cleavage activity and hydrolysis of endogenous cholesterol esters. Catechol estrogens, which may be synthesized within the follicle (49), have also been shown to stimulate steroido¬ genesis in rat granulosa cells in vitro (266) and corpora lutea (267).

Androgens With the establishment of the obligatory role of androgens as substrates for estrogen biosynthesis, other possible roles of androgens in follicular function at first received little attention or were dismissed. The antiandrogen, hydroxy flutamide had little or no effect on FSH-induction of enzyme activity or LH receptors in diethylstilbestrol (DES)-primed, hypophysectomized immature rats, suggesting that andro¬ gens are not essential for FSH to initiate development of antral follicles (268). In support of this hypothesis, Neu¬ mann et al. (269) demonstrated that cyproterone acetate, another antiandrogen, did not disrupt estrous cycles or in¬ terfere with ovulation in adult rats. In addition, Lyon and Glenister (270) have reported that Tfm/O mice, a strain in which females carry a gene conferring androgen resistance, have normal reproductive cycles; follicular maturation, con¬ ception, and pregnancy occur. More recently, however, evidence of several sorts has

appeared providing convincing evidence of other regulatory functions of androgens in the follicle. Specific androgen binding sites have been identified in ovaries from estrogenprimed, hypophysectomized immature rats (271) and later localized to the granulosa cell compartment (272). Similar androgen-binding proteins are found in sheep granulosa cells (273) and human ovarian cytosol (274). In healthy, rather than atretic, follicles there is an inverse relationship between intrafollicular concentrations of an¬ drogens and estrogens. High ratios of androgens to estrogens in follicular fluid have been associated with nonovular and atretic follicles (229,275,276). However, from these data it is difficult to determine whether the predominance of an¬ drogen over estrogen is the cause of atresia or merely a result of the process. Both androgens (221,277) and hCG (278), which stimulate ovarian androgen synthesis, have been shown to promote atresia in rats. Coadministration of antiandrogens or antiserum raised against androgen (278) alleviated this effect of hCG. Further, treatment of imma¬ ture, hypophysectomized, PMSG-treated rats with DHT in¬ duced atresia (279). This latter effect could be at least par¬ tially overcome by estradiol. Recently, Opavsky and Armstrong (280) showed an inhibitory effect of LH on the superovulatory response of immature rats to FSH. Although there is considerable circumstantial evidence to implicate androgens in the process of follicular atresia, the mecha¬ nisms of this process and the specific role of androgens in controlling the process are poorly understood. In view of the rather substantial evidence in favoring a negative role for androgens in follicular maturation, it is perhaps surprising to find that androgens have positive ef¬ fects on follicular growth. Ovarian degeneration occurred in androgen-resistant Tfm/O mice (281), and an antiandro¬ gen accelerated atresia in preovulatory rat follicles (282). Somewhat similarly, treatment of diestrous rats with flutamide resulted in decreased growth and maturation of ovar¬ ian follicles (283). Also, an inhibitory action of androgen antisera on hCG-induced ovulation has been reported in hypophysectomized rats (284). In trying to reconcile the apparent discrepancies between androgen effects within the ovarian follicles, it is worth noting that the antagonistic action of androgen on follicular events, as has been implicated in atresia, is believed to affect only those follicles at the preantral and early antral stages of development. The facilitory effect of androgen may be reserved for those large follicles that have already entered the final stages of development (285). In culture, androgens have been shown to stimulate pro¬ gesterone biosynthesis by intact follicles dissected from ova¬ ries of cycling ewes (286) and cows (260) and by granulosa cells isolated from pig (255,256), rat (257,287) and mouse (288) ovaries. In addition, administration of androgens to intact immature rats increased subsequent progesterone ac¬ cumulation by their isolated ovarian cells in vitro (289). Since both aromatizable and nonaromatizable androgens were effective, the response appears to be androgenic rather than dependent on aromatization of androgens to estrogens. This

Follicular Steroidogenesis and Its Control / hypothesis is supported by data for the pig, in which im¬ plants of antiandrogens (flutamide or hydroxyflutamide) placed in the ovarian interstitium decreased progesterone secretion by isolated granulosa cells in vitro (290). Hydroxyflutamide and cyproterone acetate suppressed the stimulatory effect of testosterone on progesterone production by rat granulosa cell incubations (257). Androgens, in addition to having their own stimulatory effects, enhance FSH-stimulated progestin synthesis (MSMS,291). This action was blocked by hydroxyflutamide (292), and the potency of various androgens appeared to be correlated with the extent to which they were converted to testosterone or DHT (195). Depending on the animal model used, androgens have been shown to act at both pre- and post-cAMP sites. Using granulosa cells isolated from ovaries of immature hypophysectomized, estrogen-treated rats, androstenedione en¬ hanced stimulation of progestin production by the cAMP analog dibutyryl cAMP but had no effect on [125I]FSH bind¬ ing to the cells, FSH-stimulated cAMP production, or con¬ version of cAMP to AMP by the phosphodiesterase enzyme (144). On the other hand, in granulosa cells isolated from ovaries of intact immature rats, androgens enhance FSHresponsiveness, as measured by the FSH stimulation of cAMP production (292-294) and [125I]FSH binding (294,295), and suppress cAMP metabolism (292). Increased C2i-steroid production in the presence of an¬ drogen is not a reflection of decreased catabolism by 5areductase (144), although androgens have been reported to regulate catabolism of progesterone (296,297) through in¬ hibition of 20a-hydroxysteroid dehydrogenase (298). An¬ drogens alone or in combination with FSH have no effect on levels of free or esterified cholesterol in cultured rat granulosa cells (299). However, in these cells, androgen and FSH act synergistically to enhance lipoprotein utiliza¬ tion (300). Both FSH and testosterone independently en¬ hance conversion of cholesterol to pregnenolone, indicating a stimulatory action on cholesterol side-chain cleavage. Combined treatment results in synergism (153,291,299). Transport of cholesterol into mitochondria is unaffected by FSH or androgen (299). Effects of androgens on 3(3-hydroxysteroid dehydrogenase are equivocal. Some data sug¬ gest that androgens act synergistically with FSH to increase conversion of pregnenolone to progesterone (291), while other data indicate that androgens are ineffective at this site (Ml). In contrast to the work using rat tissue, aromatizable androgens have a negative influence on progesterone pro¬ duction by human granulosa cells (124) and on FSH-stim¬ ulated progesterone accumulation by granulosa cells isolated from porcine ovaries (171,301). Although estradiol had a similar inhibitory action and nonaromatizable androgens were ineffective, the effect of aromatizable androgens could not be accounted for by conversion to estrogens since 4-acetoxy4-androstene-3,17-dione, an inhibitor of aromatase, failed to prevent the testosterone-induced decrease in progesterone production (301).

349

Similar to the action of FSH, dibutyryl cAMP stimulated progesterone production by porcine granulosa cells. Addi¬ tion of testosterone to cultures suppressed the stimulatory effect of dibutyryl cAMP, indicating that testosterone acts at a site distal to cAMP generation (171). Further studies revealed that androgens had no effect on progesterone me¬ tabolism (301); however, they did enhance pregnenolone synthesis in FSH-treated granulosa cell cultures (171). Thus, decreased progesterone production in the presence of an¬ drogen appears to be due to restricted conversion of preg¬ nenolone to progesterone through inhibition of A5-3(3-hydroxysteroid dehydrogenase:A5~4-isomerase activity. A direct inhibitory action of testosterone on this enzyme has recently been demonstrated (302). Very little work has been done to investigate intraovarian effects of androgens on thecal steroidogenesis. Androgen has been shown to enhance progesterone secretion by human thecal tissue in culture. The effect on granulosa cell pro¬ gesterone synthesis was negative, and in combined incu¬ bations of theca or stroma plus granulosa cells, no effect of androgen could be discerned (124). As discussed above, FSH has a regulatory role in ovarian estrogen secretion through induction of aromatase activity in rat granulosa cells. Studies using cultured rat granulosa cells from intact immature rats have demonstrated that, in addition to acting as substrates for FSH-stimulated aroma¬ tase, androgens also enhance FSH-induction of enzyme ac¬ tivity (303). Both aromatizable and nonaromatizable andro¬ gens were effective, although nonaromatizable androgens (5a-dihydrotestosterone and androsterone) were only 50% as potent as aromatizable androgens (testosterone and an¬ drostenedione). Thus, it is clear that this action of androgens is not dependent on their conversion to estrogens. This con¬ tention is supported by experiments showing that androgen enhancement of FSH-induced aromatase activity is sup¬ pressed by hydroxyflutamide (an androgen receptor blocker) (304,305) and is not affected by 4-acetoxy-4-androstene3,17-dione (an inhibitor of aromatase) (252) or nafoxidine (an estrogen receptor blocker) (252). There is evidence to suggest that, in the intact immature rat model, androgens influence granulosa cell aromatase activity by action at a site before cAMP production. Al¬ though testosterone enhances FSH-induced aromatase ac¬ tivity, it has no effect on cAMP-induced estrogen synthesis. Androgens enhance the responsiveness of cultured granulosa cells to FSH in the production of cAMP, as well as in stimulating cellular [125I]FSH binding (294). Aromatization of testosterone to estradiol by granulosa cells, isolated from the largest follicles in ovaries of rats showing diestrous II and proestrous vaginal smears, has been shown to be competitively inhibited by 5a-reduced androgens (306) The same phenomenon was described us¬ ing human follicles excised at all stages of maturity (307). Apart from alterations in aromatase enzyme activity itself, follicular estrogen biosynthesis may be influenced by the amount of C19 substrate available and variation in the ratio of aromatizable to nonaromatizable androgens as the result

350

/

Chapter 10

of changes in 5a-reductase activity. Rat (40) and human (110,308) ovaries have been shown to convert aromatizable androgens actively to 5a-reduced androgens. Since high concentrations of estrogen in follicular fluid have been as¬ sociated with healthy antral follicles and low estrogen with apparently degenerating follicles (275), alterations in the capacity of granulosa cells to convert androgens to estrogens may be a physiologically important mechanism for regu¬ lating concentrations of intrafollicular steroid hormones and development of individual follicles. Siiteri and Thompson (309) have reported a 2.5-fold in¬ crease in 5a-reductase activity and a 5-fold decrease in aromatase activity within a few hours of exposure of ovaries of PMSG-treated rats to hCG. Katz and Armstrong (310) observed a similar decline in aromatase activity following LH treatment. The increased activity of 5a-reductase could contribute to the decreased estradiol secretion, which occurs dramatically following the LH surge, in two ways: (a) by decreasing the intrafollicular levels of aromatizable andro¬ gens through increased 5a-reductase activity and (b) by increasing intrafollicular levels of 5a-reduced androgens, which serve as competitive inhibitors of the aromatase en¬ zyme system.

Progestins It is uncertain whether progestins have any direct role in the intraovarian regulation of ovarian function, and reports are often contradictory. Some investigators have found that administration of exogenous progesterone to estrogen-primed, hypophysectomized immature rats had no effect on ovarian morphology (229,311,312), while others have presented evidence that progesterone has an inhibitory effect on fol¬ licular development when given to intact animals (313— 316). That the effect of progesterone in these latter exper¬ iments might be mediated indirectly by depression of pi¬ tuitary secretion of gonadotropins, rather than direct inhi¬ bition at the follicular level, is suggested by the observation that retardation of follicular growth by progesterone occurs only when plasma concentrations of both FSH and LH are significantly reduced (316). Goodman and Hodgen (317) attempted to avoid this problem by placing progesterone directly in the monkey ovary and suggested that their results supported a direct inhibitory action of progesterone on fol¬ licular development. In prepubertal rats, progesterone implants decrease serum LH concentrations and reduce estradiol accumulation by the isolated follicles but, surprisingly, facilitate the stimulatory effects of low-dose hCG treatment on the growth of small antral follicles and on estrogen synthesis (318). It was sug¬ gested that progesterone may facilitate LH action under physiological circumstances when basal LH is low. On the other hand, even in the presence of elevated serum proges¬ terone, preovulatory follicular maturation at the end of preg¬ nancy (i.e., follicles functionally indistinguishable from those

at proestrus) is supported by small sustained increases in serum LH, suggesting that progesterone may have no direct inhibitory effect on follicle cell maturation (319). In support of a direct action of progesterone on ovarian regulation, specific progesterone receptors have been iden¬ tified in rat ovary (320,321) and subsequently in rat gran¬ ulosa cells (322,323). Similarly, ovarian progestin binding sites have been reported in rabbit (324), guinea pig (325), cow (326), and human (274,327). Progesterone has been shown to enhance the ability of cultured rat granulosa cells to respond to FSH in the pro¬ duction of cAMP (293). In another study, synthetic pro¬ gestin (R5020) increased FSH-stimulation of progesterone and 20a-dihydroprogesterone synthesis in granulosa cells isolated from immature hypophysectomized, estrogen-treated rats. Similarly, R5020 enhanced LH-stimulated progestin production by cells that had previously been primed with FSH to induce LH-receptor formation. Furthermore, in the presence of cyanoketone, an inhibitor of A5-3p-hydroxysteroid dehydrogenase, progesterone augmented the ability of FSH to stimulate pregnenolone synthesis (328). The ma¬ jor criticism of this latter work is that the concentration (1 X 10-6 m) of synthetic progestin required to elicit a response is at the extreme upper limit of physiological progestin con¬ centrations. The possibility that the action of progestin was mediated by nonspecific binding to androgen receptors was considered; however, the apparent autoregulatory actions of progestins were not altered by treatment with antiandrogens, indicating that the effect was not due to binding of progestin to androgen receptors (329). The effects of several progestins on FSH-stimulated es¬ trogen production by cultured rat granulosa cells isolated from ovaries of immature hypophysectomized, DES-treated rats have been examined (330). FSH-enhanced estrogen se¬ cretion was reduced following treatment with progesterone, 20a-dihydroprogesterone, or R5020, a potent synthetic pro¬ gestin. A study of the relative potencies of the progestins revealed that R5020 was the most effective compound, fol¬ lowed by progesterone and 20a-dihydroprogesterone. This pattern was found to reflect the relative abilities of the pro¬ gestins to bind to ovarian progestin receptors. Later exper¬ iments investigated the mechanism by which R5020 inhibits FSH-induction of aromatase activity. It was concluded that the synthetic progestin acts at a site distal to cAMP pro¬ duction and that it is not a competitive inhibitor of aromatase (331) . In another study of the effect of progesterone on FSHstimulated estradiol synthesis by cultured rat granulosa cells, it was questioned whether progesterone has the same effect on aromatase activity once activity has been induced in vivo (332) . In granulosa cells isolated on the morning of proestrus from follicles of immature rats previously treated with PMSG (4 IU), progesterone had a slight suppressive effect on es¬ tradiol synthesis. However, it was evident that once aro¬ matizing activity had been induced estradiol synthesis was much less sensitive to inhibition by progesterone.

Follicular Steroidogenesis and Its Control

Likewise, an inhibitory effect of progesterone on estrogen secretion has been demonstrated in vivo (246). Progesterone administered to hamsters on the morning of the day of proestrus resulted in a fall in serum estradiol concentration with¬ out a concomitant change in blood levels of gonadotropins. The fact that concomitant administration of testosterone did not reverse the effect of progesterone indicated that pro¬ gesterone acted at the level of the aromatase enzyme system. This study led the authors to speculate that the inhibitory effect of progesterone is one factor in the sharp decline in the serum concentration of estrogen that occurs after the LH surge in normally cycling hamsters (333). Regulation of androgen production in the ovary by progestins has not been reported; however, both progesterone (334) and 5a-pregnane-3,20-dione (335) are effective in¬ hibitors of the C-17,20-lyase, indicating their potential in intraovarian regulation of androgen biosynthesis.

Other Regulatory Factors and Hormones Prolactin Prolactin has long been known as a “luteotrophic” hor¬ mone, particularly in rodents but also in several other spe¬ cies. As such, it is involved in initiating luteinization of granulosa cells, in maintaining their level of progesterone synthesis as luteal cells, and in inhibiting the activity of the progesterone catabolizing enzyme, 20a-hydroxysteroid de¬ hydrogenase, the latter particularly in rodents (336). The appearance of specific prolactin receptors in granulosa cells late in follicular development and their induction by FSH in culture (249) indicate the likelihood that prolactin may exert a physiological action on granulosa cells at the stage of terminal differentiation, when they are transformed into luteal cells. In support of this, prolactin has been demon¬ strated to enhance progesterone production in cultured gran¬ ulosa cells obtained from preovulatory rat follicles (337) and porcine follicles (338). Striking morphological changes, characteristic of luteinization, were also induced by prolac¬ tin in cultured rat granulosa cells (337). Stimulatory effects of prolactin on steroidogenesis in pre¬ pubertal (i.e., nonluteinized) ovaries have also been re¬ ported. Thus, prolactin injections (339) or hyperprolacti¬ nemia induced by in vivo administration of dopaminergic receptor blockers (339,340) have been found to induce pre¬ cocious puberty, as well as to increase ovarian responsive¬ ness to LH in immature rats. The latter effect appeared to be mediated, in part, by an increase in ovarian LH receptors (339). There is also evidence for a role of prolactin in induction and maintenance of LH receptors on luteal cells at late gestational stages in the rat (341). In contrast to the stimulatory action of prolactin on pro¬ gesterone secretion by granulosa cells at a late stage of differentiation, progesterone production by granulosa cells from small immature porcine follicles was markedly inhib¬

/

351

ited by physiological concentrations of prolactin (338). Ex¬ posure of the latter granulosa cells to estradiol reversed this inhibitory effect (342). Another inhibitory effect of prolactin, on estrogen secre¬ tion, was reported for cultured rat granulosa cells obtained from follicles at both preantral and preovulatory stages (343,344). Decreased estrogen secretion in vitro appears to be due, at least in part, to an inhibitory action of prolactin on FSH induction of aromatase activity (345,346). The site of this action of prolactin appears to be distal to adenylate cyclase, as prolactin also inhibited the stimulatory action of dibutyryl cAMP. The inhibitory effect of prolactin on rat follicle aromatase activity has also been demonstrated by in vivo exposure of intact rats to the hormone (347). In ad¬ dition, prolactin has been reported to suppress basal and gonadotropin-stimulated estradiol secretion by human ova¬ ries perfused in vitro (348). Evidence for an inhibitory influence of prolactin on an¬ drogen secretion, presumably by action on the theca and/or interstitial cells, of rat ovaries has also been reported. Levels of androstenedione in preovulatory follicles of adult rats were significantly decreased by prolactin (347), whereas hypoprolactinemia, induced in vivo by bromoergocryptine, was accompanied by markedly increased secretion of 5aandrostane-3a, 17(3-diol, the major androgen secreted by the prepubertal rat ovary in response to hCG stimulation (349). Addition of prolactin to cultured rat theca-interstitial cell preparations at concentrations within the physiological range for the female rat, and consistent with the binding affinity observed for the specific binding sites (receptors) that have been found on rat theca-interstitial cells, caused a dosedependent inhibition of LH-stimulated androgen formation (androsterone, 5a-androstandiol) (350). As with granulosa cells, the inhibitory action of prolactin on theca-interstitial cells appears to be exerted at a step after adenylate cyclase, since the stimulatory action of 8-bromo-cAMP, as well as of other activators of adenylate cyclase (PGE2, choleratoxin), was similarly inhibited by prolactin.

Gonadotropin-releasing Hormone-like Peptides Evidence that systemic administration of gonadotropin¬ releasing hormone (GnRH) or its agonists paradoxically inhibit reproductive functions (351) was first attributed exclusively to indirect effects on the hypothalamic-pitu¬ itary-gonadal axis. There is indeed evidence showing that continuous or intermittent administration of GnRH or its agonists can cause the pituitary to become refractory to the releasing hormone (352-355) and, through the release of gonadotropins, can induce ovarian receptor loss and desen¬ sitization to these hormones (356-359). However, numer¬ ous other studies, particularly in the rat, have since estab¬ lished that GnRH binding sites are present in various extrapituitary tissues, including the gonads, and that direct actions of GnRH on somatic cells of ovary and testis occur

352

/

Chapter 10

(329,360). Furthermore, products with biological activities similar to those of hypothalamic GnRH have been reported in the ovary, testis, placenta, pancreas, other central nervous system (CNS) structures, and in some tumor types (360). We will review only those studies pertinent to possible intraovarian actions of GnRH-like peptides. Ovarian cell types in the rat that have been demonstrated to have specific high-affinity binding sites for GnRH are granulosa (361,362), luteal (362,363), and theca (364) cells, at all stages of differentiation (364). These binding sites, presumed to be receptors by virtue of the various biological effects reported, are localized in the plasma membrane (365). In granulosa cells, GnRH receptors have been shown to be regulated by GnRH itself (inhibitory and stimulatory effects) and maintained by FSH (362,366). Two binding compo¬ nents have been identified in granulosa cells, of which one is similar to the pituitary GnRH receptor (367,368). Only binding sites of low affinity (369,370) or moderate affinity (371) have been found in human corpus luteum (369,370), consistent with the prolonged treatment required to observe inhibitory effects on progesterone biosynthesis by human granulosa cells in vitro (372). One study reports the absence of inhibitory effects of GnRH on progesterone biosynthesis by human granulosa cells (373). Biological effects of GnRH on ovarian cells in rabbit (374), pig (375), and chicken (376,377) also suggest the presence of GnRH-sensitive mechanisms. Lack of ovarian inhibition by GnRH in mice (378) and rhesus monkeys (379) and the absence of specific binding sites in ovine, bovine, and porcine ovaries (380) and in monkey corpora lutea (381) suggest that these species may not share the same GnRH sensitivity found in the rat. The first evidence for a direct extrapituitary effect of a GnRH agonist on the rat ovary was provided by the study of Rippel and Johnson (382), which showed inhibition of the hCG-stimulated ovarian and uterine weight gains in im¬ mature hypophysectomized animals. These findings were substantiated by extrapituitary inhibitory effects of GnRH and its agonists on FSH-stimulated ovarian weight gain and aromatase activity in immature hypophysectomized, estro¬ gen-treated rats and on FSH-stimulated estrogen and pro¬ gesterone production in cultured rat granulosa cells (383,384). GnRH inhibition of 3H20 release from [l(3-3H]testosterone, as a measure of aromatase activity, has also been demon¬ strated in cells from immature estrogen-primed rats (385,386). Other studies in hypophysectomized, PMSG-treated female rats demonstrated that a potent GnRH agonist also inhibited follicular maturation and steroidogenesis (387). The effect of GnRH in decreasing FSH-stimulated pro¬ gesterone production appears to be due to actions at several sites, causing inhibition of pregnenolone production (pre¬ sumed to be an effect on the cholesterol side-chain cleavage enzyme), inhibition of A5-3(3-hydroxysteroid dehydroge¬ nase activity (153,386,388), and stimulation of 20a-hydroxysteroid dehydrogenase activity (389), thereby reducing progesterone production and increasing its conversion to 20a-dihydroprogesterone. Inhibitory influences of GnRH on

LH/hCG-stimulated estrogen and progesterone production were also demonstrated in cultured rat granulosa cells pre¬ viously primed with FSH (164,390). Differences in the mechanisms of inhibition of progesterone production were seen with different stimulatory agents. Thus, the inhibitory action on LH/hCG-stimulated progesterone production ap¬ peared to involve decreased pregnenolone production (164), whereas the inhibition of prolactin and (3-adrenergic agoniststimulated progesterone production appeared to be the result of an action of GnRH to increase the activity of 20a-hydroxysteroid dehydrogenase, without influencing pregnen¬ olone production (390,391). Direct effects of GnRH on steroidogenesis in granulosa cells are not limited to inhibition. In the absence of other stimulatory agents, GnRH and its agonists act on granulosa cells from normal or immature hypophysectomized, estro¬ gen-primed rats to increase aromatase activity (386) and the production of pregnenolone, progesterone, and 20a-dihydroprogesterone (153,170,386,388,392). These and other studies indicate that GnRH stimulates the cholesterol sidechain cleavage enzyme (153,170), the A5-3(3-hydroxyste¬ roid dehydrogenase (386,388), and 20a-hydroxysteroid de¬ hydrogenase (173). However, GnRH and GnRH agonists do not activate steroidogenesis nearly as well as FSH does. The direct effects of GnRH, either inhibitory or stimulatory, on follicular steroidogenesis in vivo appear to depend on the duration of exposure and on the stage of follicular mat¬ uration (393,394). GnRH also has diverse effects on other aspects of gran¬ ulosa cell cytodifferentiation that may be responsible for some or all of the observed influences on steroidogenesis. FSH-induced formation of LH and prolactin receptors (384,395) and stimulation of receptors for epidermal growth factor (EGF) on rat granulosa cells is inhibited by GnRH and its agonists (396). GnRH also inhibits FSH-stimulated cAMP production in rat (397-399) and porcine (375) gran¬ ulosa cells. This effect in rat granulosa cells is apparently due to two actions: (a) inhibition of adenylate cyclase (399), which has been attributed to a decrease in the FSH receptor content and to an inhibition of the FSH-regulated increase in its own receptor (400); and (b) stimulation of extracellular phosphodiesterase activity (399), although there are indi¬ cations that intracellular and total phosphodiesterase activity is decreased by GnRH (401). The inhibition of adenylate cyclase by GnRH is dependent on the type of activating stimulus, since stimulation by isoproterenol or prostaglandin E is not inhibited (400). In contrast, following FSH-stimulation the subsequent increase in cAMP production induced by LH, isoproterenol, or PGE2 is inhibited by GnRH, in a process requiring calcium (402). Since the inhibitory effects of GnRH can be seen when steroidogenesis is stimulated by cAMP analogs (or by various agents that stimulate cAMP production), it is probable that GnRH causes these effects by actions at site(s) distal to cAMP production. A GnRH agonist partially inhibits dibutyryl cAMP-stimulated aro¬ matase activity in rat granulosa cells in the presence of an

Follicular Steroidogenesis and Its Control

inhibitor of phosphodiesterase, indicating that cAMP ca¬ tabolism is unlikely to fully explain the inhibitory actions (385). Inhibition of the FSH-stimulated increase in cAMP binding sites in rat granulosa cells (170) might be one ad¬ ditional factor in the direct inhibitory effects of GnRH. Other FSH-stimulated responses that are augmented by GnRH include cellular protein production (403) and pros¬ taglandin synthesis (397). GnRH alone also stimulates cel¬ lular protein content (404) and production of lactate (405), plasminogen activator (406), prostaglandins (392), and fibronectin (407). Additional effects of GnRH on phospho¬ lipid labeling from 32P and phosphatidylinositol metabolism (408—412) and on arachidonic acid release (413) suggest that pathways involving phosphoinositide metabolism, per¬ haps related to calcium mobilization and calcium-activated enzymes (414,415), may mediate some of the actions of GnRH at sites distal to cAMP. Neither prostaglandins nor cAMP appears to mediate stimulatory effects of GnRH on progesterone production by granulosa cells from preovu¬ latory rat follicles (416), although a GnRH agonist in the presence of a phosphodiesterase inhibitor has been found to cause very low levels of cAMP to accumulate in the culture medium of granulosa cells from immature estrogen-treated rats (386). The independent stimulatory actions of GnRH on gran¬ ulosa cells may activate certain physiological mechanisms since GnRH agonists cause meiotic maturation of follicleenclosed oocytes in vitro (417) and in vivo in estrogen- or PMSG-primed, hypophysectomized rats (418,419), as well as ovulation in hypophysectomized rats (420,421). Tertiary atretic follicles seem to be most susceptible to the meiosisinducing action of GnRH (418). Degenerative changes in oocytes and premature luteinization of granulosa cells, oc¬ curring without ovulation (384,400), have been reported as a result of GnRH agonist treatment, suggesting that GnRH both stimulates and disturbs normal regulatory mechanisms. A more recent study indicates that GnRH agonist-induced oocyte maturation in the rat is not abnormal (418), in con¬ trast to earlier reports. GnRH also has inhibitory effects on ovarian interstitial cells prepared from hypophysectomized immature rats (422). LH-induced differentiation of interstitial cells is blocked by GNRH, and inhibition of androgen production is by selec¬ tive inhibition of the 17ot-hydroxylase:C-17,20-lyase en¬ zyme (423). The presence of specific receptors for GnRH in ovarian cell types in the rat is reason enough to suspect a physio¬ logical role involving locally produced GnRH factors. Yet it is apparent that GnRH decapeptide produced by the hy¬ pothalamus is unlikely to have a peripheral action on the ovary by virtue of its extremely low concentrations; the highest concentration in human plasma, collected during the periovulatory period, is on the order of 8 x 10“12 M (424). The first suggestion of a local source of GnRH-like factor(s), so-called gonadocrinins, was the report of an acid-extract¬ able factor in rat follicular fluid and in rat granulosa cell

/

353

conditioned-medium that was capable of releasing FSH and LH from rat pituitary cells in vitro but that was immunologically and chromatographically distinct from GnRH (425). However, the investigators who first reported this factor have been unable to reproduce their findings (426). Other investigators have since found two components in extracts from luteinized rat ovaries that resemble GnRH in a radio¬ receptor assay using ovarian plasma membranes but that show little immunoassayable activity (427). These ovarian GnRH-like factors are protease sensitive and have molecular weights between 1,000 and 10,000 daltons. They differ from hypothalamic GnRH in that they are more sensitive to heat (inactivated at 50 or 60°C for 5 min), and they are distinct by reverse-phase high-performance liquid chromatography. Since significant quantities of this receptor-active material have also been found in liver and kidney, it would appear that the factors may not be specific ovarian products. The physiological function(s) of ovarian GnRH-like pep¬ tides are unknown. It is conceivable that they are natural GnRH agonists that mediate paracrine or autocrine regu¬ lation within the ovary, the biological actions perhaps being similar to those already outlined for GnRH and its synthetic agonists. From this perspective, the direct antigonadotropic actions of GnRH shown in the rat might suggest a role in follicular atresia, perhaps related to selection of the domi¬ nant follicle. Alternatively, GnRH-like peptides might be GnRH antagonists. However, evidence obtained in estro¬ gen-primed, hypophysectomized rats indicates that the ac¬ tion of FSH on follicle development is potentiated by a synthetic GnRH antagonist but is inhibited by exogenous GnRH (428). This result suggests that the antagonist was competitive to a GnRH-like factor, presumably of ovarian origin, that inhibits the action of FSH on follicle recruitment to the preovulatory stage. Glucocorticoids The influence of the adrenal glands and glucocorticoid treatments on ovarian function has long been recognized from various studies in mice (429) and rats (430-433) and in women (434,435). Although many of the effects of glu¬ cocorticoids may be mediated by actions on the hypothal¬ amus and/or pituitary gland (431^133,436), a direct action of these steroids on the ovary cannot be discounted. Glucocorticoid receptors have been characterized in rat ovaries (437) and localized to granulosa cells (438). Cortisol or dexamethasone have been shown to inhibit FSH-stimu¬ lated aromatase activity (439,440) but to enhance FSHstimulated progesterone production in cultured rat granulosa cells (439,441). The action of glucocorticoids on proges¬ terone production is associated with increased A5-3(3-hydroxysteroid dehydrogenase and reduced 20a-hydroxysteroid dehydrogenase activities, resulting in enhanced synthesis and concomitant suppression of metabolism. In porcine granulosa cell incubations, cortisol has no effect on basal progesterone production but is stimulatory in the presence

354

/

Chapter 10

of insulin (442). The physiological significance of these observations is obscure. Concentrations of glucocorticoids required to elicit responses in vitro were considered phar¬ macological. Nevertheless, these observations are interest¬ ing from a pathological point of view, and the presence of an ovarian glucocorticoid receptor, as well as of cortisol¬ binding globulin concentrated in follicular fluid (443,444), suggests that glucocorticoids may be of some physiological significance in the regulation of follicular function. Growth Factors and Insulin It is now quite clear that several well-characterized “growth factors” have in vitro modulatory effects on granulosa cell differentiation, including effects on steroidogenesis in sev¬ eral species. However, it is uncertain what the concentra¬ tions of these factors are within the follicles at different stages, what the cellular sources are either within the ovary or in extraovarian tissues, and what the regulatory or per¬ missive actions are in vivo. These factors most likely influ¬ ence cytodifferentiation in its many facets, rather than spe¬ cifically affecting the steroidogenic mechanisms. A complete review of this expanding area of investigation is beyond the scope of this chapter, and readers are referred to the reviews available (329,445). We outline below the principal actions of growth factors on steroidogenesis by ovarian cell cultures. EGF has been shown to bind to high-affinity, low-ca¬ pacity receptors (446) and to have different influences on the steroidogenic pathways for estrogen and progesterone production in rat granulosa cells. EGF inhibits FSH-stimulated estrogen biosynthesis (i.e., aromatase activity) in granulosa cells from immature hypophysectomized, estro¬ gen-treated rats (446,447), while it augments FSH-stimulated progestin biosynthesis and independently stimulates production of pregnenolone, progesterone, and 20a-dihydroprogesterone (446). EGF has also been reported to impair FSH-stimulated progesterone production in rat granulosa cells by increasing catabolism to 20a-dihydroprogesterone (448) . The stimulatory actions of EGF (and perhaps its inhibitory actions as well) may involve sites distal to cAMP production since EGF has been reported to decrease FSH-stimulated cAMP production and to increase cAMP catabolism (448). In studies with pig granulosa cells, EGF inhibited FSH-stimulated estrogen biosynthesis but consis¬ tently inhibited FSH or hCG-stimulated progesterone pro¬ duction only after longer periods of treatment in culture (449) . In contrast with these actions on rat and pig cells, EGF acts on cultured hen granulosa cells to inhibit LHstimulated progesterone biosynthesis, apparently by inhib¬ itory actions at the levels of cAMP production, as well as distal to this step but before C27 side-chain cleavage (450). Other studies in the rat have shown that the EGF receptor content of granulosa cells is regulated by LH and FSH and is maximum at proestrus of the cycle (396,446), indicating that EGF action may be coordinated with other hormonally regulated events. EGF has also been shown to inhibit an¬

drogen production by rat ovarian theca-interstitial cells (451). The only in vivo effect of EGF on the follicle so far dem¬ onstrated is tlje retardation of development of early stage follicles when administered to neonatal mice (452). How¬ ever, there is no evidence to relate this effect to inhibition of estrogen biosynthesis, and it could be due to inhibitory effects on the pituitary. The source of EGF would appear to be extraovarian, and it has been found in several tissues, as well as in mouse plasma and milk (453). EGF has been purified from sub¬ maxillary glands of mice and rats (454,455) and from human urine (456); the structures of the material isolated from these sources are similar but not identical. The fact that androgens greatly stimulate EGF levels in submaxillary glands and plasma (453,457) suggests a possible mechanism for self¬ regulation of androgen production via EGF action on the theca-interstitial cells. The possibility that EGF-like peptfdes might be produced locally and have autocrine actions affecting follicular function must also be considered. The EGF-like growth factor, a transforming growth factor (TGFa), is known to interact with EGF receptors and has been found in several normal tissues (458^460) Also, normal bovine pituitary cells in culture have been reported to pro¬ duce EGF, EGF-like, and TGF-a-like peptides (461). Platelet-derived growth factor (PDGF) also increases FSHstimulated progestin production, which may involve in¬ creased sensitivity to FSH and cAMP (448). This in vitro action suggests that PDGF release from platelets during follicular rupture may act to stimulate luteal cell production of progesterone. Several modulatory actions on steroidogenesis have also been shown for insulin and the insulin-like growth factors (IGFs); these latter factors, classically thought to be of he¬ patic origin, are known as IGF-I [or somatomedin C (SmC)] and IGF-II in humans [or rIGF-II/MSA (multiplication stimulating activity) in the rat]. Receptors for SM-C/IGF-I have been reported in granulosa cells of immature pigs (462,463) and of rat (445). Insulin receptors have also been reported on granulosa or luteal cells of rat, pig, and human (464^468). It is not clear whether the effects of insulin on granulosa cells are mediated through insulin receptors or type 1 IGF receptors (which preferentially bind IGF-I but also bind IGF-II and insulin). However, IGF-I does not appear to interact with insulin receptors. There is evidence in the rat (445) and pig (469,470) for IGF-I production by granulosa cells in culture and for mea¬ surable quantities of IGF-I in follicular fluid from porcine (471,472) and human ovaries (445). Moreover, immunoreactive IGF-I in ovarian extracts increases following treat¬ ment of hypophysectomized, estrogen-treated rats with growth hormone (GH) (473), suggesting regulation of ovarian IGF production. Insulin has also been measured in fluid from porcine (472) and human (474) follicles. The effects of IGF-I on rat granulosa cells cultured under serum-free conditions are to augment the stimulation by FSH of progesterone and 20a-dihydroprogesterone production

Follicular Steroidogenesis and Its Control / (475,476) and of aromatase activity (477). In porcine gran¬ ulosa cells, IGF-I alone or synergistically with FSH in¬ creases progesterone production (462) and also appears to stimulate pregnenolone, progesterone, and 20a-dihydroprogesterone independently, without synergism with FSH (463). Independent stimulatory effects of rIGF-II/MSA have been shown on progesterone production by porcine granulosa cells when cultured in the absence of serum (478), although another study has shown only a synergism with FSH (462). Insulin has also been shown to act synergistically amplifying the stimulatory effect of LDL on progesterone biosynthesis by porcine granulosa cells, the mechanisms involving in¬ creased binding, internalization, and degradation of LDL (479). GH has been shown to augment FSH-stimulated proges¬ terone and 20a-dihydroprogesterone production by cultured granulosa cells from hypophysectomized, estrogen-treated rats by mechanisms involving increased cAMP, as well as stimulation at a site distal to cAMP (480). It has not yet been determined whether this is a direct effect of GH or whether GH-stimulated production of IGF is responsible. Insulin has steroidogenic effects similar to those of IGFI, and since effects of insulin and IGF-I are not additive, it is likely that they employ the same mechanism. Progester¬ one production by rat (464,481) and pig (442,466,467,478, 482,483) granulosa cells is stimulated by insulin, as is FSHstimulated aromatase activity in rat (481) and human (484) granulosa cells. In contrast, insulin inhibits or is without effect on aromatase activity in porcine granulosa cells (478,485).

Neuroregulatory Substances and Ovarian Innervation The innervation of the mammalian ovary has been well documented. Although considerable variation exists among species, sympathetic nerves, to a large extent accompanying blood vessels, innervate the ovarian interstitial tissue and perifollicular regions in several species, with follicles at all stages of development having adrenergic nerve terminals in close proximity to the blood vessels of the theca externa. Neither blood vessels nor nerves penetrate the basement membrane to reach the granulosa layers of follicles at any stage of development (486). Although the physiological significance of the ovarian nerves remains to be established, considerable research has centered around the possible role of adrenergic neurotrans¬ mitters in controlling follicular function. Studies by Bahr and Ben-Jonathan (487) have shown that stimulation of the rat ovary with gonadotropin depletes the ovary of the cate¬ cholamine. A 40% reduction of ovarian concentrations of noradrenaline occurred 12 hr after PMSG administration to prepubertal rats; this was followed by a further 40% reduc¬ tion 4 hr after the preovulatory LH surge. FSH, rather than LH or PRL, was found to be the pituitary hormone primarily involved in depletion of noradrenaline from Graafian fol¬

355

licles (488). Noradrenaline levels in porcine follicular fluid have been reported to vary with the stage of the estrous cycle, the highest levels being observed during the follicular phase (days 16-20)(489). In vivo evidence for possible involvement of ovarian nerves in regulation of steroid secretion has come from ovarian denervation and ovarian nerve stimulation experiments. De¬ creased activity of A5-3(3-hydroxysteroid dehydrogenase was seen in both the interstitial gland cells and corpus luteum of the pregnant rat following ovarian denervation (490). Subsequently, Capps et al. (491) showed that electrical stim¬ ulation of the nerves in the ovarian plexus of hypophysec¬ tomized rats caused the interstitial cells to hypertrophy and develop ultrastructural features typical of active steroid-se¬ creting cells. The possible role of catecholamines in the direct regu¬ lation of steroid biosynthesis by follicle cells has been ex¬ amined both in vivo and in vitro. In vivo experiments in¬ volved intrafollicular injection of adrenergic agonists and antagonists in rabbits. Beta-adrenergic, but not a-adrenergic, agonists resulted in increased progesterone output by the ovary, without influencing estrogen secretion (492). In vitro studies have shown catecholamine stimulation of pro¬ gesterone production by dispersed luteal or granulosa cells in culture (492^-96). These effects could be blocked by the (3-adrenergic antagonist propranolol. The steroidogenic re¬ sponse (progesterone synthesis) of cultured rat granulosa cells to noradrenaline and isoproterenol was markedly en¬ hanced by pretreatment of the cells with FSH in vivo or in vitro (496). Perhaps related to this is the observation of increased levels of high-affinity (32-adrenergic receptors on granulosa cells during the proestrus associated with puberty in the rat (497). Catecholamine responsiveness of granulosa cells in preovulatory follicles of the rat apparently develops only after the preovulatory LH surge (498). The interstitial tissue, the most richly innervated com¬ ponent of the ovaries, also contains p2-adrenergic receptors (497), and addition of the |32-agonist zinterol to cultured rat interstitial tissue (“residual” tissue, after removal of most granulosa cells) increased output of testosterone and androstenedione from this tissue on the day of proestrus, the peripubertal stage in which the greatest concentration of p2receptors were present (497). Addition of epinephrine, nor¬ epinephrine and isoproterenol to cultured theca-interstitial cells from hypophysectomized rats also markedly enhanced the secretion of certain androgens in the presence of hCG stimulation. The catecholamines were ineffective in stim¬ ulating basal (i.e., in the absence of hCG stimulation) se¬ cretion of steroids by the theca-interstitial tissue (despite the fact that they are effective, by themselves, in stimulating cAMP output) (499). Other neurotransmitters that have been detected in the ovary include acetylcholine (based on dem¬ onstration of acetylcholinesterase activity) (500), dopamine (487,501), substance P (502,503), VIP (504), and y-aminobutyric acid (GABA) (329). Although high-affinity bind¬ ing sites in ovarian tissue have been reported for some of

356

/

Chapter 10

these compounds and certain in vitro effects on steroido¬ genesis occur (505), suggesting possible regulatory roles, information is lacking about the specific effects of these neurotransmitters on steroid biosynthesis in the follicle.

ulogenesis, the process whereby germ cells associate with the follicular cells to form primordial follicles. This pre¬ follicular phase can occur in embryonic life, in the neonate, or even in later prepubertal stages depending on the species.

STEROIDOGENESIS AND ITS CONTROL BY FOLLICLE TYPE

The Prefollicular Ovary

Introductory Description of Follicle Types In the foregoing sections we have described the charac¬ teristic steroidogenic functions and control of the separate cell types of the mature ovarian follicle. The purpose of this section is to reexamine the evidence from the perspective of follicular development stage, drawing on related exam¬ ples from human and animal studies to illustrate principles. It is intended to place the previous discussion of cellular details of steroid formation and regulation into the context of the follicle as a developing biosynthetic structure. We shall consider the dynamic processes of steroid synthesis and secretion occurring within the follicles as follicular mat¬ uration proceeds under the control of the pituitary gonad¬ otropic hormones. Although the follicular contribution to ovarian steroid production is at any time the composite secretion of steroids by follicles at various functionally dif¬ ferent stages, the most important contribution eventually comes from one or several dominant follicles forming a cohort destined to ovulate. The stages of follicular development have been described by several classifications that distinguish follicles in terms of oocyte morphology and size, and the number of sup¬ porting granulosa cells and their organization. Similar schemes for mouse (506) and human (507) ovaries have defined eight principal types of follicle. The general categories of follicle types are the following: small nongrowing follicles (i.e., primordial follicles classified as types 1-2 in human and 13a in mouse); preantral follicles (i.e., single-layered primary and multilaminar secondary follicles classified as types 35 in the human and types 3b-5b in the mouse), which are characterized mainly by increases in oocyte size and number of granulosa cells; and antral follicles (i.e., tertiary or Graa¬ fian follicles classified as types 6-8 in human and mouse), which feature formation of a fluid-filled antrum and further increases in granulosa cell number. In addition, antral fol¬ licles of several species have been referred to previously as either small or large to indicate the extent of their expansion, so this description is also used here. Moreover, large antral follicles are described as either nonovulatory or preovulatory to indicate their maturity according to accepted functional criteria. We shall discuss the differing steroidogenic func¬ tions of follicles according to several of these morphological categories. Initially, a prefollicular phase will be considered. This phase does not represent a follicular stage but, rather, a brief period in ovarian organogenesis before the onset of follic-

The prefollicular phase of ovarian development would appear to be largely devoid of steroidogenic function in some, but not all, species. However, this view may reflect the great difficulty in assessing steroid production at early stages in certain smaller animals. Steroid synthesis in rodent ovaries is undetectable or very limited before follicles form (70,508). The rat ovary, which begins forming follicles on postnatal day 1 (509,510), was shown to be unresponsive to gonadotropic stimulation of ^estrogen production during the fetal period (511). Histochemical activity of A5-3(3-hydroxysteroid dehydrogenase has been demonstrated in prefollicular ovaries of rat (512) but only in the interstitial cells, with no activity apparent in granulosa cells. In neonatal mouse ovaries, in which folliculogenesis begins on postpartum day 2 (513-515), sig¬ nificant histochemical activity for A5-3 (3-hydroxysteroid de¬ hydrogenase first appears in the intraovarian rete cells at 7 days of age (70). However, in all these studies, the presence of histochemical enzyme activity does not establish that the enzyme is functioning as part of a biosynthetic pathway. In contrast to studies in rodents, prefollicular ovaries of rabbit (516), sheep (517), and cow (518) apparently syn¬ thesize or have certain functional enzymes required to syn¬ thesize steroids even before initiation of meiosis in the oo¬ cytes, which precedes follicle formation. Studies in the rabbit demonstrated conversion of testosterone to estradiol by ova¬ ries at day 18 of gestation, but the presence of endogenous androgen substrate was not determined (516). Other studies showed that gonadotropins were able to stimulate proges¬ terone production by cultured rabbit granulosa cells, isolated from ovaries at the early postnatal period, but only after follicles had formed (519); folliculogenesis begins in this species at 14 days of age (520—522). In bovine ovaries, in which folliculogenesis begins about day 95 of fetal life (523), estradiol production has been detected as early as 45 ± 3 days of fetal age but was biphasic and subsequently declined (518,524). The onset of ovarian estradiol produc¬ tion occurred when the sex of the embryonic gonad was first distinguishable. De novo synthesis is believed to occur at this early fetal age since both testosterone and estradiol have been shown to be present in the ovaries. Furthermore, estradiol production in vitro is stimulated by LH. Proges¬ terone and prostaglandins are produced even earlier, at day 30 of gestation (518). At much later gestational stages in the bovine, at least a month before term, ovaries in vitro convert [14C]progesterone to several hydroxylated metab¬ olites of C2i-steroids and to Cig-steroids, as well as convert [ 14C]androstenedione to testosterone and estradiol (525).

Follicular Steroidogenesis and Its Control / These conversions most probably involve enzymes present in both follicular and interstitial cells. Steroidogenic function in prefollicular human ovaries has not been adequately investigated, but histochemical enzyme activity for A5-3 (3-hydroxy steroid dehydrogenase has been demonstrated at this stage in the interstitial cells (526-528). Evidence for early steroidogenic function in human fetal ovaries following folliculogenesis is considered later (see section on preantral and early antral follicles, below). Morphological evidence suggests biosynthetic and secre¬ tory activity in granulosa cells in prefollicular ovaries of various species. In general, these cells have numerous ri¬ bosomes and branching mitochondria and well-developed Golgi complexes, and they often contain lipid droplets. However, these features are not specific indicators of ste¬ roid-synthesizing cells, and they may reflect other biosyn¬ thetic functions associated with granulosa cell-oocyte co¬ operation. In the prefollicular ovary, oocytes and granulosa cells exist together within the irregular cords and nests of the ovarian cortex. The arrangement of oocytes and gran¬ ulosa cells is closely packed, with extremely close appo¬ sition of plasma membranes, since at this time there is no zona pellucida to separate the granulosa cells from the oo¬ cyte. At this stage, therefore, granulosa cells may serve primarily to provide nutrients to the oocyte, a function that is certainly continued, in the developing follicles of adults, by the specialized granulosa cells of the corona radiata (60). Granulosa cells in prefollicular ovaries also appear to be active in the phagocytosis of degenerating germ cells. Oocytes and granulosa cells are entirely interdependent in the formation of follicles, without which neither sub¬ stantial steroidogenesis nor oocyte growth can occur. There¬ fore, a most important aspect of the prefollicular phase is the establishment of granulosa cell and oocyte associations leading to folliculogenesis. When the number of oocytes in the fetal ovary is greatly decreased by a genetic defect or destructive treatments, follicles rarely form, and the ovary is deficient in normal steroid production. An example is Turner’s syndrome in humans, where the 45,X chromosome constitution results in a failure to differentiate oocytes and follicle organization is disrupted (529-531). The XO con¬ dition in mice is far less severe, and females are fertile and appear normal with the exceptions of reduced numbers of oocytes, a shorter reproductive life span (532), and devel¬ opmental retardation (533). Destruction of oocytes in ova¬ ries of fetal rats by treatment with the antimitotic drug busulphan (534,535) or irradiation (536) leads to a similar conclusion; Without oocytes, follicles do not form, and the resultant ovary consists mainly of stroma containing net¬ works of cords and tubules resembling the intraovarian rete ovarii. Such ovaries lack steroidogenic functions and fail to develop responsiveness to gonadotropins (537). In summary, the cells of the prefollicular phase in some, but not all, species appear to be steroidogenically inactive and to be unresponsive to gonadotropins in terms of steroid production. However, it is necessary to interpret the sig¬

357

nificance of these findings cautiously since apparent inac¬ tivity may reflect inadequate sensitivity of the assay pro¬ cedures, while apparent activity of certain enzyme steps need not necessarily indicate biologically important de novo production of steroid hormones. Early differentiation of granulosa cells and their organization into follicles would appear to be influenced largely by interactions of these cells with oocytes and gonadotropins, but little is known about other local factors controlling early ovarian differentiation and folliculogenesis, including steroidogenesis.

Preantral and Early Antral Follicles Preantral follicles develop from primary follicles by en¬ largement of the oocyte and proliferation of the supporting granulosa cells. The factors responsible for initiation of growth of follicles from the nongrowing pool are unknown but have previously been discussed in terms of regulation by the oocyte or granulosa cells (538). Subsequent devel¬ opment of preantral follicles is not dependent on gonado¬ tropins and continues after hypophysectomy (539). As the follicle enlarges, theca cells differentiate from cells within the ovarian stroma, thereby forming a sheath of flattened cells around the follicular basement membrane. Formation of the theca layer is quite variable among individual follicles in the mouse but becomes distinct at the multilaminar stage. In the hamster, theca appears to differentiate in follicles with seven or eight layers of granulosa cells (540). In the mouse, only later, when the antrum forms, does the theca differentiate further into theca interna and externa (541,542). Evidence of various types, largely indirect, suggests that in several species limited steroidogenic activity is probably acquired at the preantral stage of follicle development, al¬ though it is weakly expressed until the antral stage. FSH receptors are found on rat granulosa cells at all follicular stages, including early preantral follicles (543,544). How¬ ever, the precise contributions of preantral follicles to ovar¬ ian steroid production are not well understood, primarily because of difficulties in studying the relatively low secre¬ tory activities of these follicles in vivo and because meth¬ odology for isolating preantral follicles is not well devel¬ oped. Only recently has an improved enzymatic and mechanical method been developed for the isolation of var¬ ious stages of intact preantral follicles in the hamster (540). Research on early follicles has also been neglected in favor of the functionally more important Graafian follicle. Several different approaches for deducing the steroidogenic function of preantral follicles are discussed below. The formation of preantral follicles in the neonatal period of the rat provides an opportunity to examine gonadotropin responsiveness and steroidogenesis in ovaries containing only small follicles, early preantral follicles, and interstitial cells. Since theca cells from the immature rat produce neg¬ ligible amounts of estrogens (190) and rat ovarian interstitial cell preparations in culture produce only small quantities of

358

/ Chapter 10

estrogens (86), neonatal ovarian synthesis of estrogens in vitro may be an indicator of aromatase activity in the gran¬ ulosa cells. However, it is also possible that neonatal thecal and interstitial cells at this age differ from those in older rats and are capable of significant estrogen biosynthesis. Follicle growth in the neonatal rat begins by postpartum day 2, with multilaminar follicles forming by day 6, but it is not until about day 10 that the first follicles containing antra appear in significant numbers. The available data sug¬ gest that preantral follicles of the neonatal period may re¬ spond to gonadotropins and produce steroids. Quattropani and Weisz (545) have shown that A5-3(3-hydroxysteroid de¬ hydrogenase activity is present histochemically in interstitial cells of day 4 rat ovaries and that as early as day 5, ovaries are capable of converting [3H]progesterone to estrogens. These investigators suggested that the interstitial cells were solely responsible for these steroidogenic activities in the neonatal rat. However, FSH binding sites, which are thought to be specific for granulosa cells, have also been demon¬ strated in the ovary as early as day 5 (546), and FSH has been found to stimulate both cAMP and steroid production by ovaries of 4-day-old rats (547). Other studies with cultured ovaries from 4-day-old rats have shown that progesterone production is stimulated by (Bu)2cAMP and conversion of testosterone to estradiol is stimulated by (Bu)2cAMP or FSH (548). Furthermore, es¬ trogen production in these studies was not substantially stim¬ ulated by FSH alone, but was greatly augmented by FSH in combination with LH (548), which is consistent with a “two-cell type, two-gonadotropin” form of regulation at this early developmental stage. Recent studies involving similar in vitro incubations of rat ovaries have established that purified FSH, LH, or PGE2 will each stimulate, to a different extent, the production of progesterone, androstenedione, and estradiol by ovaries at day 6 but not earlier (547). In these studies, secondary preantral follicles were the most advanced stages present at this time, suggesting that gonadotropin-regulated de novo synthesis of the three major classes of steroids might be produced by the developing preantral follicles in cooperation with the interstitial cells. In these studies, LH was by far the most potent stimulus for androstenedione and estradiol production, but addition of the phosphodiesterase inhibitor 3-isobutyl- 1-methylxanthine (MIX) resulted in enhanced re¬ sponsiveness to both FSH and LH, with FSH then having a potency equal to that of LH. The contention that phos¬ phodiesterase activity limits steroidogenic responsiveness to gonadotropins in early ovaries was confirmed in incubations of 4-day-old ovaries, which only in the presence of MIX allowed increases in androstenedione secretion in response to PGE2, LH, and FSH and increases in estradiol in response to PGE2 alone (547). These results indicate that steroido¬ genic capabilities differentiate in neonatal rat ovaries even before acquisition of full responsiveness to gonadotropins, and certainly well before formation of antral follicles. The responsiveness to LH might also be regulated by local ste¬

roid action since other investigators have shown that estro¬ gen treatment of neonatal rats advances the age at which LH-stimulated cAMP can be demonstrated in vitro (549). The extent to which steroidogenic activities seen in neo¬ natal rat ovaries might be influenced by the high circulating levels of FSH (550,551) and other pituitary hormones (552) present during this period is uncertain. It can only be spec¬ ulated that the steroidogenic pathways seen in neonatal rat ovaries might be similarly expressed as a result of the co¬ operation of preantral follicles and interstitial cells present in ovaries at later developmental ages, including adults. Steroids secreted by preantral follicles at all developmental ages might influence local concentrations within the ovary, thereby affecting follicular function or maturation. How¬ ever, the studies described above with whole neonatal ovary incubations do not provide conclusive evidence that signif¬ icant steroidogenesis occurs within the follicular compart¬ ment, as opposed to the interstitial cells. Similarly, there is no direct information on the steroi¬ dogenic function of preantral follicles in the adult human ovary. However, studies have investigated steroidogenic activity of fetal ovarian tissue in vitro. Most of the studies have obtained ovarian tissue at fetal ages later than the beginning of folliculogenesis, which occurs at about 8 weeks of gestation in the human (553). Therefore, steroidogenic activities reported might partly reflect functions of early follicles. However, another potentially important steroido¬ genic cell population is found in the interstitium of ovaries from human fetuses at 12 to 20 weeks gestation (554). These interstitial cells of the embryonic ovary, also called primary interstitial cells (97), are located in the medullary region beneath the cortical cords and are often adjacent to blood vessels. Their fine structure is typical of steroidogenic cells, and they are relatively large (>30 p,m greatest diameter). The evidence for steroidogenesis in human fetal ovaries is based on experiments in which fetal ovarian tissues were incubated in the presence of radioactive steroid precursors. The ability to convert [3H]testosterone or androstenedione into estradiol and estrone was not apparent in undifferen¬ tiated gonads of human fetuses at approximately 6 to 8 weeks gestation but was acquired at about 8 to 10 weeks and continued to be present until at least midgestation (555). However, George and Wilson cautioned that their results alone do not give evidence of de novo estrogen synthesis from endogenous substrates. Earlier studies with ovaries from more advanced fetuses (16-22 weeks gestation) did not find evidence of C19-steroid synthesis from [14C]progesterone (556) or from [1-14C]acetate (557), but in the latter study acetate was converted to lanosterol, cho¬ lesterol, and smaller quantities of pregnenolone and pro¬ gesterone. A further study in which fetal ovarian tissue was incubated with [14C]pregnenolone found progesterone to be the main product, but products of the 5-ene-3(3-hydroxysteroid pathway, 17a-hydroxypregnenolone and dehydroepiandrosterone, were also identified (558). An earlier study demonstrated the active conversion of [14C]progesterone to

Follicular Steroidogenesis and Its Control / 20a-dihydroprogesterone in fetal ovaries at 19 weeks but not at 12 to 15 weeks of gestation (559). More recently, in organ cultures of ovaries from three fetuses (12, 20, and 22 weeks gestation), release of progesterone, dehydroepiandrosterone, androstenedione, estrone, and estradiol (560) was reported. It was also found that progesterone production in vitro was stimulated by (Bu)2cAMP and, to a lesser ex¬ tent, by LH/FSH. A most interesting finding is the substantial conversion by fetal human ovaries (at approximately 12.5-17.5 weeks gestation) of [3H]pregnenolone sulfate into pregnenolone, 17a-hydroxypregnenolone, dehydroepiandrosterone, and androstenedione but without formation of progesterone, tes¬ tosterone, or estrogens (561). This indicates the presence of steroid sulfatase activity in the fetal ovary and the ability to convert C2i-steroids to Ci9-steroids via the 5-ene-3(3hydroxysteroid pathway, similar to the preferred pathway to androstenedione in the adult ovary (84). The presence of a high concentration of pregnenolone sulfate in fetal blood (562) suggests there is a natural substrate available. These studies indicate that the human fetal ovary is steroidogenically active at developmental stages after initiation of folliculogenesis but when embryonic interstitial cells are also present. It is possible that both early follicular cells and interstitial cells contribute to the conversions observed. However, the presence of the interstitial cells precludes deductions regarding the steroidogenic functions of early stages of human follicles. As mentioned previously, inter¬ stitial cells in the human ovary are histochemically reactive for A5-3(3-hydroxysteroid dehydrogenase even in prefollicular stages. One study with fetal ovaries of rhesus monkeys indicates that the capacity for de novo production of estradiol is ac¬ quired only in late gestation, when multilayered and antral follicles have already developed (563); this suggests that preantral follicles may not contribute significantly to estro¬ gen production in this species, at least during fetal life. A more direct, but little used approach to assess steroido¬ genesis in preantral follicles at various stages of develop¬ ment has been to obtain dispersed follicles by enzymatic and/or mechanical procedures. In one study, dispersed ovar¬ ian tissues from immature mice were cultured in vivo as transplants, either implanted subcutaneously in gelfoam sponges or diffusion chambers or as cells implanted into intraocular sites (564). Intraocular transplants were accom¬ panied by a fragment of vaginal epithelium as a biological indicator of estrogen production. These studies demon¬ strated that subcutaneous transplants into ovariectomized mice caused vaginal opening and onset of estrous cycles similar to that seen in nonovariectomized controls, and in¬ traocular transplants promoted comification of the vaginal fragment. In the latter situation, follicles transplanted to¬ gether with dispersed ovarian cells, presumably containing androgen-secreting interstitial cells, were far more effective in causing vaginal comification. Most importantly, it was noted that vaginal comification, taken as a measure of es¬

359

trogen production, was apparent only when growth of fol¬ licles and concomitant maturation of antra were observed. Therefore, while preantral follicles might have contributed to estrogen production by transplants of dispersed ovarian tissues, these studies suggest that substantial secretion of estrogens requires antral follicles. Enzymatic dispersion of the ovarian compartments has also been used successfully to harvest all but the most mature antral follicles from rabbit ovaries (565). Measurements of follicular steroid content, which undoubtedly reflect steroid uptake from other compartments and follicles at different stages, as well as biosynthesis, suggest a progressive in¬ crease in levels of progesterone and estrogen as follicles develop from small primary to large antral stages. However, since de novo synthesis was not conclusively demonstrated, these results do not necessarily establish the steroidogenic abilities of early rabbit follicles. Preantral follicles have been isolated from ovaries of cyclic hamsters and incubated in vitro to assess LH-stimulated steroid production (566). These studies indicate that isolated preantral follicles show a shift in LH-stimulated steroid products, depending on the time of follicle isolation relative to the endogenous LH surge. Before the LH surge during proestms, androstenedione production predominated; im¬ mediately following the endogenous LH surge, the isolated preantral follicles produced only progesterone. The ability to produce androstenedione, and additionally estradiol, reappeared in follicles isolated on the afternoon of estrus. Subsequently, isolated small antral follicles, which had de¬ veloped from preantral follicles present at estrus, produced mainly estradiol and androstenedione, with only smaller amounts of progesterone. It is apparent that steroidogenesis can occur in isolated preantral hamster follicles and that in vivo the LH surge may act to synchronize steroidogenesis by these follicles (566). Many in vivo studies with immature normal or hypophysectomized rats indicate that ovaries consisting primarily of larger preantral follicles are steroidogenically responsive to gonadotropins (FSH and/or LH). However, these animal models do not allow the action of FSH in inducing steroid¬ ogenic activity to be distinguished from the concurrent effect of FSH on formation of follicular antra. Large preantral follicles that form in hypophysectomized rats do not show autonomous synthesis of estrogens but re¬ quire stimulation with FSH and either LH or aromatizable androgen (209). It would appear that significant aromatase activity occurs only as antral follicles form in response to FSH stimulation. Several models exist for the culture of granulosa cells from ovaries containing primarily small and preantral fol¬ licles. Granulosa cells can be obtained either from normal immature rats before large antral follicles have formed or from immature hypophysectomized rats, in which very few small tertiary follicles will form in the absence of gonad¬ otropin stimulation. In both models, treatment with estrogen is often used to stimulate the numbers of relatively uniform

360

/

Chapter 10

preantral follicles; in addition, this treatment greatly in¬ creases the responsiveness of the cells to FSH. Regardless of the model employed, granulosa cells isolated from ovaries of these animals show minimal production of progestins or estrogens when cultured without gonadotropins, even in the presence of aromatizable substrates. Other features of gran¬ ulosa cytodifferentiation, such as LH receptor formation, are also absent in cells cultured without gonadotropins, in¬ dicating that these cells are minimally differentiated. There¬ fore, it is apparent that key steroidogenic enzymes are lack¬ ing or weakly expressed in granulosa cells of preantral follicles in hypophysectomized or normal immature rats. Since preantral follicles can, and probably do, develop without gonadotropin support, it is likely that the granulosa cells of at least early preantral stages are not very active steroid producers. However, it is probable that cyclical increases in circulating gonadotropins act in a coordinate fashion to stimulate cytodifferentiation and steroidogenesis in granu¬ losa cells of late preantral follicles. Granulosa cells from preantral follicles of rats respond in culture to FSH with increased secretion of C2i-steroids and increased aromatase activity. These steroidogenic effects in vitro usually appear after a delay of about 24 hr, suggesting the need for in¬ duction of steroidogenic enzymes. Certain steroidogenic enzymes are present (although per¬ haps not maximally expressed) in preantral follicles that have not been stimulated with gonadotropins. Granulosa cells isolated from immature rats and placed in culture are able to convert exogenous pregnenolone to progesterone, 20a-dihydroprogesterone, and, probably in lesser amounts, 5a-reduced C21-steroid metabolites (47). They also convert testosterone rapidly to its 5a-reduced metabolites, including 5a-androstane-3a,17(3-diol and androsterone (141). Histochemical demonstration of A5-3p-hydroxysteroid dehy¬ drogenase activity in the rat ovary indicates that this enzyme is strongly expressed in granulosa cells of preovulatory fol¬ licles but is only weakly expressed in earlier antral stages (567). This is consistent with the biochemical evidence that induction of this enzyme requires FSH stimulation of the granulosa cells (141). A variety of factors might be responsible for limiting the steroidogenic competency of preantral follicles. Although the granulosa cells have FSH receptors that, at least in late preantral follicles, are functionally coupled to adenylate cy¬ clase, these receptors do not function optimally since es¬ trogen is required to augment FSH-stimulated cAMP pro¬ duction. This increased cAMP response in turn allows FSH to increase aromatase activity and estrogen synthesis and to increase the number of gonadotropin receptors and cAMP binding sites (568). These changes are associated with de¬ velopment of the antral follicle. Further, although it is gen¬ erally assumed that circulating FSH and LH are similarly available to the granulosa cells of preantral and later-stage follicles by diffusion from capillaries of the theca layer through the follicular wall and into the granulosa cell layers, this assumption has not been critically tested. However,

molecular size per se is not a limiting factor for gonadotropin penetration of the follicular membrane (569). The ability of healthy and steroidogenically active follicles to concentrate gonadotropins in follicular fluid suggests the probable im¬ portance of the availability of FSH to steroid production. Thus, while it is clear that follicle cells isolated from large preantral follicles respond to gonadotropins and require them for steroid production, it is uncertain to what extent preantral follicles from various species secrete steroids in vivo in response to normal cyclical fluctuations in circulating go¬ nadotropin levels.

Antral to Early Preovulatory Follicles There is general agreement that the antral stages of fol¬ licular development, and especially the early preovulatory follicles, are by far the most important source of ovarian steroids during the female cycle. This changing pattern of steroid secretion is associated with increased follicle cell growth and cytodifferentiation and considerable expan¬ sion of the antrum, which occur in response to changing levels of FSH and LH. The ability to secrete significant quantities of estrogens, a function acquired as antral follicles form, is of key importance in increasing the responsiveness of the granulosa cells to FSH and subsequently to LH (570572). The direct action of estrogen at the follicular level and the additional indirect effect of feedback inhibition of gonadotropin secretion are important contributing factors in follicular selection. The formation of the fluid-filled antrum provides a con¬ venient morphological marker of the stage in follicle de¬ velopment when gonadotropin-dependent growth of the fol¬ licle approximately begins and steroid secretion greatly increases. Antrum formation is induced by the action of FSH, but not LH alone, and presumably reflects changes in the FSH-dependent secretory function of the granulosa cells, as well as influx of interstitial fluid (569,573). Increased follicular vascularization and steroidal effects on follicular vascular permeability could also be factors in antral fluid accumulation (569). However, antrum formation is appar¬ ently not a necessary feature of preovulatory follicular mat¬ uration in all species, since in several exceptional families (i.e., Tenrecidae and Erinaceidae), ovaries do not form antral follicles (574). Granulosa cells of early antral follicles do not generally have ultrastructural features characteristic of steroid-secret¬ ing cells. This was originally taken as evidence of their nonsteroidogenic function until later histochemical and bio¬ chemical evidence changed that view. Only after the LH surge are substantial ultrastructural changes seen in organ¬ elles associated with steroidogenesis in the granulosa cells. These changes, which are considered the earliest signs of morphological luteinization and may occur prior to ovula¬ tion in certain species (575), are discussed in the next sec¬ tion.

Follicular Steroidogenesis and Its Control

We have already reviewed the evidence for steroid path¬ ways present in mature granulosa and theca cells of antral follicles and considered the types of cellular cooperation involved in regulating steroid production at this stage. In this section, we consider only the additional evidence in¬ dicating that the major sources of ovarian estrogens and progesterone during the follicular phase are the large antral and preovulatory follicles, and that both small and large antral follicles are significant sources of androgens prior to the LH surge. This evidence comes mainly from measure¬ ments comparing steroid levels in ovarian venous plasma or in follicular fluid with those in peripheral plasma. Steroid measurements in ovarian venous plasma reflect rates of ovarian secretion into the circulation. For example, a close relationship between follicular production of certain steroids and secretion into blood is apparent, since pulsatile release of LH in sheep is followed rapidly by steroid se¬ cretion into the ovarian vein, with peak levels of estradiol and androstenedione occurring 30 min after each pulse (576). In comparison, steroid measurements in follicular fluid pro¬ vide a static profile of the pool of steroids secreted primarily by the granulosa and theca cells of one follicle. However, the concentrations of steroids in follicular fluid reflect not only secretion into the fluid, but also changes due to me¬ tabolism and dilution as fluid accumulates. Transfer of ste¬ roids from follicular fluid to venous plasma occurs only slowly. As demonstrated in the mare, the hourly transfer of radiolabeled pregnenolone and androstenedione from fol¬ licular fluid to ovarian vein was approximately 3% to 9% (189). The export of free and conjugated steroids from fol¬ licular fluid might also be impaired by binding globulins in the fluid (577,578). In this regard, species differences occur since sex hormone-binding globulin is present in blood plasma and follicular fluid of the cow and sheep but not the pig (579).

Estrogens Estrogen secretion in women is highest during the late follicular phase of the menstrual cycle (580). At this time, estrogen levels are significantly higher in ovarian venous plasma from o varies containing a large antral follicle (>8 mm diameter) compared with contralateral ovaries contain¬ ing small antral follicles (1.3 mU/ml), indicating the key regulatory role of FSH (582). Second, the importance of granulosa cells in estrogen production is further indicated by the positive relation between the number of granulosa cells and the intrafollicular estradiol concentration for a given size of antral follicle (588). Thus, it appears that preovulatory follicles prior to the midcycle LH surge are characterized by much higher estrogen biosynthetic activity and estrogen content, higher FSH concentrations, and more granulosa cells than in nonovulatory or atretic follicles (131,584,589,590). Studies of numerous other species—including sheep (591), cows (109), pigs (592), rabbits (593), and rats (594)—report similarly high levels of estrogens in follicular fluid from

362

/ Chapter

10

large antral follicles, with maximal concentrations occurring in the preovulatory period, before or immediately after the LH surge. On the other hand, fluid from atretic follicles of various species, like the human, characteristically exhibits substantially lower estradiol concentrations (109,276, 588,595,596). Over 90% of the circulating estradiol secreted during the late follicular phase of monotocous species originates from the dominant follicle (597,598). In women, normally only one antral follicle matures to preovulatory eminence. Es¬ tradiol levels in the fluid of the dominant follicle before the midcycle LH surge is on the order of 1 to 2 p-g/ml (131,569,597), and similar estrogen levels have been found in fluids from ovaries of cyclic (597,599-602) and gonad¬ otropin-stimulated (603,604) women. Results from various other monotocous species are comparable (187,213,603,605607). Therefore, this evidence indicates that the dominant follicle is the principal source of the preovulatory estrogen surge that appears in plasma. This conclusion has been directly supported by studies in women that measured the large decrease in ovarian venous plasma levels of estradiol and estrone following surgical enucleation (i.e., intact re¬ moval) of the largest follicle (608). This preovulatory es¬ trogen production is apparently regulated by LH-stimulated production of aromatizable androgens in the theca cells (dis¬ cussed below; see also the section on post-LH surge pre¬ ovulatory follicles), in agreement with evidence that gran¬ ulosa aromatase activity is already maximally stimulated in early preovulatory follicles and cannot be further stimulated by FSH in cells from follicles >12 mm in diameter at midfollicular phase (586). Some insight into the physiological factors regulating fol¬ licular estrogen biosynthesis during the preovulatory period has been provided from in vitro incubations of follicles at different stages of the estrous cycle. In the rat, regulation of estradiol production by isolated preovulatory follicles appears to be principally at the level of production of aro¬ matizable androgens. When follicles from ovaries of 5-day cycling rats were incubated individually, accumulation of estradiol in the culture medium increased from low levels at diestrus I to high levels at proestrus (609). Estradiol accumulation by follicles obtained at either diestrus or proes¬ trus was increased by addition of androstenedione or tes¬ tosterone (indicating the presence of aromatase activity) or by addition of 17a-hydroxyprogesterone or progesterone (indicating the presence of 17a-hydroxylase:C-17,20-lyase enzyme required for conversion of progesterone to andro¬ gens). In these studies measurements of follicular enzyme activities at diestrus and proestrus demonstrated little change in lyase activity and a small increase in aromatase activity at proestrus. Since C-17,20-lyase activity was always more limiting than aromatase activity, it was concluded that in¬ creased estradiol production at proestrus might be due to an increase in endogenous production of progesterone, with greater conversion to androgens. This was supported by the finding that pregnenolone accumulation in medium from

follicle incubations, carried out in the presence of cyanoketone to inhibit metabolism to progesterone, increased be¬ tween diestrus and proestrus (609). Other studies (as already discussed in the section on the theca) have indicated that increased thecal cell activities of 17a-hydroxylase:C-17,20lyase are principally responsible for the increases in andro¬ gen production that occur as small antral follicles become preovulatory (102). Therefore, together, these results are consistent with the idea that LH-stimulated production of thecal androgens (via affects on C2i and C27 side-chain cleavage) is the controlling factor in estrogen secretion at proestrus in the rat. In this respect, the regulation of pre¬ ovulatory estrogen secretion in the rat and human appears to be similar. The regulation of follicular estrogen biosynthesis in de¬ veloping follicles might in part be accomplished by secreted regulatory protein(s) that have been identified in fluids of -preovulatory follicles in the human (610,611) and pig (612) and that have been shown to inhibit granulosa cell aromatase activity and reduce responsiveness to gonadotropins (611— 613). The regulatory protein(s) might be one means by which the “selected” or dominant follicle in monotocous species causes neighboring follicles to become atretic. The positive relationship in preovulatory follicles between es¬ trogen concentrations in follicular fluid and levels of aromatase-inhibiting activity suggests that the dominant follicle is insensitive to this inhibitor (613). Evidence in women that this factor is secreted into the venous blood of ovaries containing a preovulatory follicle but not of contralateral ovaries suggests that interovarian, as well as intraovarian, regulation may be involved (614).

Androgens Follicular fluid measurements demonstrate that antral fol¬ licles at all stages of the menstrual cycle in women are significant sources of C19-steroids including androstenedi¬ one, testosterone, and their 5a-reduced metabolites. Since androstenedione and testosterone are not synthesized de novo in significant quantities by granulosa cells, they must be secreted into the follicular fluid by the theca cells of the follicle and perhaps also by the interstitial cells. However, granulosa and theca cells are capable of 5a-reduction of testosterone to 5a-dihydrotestosterone and further metab¬ olism to 5a-androstane-3a,17(3-diol and androsterone, so these metabolites might arise partly by conversion in the granulosa cells. Follicular fluid of women (275,615) and pigs (616) and extracts of rat ovary (79) contain 5a-dihydrotestosterone. The relative levels of androstenedione and testosterone in follicular fluid are probably also influenced by interconversion via 17(3-hydroxysteroid dehydrogenase in the granulosa cells, as discussed earlier. In women, the dominant androgen, androstenedione, has been measured in small and large antral follicles throughout the menstrual cycle (581). Concentrations of androstene-

Follicular Steroidogenesis and Its Control

dione in follicular fluid are between 100 and 500 times that in peripheral plasma (569). The highest concentrations of androstenedione in follicular fluid are found at midfollicular phase (745 ng/ml), with a substantial decrease occurring by the late follicular phase (120 ng/ml) (584). At the late fol¬ licular phase there is a significantly greater concentration of androstenedione in small antral follicles (726 ng/ml) than in large antral follicles (266 ng/ml) (581). At all other phases of the menstrual cycle, the differences between androgen concentrations in large and small antral follicles are not significant. Concentrations of testosterone, although con¬ siderably lower than androstenedione, show a similar pattern of change, with the highest levels occurring in small antral follicles at midcycle (104 ng/ml) but with lower concentra¬ tions in large antral follicles throughout the follicular phase (25-38 ng/ml) (581). Androgen concentrations might be less in large antral follicles at the late follicular phase partly because of sub¬ stantially greater conversion to estrogens and partly because of greater secretion into the blood due to the increased vascularization of the theca (581,617). Another contributing factor in the declining level of androgens observed in fol¬ licular fluid during the later stages of preovulatory follicle growth may be the inhibitory influence of the increasing estradiol concentrations on androgen biosynthesis in the theca cells (618,619). This possibility is supported by observa¬ tions that the decline in androstenedione concentration oc¬ curring between the middle and late follicular phases is seen only in normal healthy follicles characterized by detectable levels of FSH in follicular fluid, while increased andro¬ stenedione concentrations are observed in follicles with un¬ detectable FSH (584). Therefore, the role of FSH in in¬ ducing aromatase activity in granulosa cells appears to be an important factor in regulating follicular androgen con¬ centrations. Estradiol, produced as a result of FSH-induced aromatase activity in granulosa cells, may feed back on the theca cells to inhibit production of its precursors, the aromatizable androgens. Such an intrafollicular negative-feed¬ back system could be significant in limiting the rate of increase of estrogen secretion during the final stages of follicular growth; this would provide adequate time for com¬ pletion of the process of cytoplasmic maturation of the oo¬ cyte before ovulation. An intrafollicular negative-feedback mechanism such as this could be particularly significant in polytocous species as a means of holding in check the most mature follicles in the selected ovulatory population: The less mature of the selected follicles could thereby continue to develop, thus resulting in a synchronous population by the time of the LH surge. In bovine antral follicles, as in the human, changes in androgen and estrogen concentrations in follicular fluid are inversely related as follicles mature. Estradiol concentra¬ tions increase greatly as bovine follicles progress from small to large sizes, while androstenedione and testosterone con¬ centrations decrease (620). A sharper decline in testosterone than androstenedione suggests that, as previously demon¬

/

363

strated (621,622), testosterone is the preferred substrate for aromatization in bovines, in contrast to the situation in the human. Another apparent difference from humans is that the concentration of FSH in bovine follicular fluid is not related to the high ratio of estradiol to androgen found in large follicles (620). This suggests that intrafollicular FSH is less likely to be a limiting factor for aromatization in bovine follicles than in human follicles. The bovine follicle may be exceptional also in another respect, since granulosa cells in this species have been reported to convert pregnen¬ olone to 17a-hydroxprogesterone, and 17a-hydroxypregnenolone to androstenedione, indicating a possible role for these cells in C)9-steroid formation (85). In the hamster, a different relationship of whole follicular concentrations of androgens and estrogens was seen in antral follicles stimulated with PMSG (623). After treatment with PMSG, testosterone and estrogen levels initially increased, the greatest concentrations occurring in middle-sized folli¬ cles; at longer intervals, however, testosterone concentra¬ tions decreased sharply in medium and large follicles, while estradiol and estrone concentrations declined less abruptly. Therefore, according to this model system in the hamster, a possible difference from other species is that androgen availability might become limiting to estrogen biosynthesis even before large antral follicles have formed. Nonaromatic Ci8-steroids are present in the follicular fluid of the mare and sow and are presumed to be products of follicular steroidogenesis. Equine follicular fluid was first shown to contain high levels of 19-norandrostenedione, with concentrations of this steroid at estrus being 30 to 160 ng/ml (4). More recently, 19-nortestosterone was also identified as a minor component in equine follicular fluid (6). Equine granulosa cells convert testosterone primarily to estrogens but also produce 19-nortestosterone and, apparently, lower amounts of 19-norandrostenedione (126), indicating an in¬ trafollicular site of 19-norsteroid biosynthesis. Limited stud¬ ies on follicular stages in the mare report that the highest concentrations of 19-norandrostenedione are present in fluids from large and preovulatory follicles, with lower levels pres¬ ent in less mature and atretic follicles (7). Porcine follicular fluid from preovulatory follicles obtained on days 19 and 20 of the estrous cycle also contains high concentrations of 19-norandrostenedione (21-25 p,mol/liter, 5720-6810 ng/ml), with 19-nortestosterone being tentatively identified as a mi¬ nor component (8). Follicular fluid concentrations of 19norandrostenedione were increased after treatment of im¬ mature pigs with PMSG and decreased following subsequent hCG treatment (624). The physiological significance of 19norsteroids in the follicle is not known, but they have been proposed as possible intermediates in estrogen biosynthesis in the mare (7) and as competitive inhibitors of aromatization in the pig (8). Furthermore, 19-norandrostenedione (0.5 (xmol/liter) acts synergistically with cAMP to inhibit porcine oocyte maturation in vitro, suggesting another possible reg¬ ulatory role in this species (625). Human ovarian tissue in vitro has been reported to convert testosterone to 19-nor-

364

/

Chapter 10

steroids (626), but it has not been established that these steroids are present in human ovaries.

Progestins Progesterone concentrations in follicular fluid of human follicles are much lower than in tissue of corpora lutea (627), but they vary substantially during the menstrual cycle and in small and large antral follicles. During the late follicular phase, concentrations are significantly higher in large antral follicles (~1300 ng/ml) than in small ones (~270 ng/ml) (582). Furthermore, the increase in follicular fluid proges¬ terone concentration in large antral follicles occurs during the transition from midfollicular (760 ng/ml) to late follic¬ ular phases (1720 ng/ml), and this increase is associated with the presence of detectable levels of LH in the fluid (582). These increases in progesterone in the late follicular phase presumably reflect the increased secretion by gran¬ ulosa cells; increased granulosa stimulation by LH would be expected as LH levels progressively rise in the second half of the follicular phase, at a time when FSH levels decline (628). LH has been shown to stimulate progesterone secretion by human granulosa cells from large antral follicles treated with FSH and estrogen (629). This effect of LH on human granulosa cells is presumably as a consequence of LH receptor induction, similar to that demonstrated in rat granulosa cells in vivo (87) and in culture (329). Luteinization of human granulosa cells appears to begin before ovulation since cells isolated in the preovulatory period pro¬ duce large amounts of progesterone (630). Pregnenolone, the immediate precursor of progesterone, is also substantially increased in fluid from large antral fol¬ licles at middle and late follicular phases in women (582). Levels on the order of 10 |xg/ml are reported. Pregnenolone concentrations are also elevated in fluid from human pre¬ ovulatory follicles aspirated 32 to 33 hr after treatment with hCG at midcycle, before the LH surge (631). Pregnenolone has been measured in bovine follicular fluid, with concen¬ trations exceeding those of progesterone in the large pre¬ ovulatory follicles present at proestrus (109). Although the cellular origin of follicular fluid pregnen¬ olone is uncertain, the likelihood that the granulosa cells are the source is supported by observations of high rates of pregnenolone secretion by cultured porcine granulosa cells (215). Evidence has been presented to suggest that preg¬ nenolone in follicular fluid, secreted by granulosa cells, may undergo metabolism to progesterone by bovine theca cells (632) and to aromatizable androgens by porcine theca cells (215).

Preovulatory Follicles After the LH Surge The ovulation-inducing LH surge causes concomitant changes in follicle cell structure, proliferative activity, and steroidogenesis of the preovulatory follicles. These changes

occur in the interval between the LH peak and ovulation, this interval being relatively constant for each species; for example, 12 to 15 hr in mouse or rat and approximately 28 to 36 hr in human (633). Ultrastructural changes occurring in granulosa cells of women prior to ovulation include a continuous change from granular to smooth endoplasmic reticulum (634,635) and a change to a more homogeneous chromatin structure (636). Typical characteristics of luteinization, such as an increase in smooth endoplasmic reticulum and the appearance of mitochondria with tubular cristae, begin to appear even be¬ fore ovulation in many species (575). The ovulatory stim¬ ulus is also associated with changes in the quantities and types of gap junctions present between granulosa cells, as in the rabbit (637,638). Mitoses are abundant in granulosa cells of large preovu¬ latory follicles of certain species, including human (639), hamster (640), and guinea pig (641), but mitotic activity declines sharply between the LH surge and ovulation. In women the high mitotic index and nucleocytoplasmic ratio of granulosa cells in preovulatory follicles are apparently lowered even before the beginning of the LH surge (639). In the estrous rabbit, mitotic activity is greatest in cells of nonvesicular follicles but similarly declines following the mating-induced LH surge (642). These observations suggest that either the LH surge or perhaps related events preceding it are involved in suppressing granulosa cell proliferative activity as cells begin to luteinize. It is unknown whether these proliferative and steroidogenic changes are causally related. However, in vivo studies of PMSG/hCG superovulated rats indicate that increased steroidogenic activity develops only after DNA synthesis arrests (158,643), and recent in vitro studies with rat granulosa cells suggest that cell proliferation in culture and steroidogenic expression are inversely related (186). Histochemical evidence of increased steroidogenic activ¬ ity is also apparent as ovulation approaches. Activity of A53(3-hydroxysteroid dehydrogenase is weak in granulosa cells of developing antral follicles but increases about the time of the estrogen surge (644,645). Similarly in the hamster, activity of this enzyme increases towards the time of ovula¬ tion (640,646,647), when follicular progesterone synthesis increases (648). This enzyme is also active in granulosa cells of preovulatory human follicles, whereas it is limited to thecal cells in smaller antral follicles (585,649). In con¬ trast with these changes in the granulosa cells, theca interna cells show high activity of A5-3(3-hydroxysteroid dehydro¬ genase throughout the maturation of the follicle. Steroid production is substantially altered in the pre¬ ovulatory period following the LH surge. Within minutes of the LH surge there is a transient increase in progesterone and estrogen secretion in several animal species, including rat, monkey, and rabbit, followed by a marked decline to basal levels within several hours (241,310,593,650-653). This inhibitory phase following the LH surge or after hCG treatment is marked by decreased concentrations of aro-

Follicular Steroidogenesis and Its Control

matizable androgens and estrogens in follicular fluid or ova¬ ries of all species studied, including rat (243), hamster (654), sheep (655), bovine (656), and human (657,658). In the rat, a large preovulatory peak of progesterone and 20a-dihydroprogesterone occurs close to the time of the LH peak. These extremely high levels of progestins (i.e., greater than in the short luteal phase at diestrus) are probably de¬ rived from the granulosa cells as they begin the process of luteinization, although the extent of contributions from thecainterstitial cells is uncertain. LH stimulates progesterone secretion in vitro by mature granulosa cells of the rat (293,571), and in vivo pretreatment with testosterone sig¬ nificantly enhances this secretion (289). Measurements of steroids in follicular fluid of preovulatory follicles have been made in rats first given low-dose PMSG treatment to induce follicle maturation and then LH to induce ovulation (243). In these studies, the progesterone concentration in follicular fluid increased sevenfold within 1 hr of LH treatment and remained elevated throughout the 10-hr study. Androstenedione and estradiol concentrations in fluid were elevated before LH treatment, increased slightly 1 to 2 hr after LH, and then fell to extremely low levels by 6 hr. In other studies, inhibition of steroidogenesis, and in particular of estradiol production, occurs during the latter half of the preovulatory period in rat (104,310,650,653), sheep (659), and pig (660). In the rabbit, where the LH surge is a reflex response induced by mating, follicles isolated at 2 and 12 hours after mating showed, respectively, a rapid rise then a fall in progesterone and estradiol synthesis from [14C]acetate (651). A similar type of steroidogenic regulation occurs in the human ovary following the LH surge. Human ovarian slices initially showed increased conversion of [14C] acetate into pregnenolone, progesterone, 17a-hydroxyprogesterone, androstenedione (principal androgen), and estrogens (661). Subsequently, there was substantial reduction in the syn¬ thesis of pregnenolone and 17a-hydroxyprogesterone and, at the same time, almost complete inhibition of androgen and estrogen production (661). Fluid from human follicles after the LH surge also shows a progressive decrease over 38 hr in concentrations of estradiol and androstenedione, testosterone, and dehydroepiandrosterone (657). Similar re¬ sults have been obtained in other studies (588,615,631). Despite the decrease in estradiol concentrations in human follicular fluid following the LH surge, preovulatory folli¬ cles still have the ability to aromatize androgens in vivo (583) and in vitro (662). Progesterone production decreases only slightly in the preovulatory period and then rises stead¬ ily towards ovulation (630,663), while estradiol synthesis remains low (651). Luteinization is well advanced 15 to 20 hr after the beginning of the LH surge, as indicated by high progesterone concentrations in follicular fluid ( 13 p.g/ml), with a further increase occurring ( 18 |xg/ml) after more than 27 hr (662). Progesterone concentrations in follicular fluid have been observed to peak between 5 and 25 hr after the onset of the LH surge, while the concentrations of 17ahydroxyprogesterone change little (657). Measurements of

/

365

steroids in blood during the preovulatory period in women provide similar evidence of changes in ovarian steroid se¬ cretion (630). Regulation of the transient inhibition of steroid production in the preovulatory period of all species probably occurs at several levels. Evidence from luteal cells (356,357,664) and granulosa cells (358,359) indicates that relatively high con¬ centrations of LH cause a transient desensitization at the gonadotropin receptor level such that target cells become refractory to gonadotropin stimulation of adenylate cyclase and steroid production. Desensitization in granulosa cells is followed by a down-regulation of receptors for LH, FSH, and estrogen (665,666). Normally during the luteal phase new receptors form and cells regain their sensitivity to LH at a time when progesterone production greatly increases. Pulsatile release of tonic levels of LH during the follicular phase may be important in preventing desensitization and down-regulation, which naturally occur in response to high concentrations and prolonged exposure to LH at midcycle. Inhibition of estradiol production following the preovu¬ latory LH surge is primarily the result of decreased androgen production in theca and interstitial cells. In an in vitro study with rat Graafian follicles in culture, inhibition of androgen secretion appeared to result from reduced 17a-hydroxylase and/or C-17,20-lyase activity since there was a concomitant inhibition of estradiol, androstenedione, and testosterone accumulation 4 to 6 hr after LH treatment (104). The authors of this study suggested that an LH-stimulated protein factor might inhibit enzymes required for cleavage of the C-17 side chain of progesterone. A subsequent study with PMSGprimed immature rats showed that a maximal ovulatory dose of LH in vivo caused an initial increase in ovarian androgen after 2 hr and then a reduction in androgens and estradiol 4 to 8 hr after LH injection, with serum concentrations declining after 6 hr (310). Aromatase activity in the ovarian microsomal fraction was also inhibited in a noncompetitive manner between 4 and 8 hr after LH treatment in vivo, consistent with either inactivation of the aromatase complex or reduced biosynthesis and rapid turnover of this enzyme. These results led to the suggestion that LH-induced inhi¬ bition of estrogen production in the rat occurred mainly at the level of 17a-hydroxylase:C-17,20-lyase, followed by a secondary decrease in aromatase activity (310). Other in vivo and in vitro studies in rats provide clear evidence that the major cause of the rapid decline in estrogen secretion after the LH surge is limited availability of aromatizable androgens, due to the decreased rate of C2i side-chain cleav¬ age. The decreased ovarian content of estradiol following injection of hCG could be completely prevented by in vivo administration of testosterone (667) or by in vitro incubation of isolated follicles with aromatizable substrate (668). The decreased levels of 17a-hydroxylase:C-17,20-lyase respon¬ sible for the declining androgen production have been found to be associated with a marked reduction in the microsomal cytochrome P-450 levels in the preovulatory ovary. These experiments involving inhibitors of RNA and protein syn-

366 / Chapter 10 thesis provided evidence that the LH-induced decline in activity of these rate-limiting enzymes was dependent on continuing synthesis of RNA and protein (669). Recent studies indicate that the inhibition of the C2i sidechain cleavage reaction following treatment of proestrous rats with an ovulatory dose of hCG is at the levels of NADPHand NADH-linked lyase reactions and selectively at the NADH-linked 17a-hydroxylase (670). It is possible that these apparently distinct lyase and hydroxylase activities detected after cell disruption do not reflect the true nature of the C2i side-chain cleavage system. As mentioned pre¬ viously, direct immunoblot measurements of cytochrome P450 for this enzyme in bovine follicles established that the enzyme protein decreases to undetectable levels at the ear¬ liest stage of corpora lutea formation (105). Inhibition of aromatase cannot be ruled out as a minor regulatory mech¬ anism following LH treatment. Preovulatory estrogen se¬ cretion, which peaks slightly before the LH surge, has been suggested to mediate the inhibitory action on androgen pro¬ duction by a direct intraovarian action on the ovary (233). Support for this idea has been provided by results of in vitro studies indicating that estradiol inhibits androgen production by rat ovarian interstitial cell preparations (86). Evidence for a similar inhibitory effect of estradiol on androgen pro¬ duction by isolated porcine theca cells has also been reported (619) and recently confirmed (235). Morphological signs of theca cell regression suggest that the activity of this cell type is reduced in the late pre¬ ovulatory period in sheep (671) and human (672). Other studies of advanced human preovulatory follicles show that there are fewer theca cells and that they are associated with hemorrhagic lesions when androgen synthesis is minimal (661). However, in the rat, estrogen-priming in vivo actually increases the in vitro LH-stimulated progesterone response of theca cell preparations, suggesting that viability of theca cells is maintained despite reduced androgen production (234). Prostaglandins may also be involved in steroidogenic reg¬ ulation during the preovulatory period. High levels of pros¬ taglandins (F and E series) are found in follicular fluid of the late preovulatory follicles of many species, including the rabbit (673,674), rat (675-677), pig (678,679), and human (680). Prostaglandin synthesis is presumably stim¬ ulated via the action of LH in increasing cAMP. PGF is undetectable in human follicular fluid prior to the LH peak but is high in ovulatory follicles and at ovulation (569,678). Prostaglandins may be involved in desensitization to go¬ nadotropins. However, PGE2, which stimulates progester¬ one production by human granulosa cells in vitro (681), may also maintain or increase progesterone secretion in the late preovulatory period. Furthermore, PGE2 stimulates an¬ drogen production by thecal cells (99,682). Adenylate cy¬ clase is desensitized to LH and FSH at this time, but gran¬ ulosa cells remain responsive to PGE2 (665). Prostaglandins might also mediate effects of LH on increased blood flow and intraovarian distribution of blood (683), thereby influ¬

encing steroid secretion into the circulation. A further im¬ portant role of prostaglandins at ovulation is probably in follicular rupture since this process is blocked by inhibitors of prostaglandin synthetase (684,685) and by systemic (686) or intrafollicular injection of antiserum against prostaglan¬ dins (675). SUMMARY AND CONCLUDING COMMENTS As reviewed in this chapter, the follicle is the principal steroidogenic unit of the ovary. Its biosynthetic activities, and the factors that regulate them, change greatly as it emerges from the pool of resting follicles and undergoes progressive stages of differentiation. This progression leads ultimately to ovulation and luteinization of a small minority of those follicles that leave the resting pool, with the vast majority being aborted in atresia. - The steroidogenic output of the follicle at essentially all stages is a function of the concerted actions of its two types of somatic cells, the theca and granulosa cells, whose ste¬ roidogenic profiles differ in some significant ways. These differences are the result of differences in hormone receptors on their cell membranes, in specific steroidogenic enzyme activities, and in compartmentalization within the follicle, which restricts vascularization to the thecal layers and thus creates substantially different microenvironments for the two cell types. The steroidogenic cells of the follicle are under primary control by the pituitary gonadotropic hormones, FSH and LH, with many of the actions of these hormones being influenced or modulated by a number of intraovarian factors, principally the steroid products of these cells. The gonad¬ otropins initiate the chain of responses in their respective target cells through interaction with specific high-affinity binding sites (receptors) on the cell surfaces. Variation in the cellular concentrations of these receptors appears to dic¬ tate the fate of a given follicle, including its level of steroid output at a given stage of development. The actions of FSH are restricted to the granulosa cells, as all other ovarian cell types appear to lack FSH receptors. In contrast, LH actions are exerted on both follicle cell types, as well as on cells of the interstitial gland and corpus luteum. Granulosa cells at all stages of follicular develop¬ ment appear to possess FSH receptors, whereas they acquire LH receptors and responsiveness only at later developmental stages. Theca cells acquire LH receptors and responsiveness at considerably earlier stages of follicular development than do granulosa cells. FSH and LH exert major effects on steroidogenesis in their respective follicle target cells at least in part through activation of membrane-bound adenylate cyclase, thereby increasing the rate of synthesis of cAMP from ATP. The resulting increased intracellular cAMP levels bring about a variety of physiological responses of the cells through cou¬ pling mechanisms that generally involve phosphorylation and activation of protein kinases. Steroidogenic responses

Follicular Steroidogenesis and Its Control

depend upon the identity and levels of rate-limiting enzymes that the particular target cells possess at the time of stim¬ ulation or acquire subsequently. The earliest steroidogenic response of undifferentiated granulosa cells (those present at early preantral stages of follicle development) to FSH is increased activity of the aromatase enzyme complex. Given an adequate supply of aromatizable androgens (testosterone or androstenedione), the granulosa cells, thus stimulated, are able to increase their rate of synthesis and secretion of estrogens. However, because of an essential absence of the androgen biosynthetic enzymes (17a-hydroxylase:C-17,20-lyase complex) in granulosa cells, estrogen secretion following FSH stimu¬ lation depends on an exogenous supply of androgen, and substrate supply becomes the rate-limiting factor in vivo and in vitro for estrogen biosynthesis by granulosa cells at early developmental stages. FSH also increases the ability of the granulosa cells to secrete progestins through induction of two other rate-lim¬ iting steps in the steroidogenic pathway, cholesterol sidechain cleavage P-450Scc and the A5-3(3-hydroxysteroid dehydrogenase:A54-isomerase enzyme complex. The choles¬ terol substrate required by the granulosa cells for progestin production in vivo (over and above that provided by low rates of constitutive synthesis in the granulosa cells them¬ selves) is provided in the form of lipoprotein-bound cho¬ lesterol reaching the follicle via the blood supply. Lack of vascularization of the granulosa cell layer and essential im¬ permeability of the basement membrane surrounding the granulosa layer to the large lipoprotein molecules limit pro¬ gesterone production by granulosa cells until later preovu¬ latory stages. As the follicle matures, plasma membrane receptors for LH and prolactin are acquired by the granulosa cells. FSH appears to be the primary inducer of these receptors, but the induction is enhanced by autocrine and/or paracrine ac¬ tions of the estrogen produced intracellularly in response to FSH stimulation and acting synergistically with FSH. Once the granulosa cells acquire receptors for LH, this gonado¬ tropin becomes an additional stimulus to cAMP-regulated processes, particularly the further stimulation of the pro¬ gesterone-synthesizing enzymes. Androgen production appears to be the principal steroido¬ genic function of the theca cells. These cells therefore play a major role in enabling production of estrogen by the fol¬ licle, as they supply the substrate that is the rate-limiting factor in estrogen biosynthesis by the aromatase enzyme complex in granulosa cells. Since LH is required for sig¬ nificant rates of androgen production by the theca cells, this gonadotropin may be regarded as the primary regulatory factor in controlling estrogen secretion by all but the most immature of follicles. It is this action of LH on theca cells, together with the action of FSH in induction of aromatase activity in granulosa cells, that forms the basis of the two¬ cell, two-gonadotropin” theory for control of estrogen se¬ cretion in the follicle.

/

367

The increased androgen secretion by theca cells, under LH stimulation, appears to be the result of increased activ¬ ities of two sets of steroidogenic enzymes—the cholesterol side-chain cleavage system and the 17a-hydroxylase:C-17,20lyase complex. Under certain physiological conditions, when the latter enzyme complex is rate-limiting, the products of the former may accumulate with the result that the theca cells become a significant source of C2rsteroid (progestin) secretion. Under other conditions, C2i-steroid synthesis may not keep pace with the activity of the 17a-hydroxylase:C17,20-lyase complex, in which case exogenous progestin of granulosa cell origin may provide significant amounts of the C2i-substrate for theca cell androgen output. This then, forms the basis of a second two-cell theory, for follicular steroidogenesis, in which granulosa and theca cells coop¬ erate in the production of androgens by the follicle. Under certain conditions, in some species more than oth¬ ers, the theca cells appear to possess a sufficient aromatase enzyme system to become a significant source of estrogen secretion as well. Various steroidal influences on the activities of the ste¬ roidogenic enzymes in both granulosa and theca cells come into play as the follicles become more steroidogenically active under the influence of FSH and LH, as summarized in Fig. 5. At least some of the modulating effects of steroids appear to be mediated via intracellular steroid receptor, i.e., the demonstrated existence of estrogen receptors in granu¬ losa cells, and the intraovarian actions of estrogens men¬ tioned above. Other steroid effects appear to occur through direct action on steroidogenic enzymes. In granulosa cells, the action of FSH in induction of aromatase is enhanced by androgens, as well as by estro¬ gens, and inhibited, under certain circumstances, by 5areduced androgens and perhaps by progesterone. The 5areduced androgens also act as competitive inhibitors of the aromatase system. Both stimulatory and inhibitory actions of androgens have been reported on FSH-induced proges¬ terone biosynthesis in granulosa cells. The stimulatory ac¬ tion, studied extensively in rat granulosa cells, appears to involve enhancement of FSH induction of the cytochrome P-450Scc and the A5-3(3-hydroxysteroid dehydrogenase: A5'4-isomerase enzymes. The inhibitory action, demon¬ strated with porcine granulosa cells, appears to be primarily at the level of the A5-3(3-hydroxysteroid dehydrogenase: A5'4-isomerase complex. Estrogens have been shown to influence androgen bio¬ synthesis by theca cells, through inhibition at the level of the 17a-hydroxylase:C-17,20-lyase system. This paracrine action of estrogen may function as an intrafollicular nega¬ tive-feedback system to limit its own production in granu¬ losa cells through regulation of substrate availability. Prolactin appears to exert inhibitory influences on estro¬ gen biosynthesis, by inhibiting both FSH-induced aromatase activity in granulosa cells and LH-induced androgen pro¬ duction by theca/interstitial cells. A further well-established action of prolactin is its role in stimulating progesterone

368

/

Chapter 10

FIG. 5. Principal sites of regulation of follicular steroidogenesis in the rat. The scheme compresses into one diagram the prin¬ cipal regulatory sites of action of gonadotropins and steroids that have been demonstrated either at various stages of fol¬ licular maturation in vivo or with isolated cell types. (See section on steroidogenesis and its control by cell type for details.) The proposed sequence of events is as follows. Undifferentiated granulosa cells initially respond exclusively to FSH, by a cAMPdependent mechanism, thereby stimulating activities of en¬ zymes required for the metabolism of cholesterol to proges¬ terone and for the conversion of theca-derived androgens to estrogens. Steroid biosynthesis in the theca cells is stimulated exclusively by LH. As the follicle matures, plasma membrane receptors for LH and prolactin (PRL) are acquired by the gran¬ ulosa cells as a result of both FSH action and stimulation by estrogen on this process. LH may initially contribute to gran¬ ulosa cell differentiation by augmenting the various cAMP-dependent processes. A variety of steroidal influences on the activities of steroidogenic enzymes may come into play as the follicles become more biosynthetically active. These effects may occur as the result of receptor-mediated actions of estro¬

production by granulosa cells undergoing luteinization dur¬ ing the periovulatory period. The resulting high levels of progesterone may further contribute to the inhibition of es¬ trogen synthesis in the follicle by prolactin. The cyclic changes in follicular steroid biosynthesis dur¬ ing the estrous or menstrual cycle—as well as the more basal secretion rates of follicular activity during other phys¬ iological states such as pregnancy, seasonal and lactational anestrus (or amenorrhea), and prepuberty—can probably be

gens, androgens, and progesterone, thereby influencing re¬ sponsiveness to FSH and LH, or steroids may directly alter enzymatic activities; these different actions of steroids are not distinguished in the above scheme. The apparent induction of aromatase activity is augmented by androgenic and estrogenic actions but is inhibited by progestins under certain circum¬ stances. 5a-Reduced metabolites of androgens may compet¬ itively inhibit the aromatase enzyme, and estrogens may inhibit 5a-reductase. Prolactin has been shown to inhibit the induction of aromatase activity in vitro and also does so in vivo under certain physiological conditions, but its role in regulating es¬ trogen synthesis during the ovarian cycle is still uncertain. An¬ drogens may also facilitate the synthesis of progesterone. Fol¬ lowing the LH surge, steroid production is briefly stimulated and then suppressed as granulosa and theca cells become desensitized to gonadotropin stimulation. Transiently in¬ creased estrogen secretion by granulosa cells causes inhibi¬ tion of follicular estrogen biosynthesis by inhibiting androgen biosynthesis in the theca cells. Progesterone biosynthetic ac¬ tivity is restored subsequently and increases greatly as follicle cells luteinize.

explained on the bases of the coordinated actions of those cellular mechanisms and regulatory agents discussed in this chapter. The physiological importance of the pituitary go¬ nadotropins and prolactin, as well as of the steroidal prod¬ ucts of the follicular cells, is widely accepted. There are a number of other regulatory molecules whose specific roles in ovarian regulation are less well established but for which there is mounting evidence, largely from in vitro studies with isolated ovarian cells, of at least a supporting or per-

Follicular Steroidogenesis and Its Control

missive role in follicular regulation. Those we have included in this review are GnRH-like peptides, corticosteroids, in¬ sulin and other growth factors, and various neurotransmit¬ ters. We have undoubtedly overlooked others of emerging importance. Whether the demonstrated effects of these sub¬ stances represent physiologically important actions or merely modulations peculiar to the in vitro systems remains un¬ certain. Nevertheless, it is abundantly clear that the follic¬ ular microenvironment, as embodied in the fluid of antral follicles, contains a wide array of varied substances (in¬ cluding steroids, eicosanoids, peptides, proteins, glycopro¬ teins, and proteoglycans), some of which have hormonal or other biological activities in vitro and which could have significant regulatory effects on gonadotropin action and/or follicular steroidogenesis.

ACKNOWLEDGMENTS We wish to thank Dr. S.A.J. Daniel for expert assistance in preparing material for portions of this chapter and Dr. M. W. Khalil for valuable discussions. We are also indebted to Mr. G. Barbe, Mrs. H. E. Ross, and Ms. U. Williams for diligent assistance in the preparation of the manuscript. The authors’ research is supported by a group grant from the Medical Research Council (MRC) of Canada; D.T.A. is a Career Investigator of the MRC, and R.E.G.-L. is the recipient of a Career Scientist Award from the Ontario Min¬ istry of Health, Health Research Personnel Development Program.

REFERENCES 1. Kellie, A. E. (1984) Structure and nomenclature. In: Biochemistry of Steroid Hormones, edited by H. L. J. Makin, pp. 1-19. Blackwell Scientific, Oxford, UK. 2. Doisy, E. A., Veler, C. D., and Thayer, S. (1929): Folliculin from urine of pregnant women. Am. J. Physiol., 90:329-330. 3. MacCorquodale, D. W., Thayer, S. A., and Doisy, E. A. (1936). The isolation of the principal estrogenic substance of liquor folliculi. J. Biol. Chem., 115:435-448. 4. Short, R. V. (1961): Steroid concentrations in the follicular fluid of mares at various stages of the reproductive cycle. J. Endocrinol., 22:153-163. 5. Dehennin, L., Blacker, C., Reiffsteck, A., and Scholler, R. (1984): Estrogen 2-, 4-, 6- or 16-hydroxylation by human follicles shown by gas chromatography-mass spectrometry associated with stable isotope dilution. J. Steroid Biochem., 20:465-471. 6. Silberzahn, P., Almahbobi, G., Dehennin, L., and Merouane, A. (1985): Estrogen metabolite in equine ovarian follicles; gas chro¬ matographic-mass spectrometric determinations in relation to follic¬ ular ultrastructure and progestin content. J. Steroid Biochem., 22:5017. Silberzahn, P., Dehennin, L., Zwain, I., and Reiffsteck, A. (1985): Gas chromatography-mass spectrometry of androgens in equme ovar¬ ian follicles at ultrastructurally defined stages of development. Iden¬ tification of 19-nortestosterone in follicular fluid. Endocrinology, 8

117:2176-2181. Khalil M W., and Walton, J. S. (1985): Identification and mea¬ surement of 4-oestren-3,17-dione (19-norandrostenedione) in porcine ovarian follicular fluid using high performance liquid chromatogra¬ phy and capillary gas chromatography-mass spectrometry. J. En¬ docrinol., 107:375-381.

/

369

9. Hill, R. T. (1937): Ovaries secrete male hormone. I. Restoration of the castrate type of seminal vesicle and prostate glands to normal by grafts of ovaries in mice. Endocrinology, 21:495-502. 10. Short, R. V. (1960): Steroids present in the follicular fluid of the mare. J. Endocrinol., 20:147-156. 11. Ryan, K. J., and Petro, Z. (1966): Steroid biosynthesis by human ovarian granulosa and thecal cells. J. Clin. Endocrinol. Metab., 26:46-52. 12. Springer, C., and Eckstein, B. (1971): Regulation of production in vitro of 5a-androstane-3a,17p-diol in the immature rat ovary. J. Endocrinol., 50:431 —439. 13. Zmigrod, A., Lindner, H. R., and Lamprecht, S. A. (1972): Re¬ ductase pathways of progesterone metabolism in the rat ovary. Acta Endocrinol. (Copenh.), 69:141-152. 14. Karakawa, T., Karachi, K., Aono, T., and Matsumoto, K. (1976): Formation of 5a-reduced Ci9-steroids from progesterone in vitro by a pathway through 5a-reduced C2i-steroids in ovaries of late pre¬ pubertal rats. Endocrinology, 98:571-579. 15. Tsuji, M., Terada, N., Sato, B., and Matsumoto, K. (1982): 5(3and 5a-reductases for 4-ene-3-ketosteroids in golden hamster ovaries at different stages of development. J. Steroid Biochem., 16:207-213. 16. Gower, D. B. (1972): 16-Unsaturated C|9-steroids. A review of thenchemistry, biochemistry and possible physiological role. J. Steroid Biochem., 3:45-103. 17. Armstrong, D. T. (1979): Alterations of progesterone metabolism in immature rat ovaries by luteinizing hormone. Biol. Reprod., 21:10251033. 18. Fieser, L. F., and Fieser, M. (1959): Steroids. Reinhold, New York. 19. Dorfman, R. T., and Ungar, F. (1965): Metabolism of Steroid Hor¬ mones. Academic, New York. 20. Makin, H. L. J. (ed.) (1984): Biochemistry of Steroid Hormones. Blackwell, Oxford, UK. 21. Strauss III, J. F., Schuler, L. A., Rosenblum, M. F., and Tanaka, T. (1981): Cholesterol metabolism by ovarian tissue. Adv. Lipid Res., 18:99-157. 22. Solod, E. A., Armstrong, D. T., and Greep, R. O. (1966): Action of luteinizing hormone on conversion of ovarian cholesterol stores to steroid secreted in vivo and synthesized in vitro by the pseudo¬ pregnant rabbit ovary. Steroids, 7:607-620. 23. Gwynne, J. T., and Strauss III, J. F. (1982): The role of lipoproteins in steroidogenesis and cholesterol metabolism in steroidogenic glands. Endocr. Rev., 3:299-329. 24. Veldhuis, J. D., Klase, P. A., Strauss III, J. F., and Hammond, J. M. (1982): Facilitative interactions between estradiol and lutein¬ izing hormone in the regulation of progesterone production by cul¬ tured swine granulosa cells: relation to cholesterol metabolics. En¬ docrinology, 111:441^447. 25. Rajendran, K. C., Hwang, J., and Menoir, K. M. J. (1983): Binding, degradation and utilization of plasma high density and low density lipoproteins for progesterone production in cultured rat luteal cells. Endocrinology, 112:1746-1753. 26. Shalgi, R., Kraicer, P., Renoir, A., Pinto, M., and Soferman, N. (1973): Proteins of human follicular fluid: the blood-follicle barrier. Fertil. Steril., 24:429—434. 27. Chang, S. C. S., Jones, J. D., Ellefson, R. D., and Ryan, R. J. (1976): The porcine ovarian follicle: I. Selected chemical analysis of follicular fluid at different developmental stages. Biol. Reprod., 15:321-328. 28. Simpson, E. R., Rochelle, D. B., Carr, B. R., and MacDonald, P. C. (1980): Plasma lipoproteins in follicular fluid of human ovaries. J. Clin. Endocrinol. Metab., 51:1469-1471. 29. Wang, S. C., and Greenwald, G. S. (1984): Effect of lipopro¬ teins, 25-hydroxycholesterol and luteinizing hormone on in vitro follicular steroidogenesis in the hamster and rat. Biol. Reprod., 31:271-279. 30. Farkash, Y., Timberg, R., and Orly, J. (1986): Preparation of anti¬ serum to rat cytochrome P-450 cholesterol side chain cleavage, and its use for ultrastructural localization of the immunoreactive enzyme by protein A-gold technique. Endocrinology, 118:1353-1365. 31. Lieberman, S., Greenfield, N. J., and Wolfson, A. (1984): A heu¬ ristic proposal for understanding steroidogeneic processes. Endocr. Rev., 5:128-148. 32. Hall, P. F. (1984): Cellular organization for steroidogenesis. Ini. Rev. Cytol., 86:53-95.

370

/

Chapter 10

33. Gower, D. B. (1984): The role of cytochrome P-450 in steroido¬ genesis and properties of some of the steroid-transforming enzymes. In: Biochemistry of Steroid Hormones, edited by H. L. J. Makin, pp. 230-292. Blackwell Scientific, Oxford, UK. 34. Kautsky, M. P., and Hagerman, D. D. (1980): Kinetic properties of steroid 19-hydroxylase and estrogen synthetase from porcine ovary microsomes. J. Steroid Biochem., 13:1283-1290. 35. Brodie, A. M., Schwarzel, W. C., and Brodie, H. J. (1976): Studies on the mechanism of estrogen biosynthesis in the rat ovary. J. Steroid Biochem., 7:787-793. 36. Mason, N. R. (1970): Steroid A-ring reduction by rat ovaries. En¬ docrinology, 87:350-355. 37. Eckstein, B., and Ravid, R. (1974): On the mechanism of the onset of puberty: identification and pattern of 5a-androstane-3(3,17(3-diol and its 3oc-epimer in peripheral blood of immature female rats. En¬ docrinology, 94:224-229. 38. Mizutani, S., Akashi, S., Terada, N., and Matsumoto, K. (1979): A comparison of metabolism of progesterone to 5a-steroids in mon¬ key, mouse and rat ovaries. J. Steroid Biochem., 10:549-552. 39. Furubayashi, Y., Terada, N., Sato, B., and Matsumoto, K. (1982): Localization of A4-5p- and 5u.reductases and 17(3-ol-dehydrogenase in immature golden hamster testis. Endocrinology, 111:269-272. 40. Inaba, T., Imori, T., and Matsumoto, K. (1978): Formation of 5a-reduced C 19-steroids from progesterone in vivo by 5a-reduced pathway in immature rat ovaries. J. Steroid Biochem., 9:11051110. 41. Zmigrod, A., and Lindner, H. R. (1969): Metabolism of progesterone by the rat ovary: formation of 3(3-hydroxy-5a-pregnan-20-one by ovarian microsomes. Acta Endocrinol. (Copenh.), 61:618-628. 42. Armstrong, D. T., Kraemer, M. A., and Hixon, J. E. (1975): Me¬ tabolism of progesterone by rat ovarian tissue: influence of pregnant mare serum gonadotrophin and prolactin. Biol. Reprod., 12:599— 608. 43. Ichikawa, S., Morioka, H., and Sawada, T. (1971): Identification of the neutral steroids in the ovarian venous plasma of LH-stimulated rats. Endocrinology, 88:372-383. 44. Eckstein, B. (1974): The origin of 5a-androstane-3a,17(3-diol and its 33 epimer in peripheral blood of immature female rats. J. Steroid Biochem., 5:577-580. 45. Eckstein, B. (1983): Blood concentrations and biological effects of androstanediols at the onset of puberty in the female rat. J. Steroid Biochem., 19:883-886. 46. Hashimoto, I., and Wiest, W. G. (1969): Luteotrophic and luteolytic mechanisms in rat corpora lutea. Endocrinology, 80:886-892. 47. Goldring, N. B., and Orly, J. (1985): Concerted metabolism of steroid hormones produced by cocultured ovarian cell types. J. Biol. Chem., 260:913-920. 48. MacLusky, N. J., Naftolin, F., Krey, L. C., and Franks, S. (1981): The catechol estrogens. J. Steroid Biochem., 15:111-124. 49. Hammond, J. M., Hershey, R. M., Walega, M. A., and Weisz, J. (1986): Catecholestrogen production by porcine ovarian cells. En¬ docrinology, 118:2292-2299. 50. Ball, R., and Knuppen, R. (1980): Catechol oestrogens (2- and 4hydroxyoestrogen): chemistry, biogenesis, metabolism, occurrence and physiological significance. Acta Endocrinol. (Copenh.), 93 (Suppl. 232): 1-27. 51. Richards, J. S. (1979): Hormonal control of ovarian follicular de¬ velopment. Rec. Prog. Horm. Res., 35:343-368. 52. Richards, J. S. (1980): Maturation of ovarian follicles: actions and interactions of pituitary and ovarian hormones on follicular cell dif¬ ferentiation. Physiol. Rev., 60:51-89. 53. Robison, G. A., Butcher, R. W., and Sutherland, E. W. (1971): Cyclic AMP. Academic, New York. 54. Kuo, J. F., and Greengard, P. (1969): Cyclic nucleotide-dependent protein kinases. IV. Widespread occurrence of adenosine 3',5'monophosphate-dependent protein kinase in various tissues and phyla of the animal kingdom. Proc. Natl. Acad. Sci. USA, 64:1349-1355. 55. Carnegie, J., and Tsang, B. K. (1983): Follicle-stimulating hormoneregulated granulosa cell steroidogenesis: involvement of the calciumcalmodulin system. Am. J. Obstet. Gynecol., 145:223-228. 56. Byskov, A. G. (1978): The anatomy and ultrastructure of the rete system in the fetal mouse ovary. Biol. Reprod., 19:720-735. 57. Byskov, A. G., and Rasmussen, G. (1973): Ultrastructural studies of the developing follicle. In: The Development and Maturation of

the Ovary and Its Function, International Congress Series No. 267 edited by H. Peters, pp. 55-62. Excerpta Medica, Amsterdam. 58. Bjersing, L. (1967): On the ultrastructure of follicles and isolated follicular granulosa cells of porcine ovary. Z. Zellforsch. Mikrosk. Anat., 82:173-186. 59. Motta, P., and Didio, L. J. A. (1974): Microfilaments in granulosa cells during the follicular development and transformation in corpus luteum in the rabbit ovary. J. Submicrosc. Cytol., 6:15-27. 60. Zamboni, L. (1974): Fine morphology of the follicle wall and follicle cell-oocyte association. Biol. Reprod., 10:125-149. 61. Byskov, A. G. (1975): The role of rete ovarii in meiosis and follicle formation in the cat, mink and ferret. J. Reprod. Fertil., 45:201289. 62. Amsterdam, A., Koch, Y., Lieberman, M. E., and Lindner, H. R. (1975): Distribution of binding sites for human chorionic gonado¬ tropin in the preovulatory follicle of the rat. J. Cell Biol., 67:894900. 63. Midgley, Jr., A. R. (1972): Gonadotropin binding to frozen sections of ovarian tissue. In: Gonadotropins, edited by B. B. Saxena, C. G. Beling, and H. M. Gandy, pp. 248-260. Wiley-Interscience, New York. 64. Zoller, L. C., and Weisz, J. (1979): A quantitative cytochemical study of glucose-6-phosphate dehydrogenase and A5-3|3-hydroxysteroid dehydrogenase activity in membrana granulosa of the ovulable type of follicle of the rat. Histochemistry, 62:125-135. 65. Zoller, L. C., and Weisz, J. (1978): Identification of cytochrome P450 and its distribution in the membrana granulosa of the preovulatory follicle using quantitative cytochemistry. Endocrinology, 103:310313. 66. Erickson, G. F., Hofeditz, C., Unger, M., Allen, W. R., and Dulbecco, R. (1985): A monoclonal antibody to a mammary cell line recognizes two distinct subtypes of ovarian granulosa cells. Endo¬ crinology, 117:1490-1499. 67. Kasson, B. G., Median, R., Davoren, J. B., and Hsueh, A. J. W. (1985): Identification of subpopulations of rat granulosa cells: sedi¬ mentation properties and hormonal responsiveness. Endocrinology, 117:1027-1034. 68. Weakly, B. S. (1966): Electron microscopy of the oocyte and gran¬ ulosa cells in the developing ovarian follicles of the golden hamster. J. Anat., 100:503-534. 69. Hoage, T. R., and Cameron, I. L. (1976): Folliculogenesis in the ovary of the mature mouse: an autoradiographic study. Anat. Rec., 184:699-710. 70. H^Syer, P. E., and Byskov, A. G. (1981): A quantitative cytochemical study of A5, 3f3-hydroxysteroid dehydrogenase activity in the rete system of the immature mouse ovary. In: Development and Function of Reproductive Organs, edited by A. G. Byskov and H. Peters, pp. 216-224. Excerpta Medica, Amsterdam. 71. Quattropani, S. L. (1973): Morphogenesis of the ovarian interstitial tissue in the neonatal mouse. Anat. Rec., 177:569-584. 72. Mori, H., and Matsumoto, K. (1970): On the histogenesis of the ovarian interstitial gland in rabbits. I. Primary interstitial gland. Am J. Anat., 129:289-306. 73. Kingsbury, B. F. (1939): Atresia and the interstitial cells of the ovary. Am. J. Anat., 65:309-331. 74. Dawson, A. B., and McCabe, M. (1951): The interstitial tissue of the ovary in infantile and juvenile rat. J. Morphol., 88:543-571. 75. Rennels, E. G. (1951): Influence of hormones on the histochemistry of ovarian interstitial tissue in the immature rat. Am. J. Anat 88 63108. 76. Deanesly, R. (1972): Origins and development of interstitial tissue in ovaries in rabbit and guinea-pig. J. Anat., 113:251-260. 77. Guraya, S. S., and Greenwald, G. S. (1968): A comparative histochemical study of interstitial tissue and follicular atresia in the mammalian ovary. Anat. Rec., 149:411-434. 78. McNatty, K. P. (1981): Hormonal correlates of follicular develop¬ ment in the human ovary. Aust. J. Biol. Sci., 34:249-268. 79. Belanger, A., Cusan, L., Caron, S., Barden, N., and Dupont, A. (1981): Ovarian progestins, androgens and estrogen throughout the 4-day estrous cycle in the rat. Biol. Reprod., 24:591-596. 80. Sridaran, R., and Gibori, G. (1983): Intraovarian localization of luteinizing hormone/human chorionic gonadotropin stimulation of testosterone and estradiol synthesis in the pregnant rat. Endocrinol¬ ogy, 112:1770-1776.

Follicular Steroidogenesis and Its Control /

81. Dennefors, B. L., Janson, P., Knutson, F., and Hamberger, L. (1980): Steroid production and responsiveness to gonadotrophin in isolated stromal tissue of human menopausal ovaries. Am. J. Obstet. Gynecol., 136:997-1002. 82. Longcope, C., Hunter, R., and Franz, C. (1980): Steroid secretion by the postmenopausal ovary. Am. 7. Obstet. Gynecol., 138:564568. 83. Dyer, C. A., Erickson, G. F., and Curtiss, L. K. (1985): Functional heterogeneity in the ability of high density lipoproteins to enhance gonadotropin-induced androgen production in cultured rat theca-in¬ terstitial cells. In: Lipoprotein and Cholesterol Metabolism in Ste¬ roidogenic Tissues, edited by J. F. Strauss III and K. M. J. Menon, pp. 141-146. Strickley, Philadelphia. 84. Aakvaag, A. (1969): Pathways in the biosynthesis of androstenedione in the human ovary in vitro. Acta Endocrinol. (Copenh.), 60:517526. 85. Lacroix, E., Eechaute, W., and Leusen, I. (1974): The biosynthesis of oestrogens by cow follicles. Steroids, 23:337-356. 86. Magoffin, D. A., and Erickson, G. F. (1982): Primary culture of differentiating ovarian androgen-producing cells in defined medium. J. Biol. Chem., 257:4507-4513. 87. Zeleznik, A. J., Midgley Jr., A. R., and Reichert Jr., L. E. (1974): Granulosa cell maturation in the rat: increased binding of human chorionic gonadotropin following treatment with follicle-stimulating hormone in vivo. Endocrinology, 95:818-825. 88. Lindner, H. R., Amsterdam, A., Salomon, Y., Tsafriri, A., Nimrod, A., Lamprecht, S. A., Zor, U., and Koch, Y. (1977): Intraovarian factors in ovulation: determinants of follicular response to gonado¬ tropins. J. Reprod. Fertil., 51:215-235. 89. Zeleznik, A. J., Schuler, H. M., and Reichert Jr., L. E. (1981): Gonadotropin-binding sites in the rhesus monkey ovary: role of the vasculature in the selective distribution of human chorionic gonad¬ otropin in the preovulatory follicle. Endocrinology, 109:356-362. 90. Oxberry, B. A., and Greenwald, G. S. (1982): An autoradiographic study of the binding of l25I-labelled follicle-stimulating hormone, human chorionic gonadotrophin and prolactin to the hamster ovary throughout the estrous cycle. Biol. Reprod., 27:505-516. 91. Shaha, C., and Greenwald, G. S. (1983): Development of steroido¬ genic activity in the ovary of the prepubertal hamster. I. Response to in vivo or in vitro exposure to gonadotropins. Biol. Reprod., 28:1231-1241. 92. Uilenbroek, J. T. J., and van der Linden, R. (1983): Changes in gonadotrophin binding to rat ovaries during sexual maturation. Acta Endocrinol. (Copenh.), 104:413-419. 93. Henderson, K. M., Kieboom, L. E., McNatty, K. P., Lun, S., and Heath, D. A. (1984): 125I-HCG binding to bovine thecal tissue from healthy and atretic antral follicles. Mol. Cell. Endocrinol., 34:91-98. 94. Hamberger, L., Hillensjo, T., and Ahren, K. (1978): Steroidogenesis in isolated cells of preovulatory rat follicles. Endocrinology, 103:771—

111. 95. Weiss, T. J., Armstrong, D. T., McIntosh, J. E. A., and Seamark, R. F. (1978): Maturational changes in sheep ovarian follicles: go¬ nadotrophic stimulation of cyclic AMP production by isolated theca and granulosa cells. Acta Endocrinol. (Copenh.), 89:158-165. 96. Tsang, B. K., Moon, Y. S., Simpson, C. W., and Armstrong, D. T. (1979): Androgen biosynthesis in human ovarian follicles: cellular source, gonadotropic control, and andenosine 3 ,5 -mono¬ phosphate mediation. J. Clin. Endocrinol. Metab., 48.153—158. 97. Erickson, G. F., Magoffin, D. A., Dyer, C. A., and Hofeditz, C. (1985): The ovarian androgen producing cells: a review of struc¬ ture/function relationships. Endocr. Rev., 6:371-399. 98. Erickson, G. F., and Ryan, K. J. (1976): Stimulation of testosterone production in isolated rabbit thecal tissue by LH/FSH, dibutyryl cyclic AMP, PGF2o, and PGE2. Endocrinology, 99:452-458. 99. Tsang, B. K., Armstrong, D. T., and Whitfield, J. F. (1980): Steroid biosynthesis by isolated human ovarian follicular cells in vitro. J. Clin. Endocrinol. Metab., 51:1407—1411. 100. Fukuda, S., Terakawa, N., Sato, B., Imori, T., and Matsumoto, K. (1979): Hormonal regulation of activities of 17(3-ol-dehydrogenases, aromatase and 4-ene-5a-reductase in immature ovaries. J. Steroid Biochem., 11:1421-1427. 101. Aono, T., Kitamura, Y., Fukuda, S., and Matsumoto, K. (1981): Localization of 4-ene-5a-reductase, 17f3-ol-dehydrogenase and aro¬ matase in immature rat ovary. J- Steroid Biochem., 14.1369-1377.

371

102. Bogovich, K., and Richards, J. S. (1982): Androgen biosynthesis in developing ovarian follicles: evidence that luteinizing hormone reg¬ ulates thecal 17a-hydroxylase and Ci7-2o-lyase activities. Endocri¬ nology, 111:1201-1208. 103. Makris, A., and Ry an, K. J. (1980): The source of follicular androgen in the hamster follicle. Steroids, 35:53-64. 104. Lieberman, M. E., Bamea, A., Bauminger, S., Tsafriri, A., Collins, W. P., and Lindner, H. R. (1975): LH effect on the pattern of steroidogenesis in cultured Graafian follicles of the rat: dependence on macromolecular synthesis. Endocrinology, 96:1533-1542. 105. Rodgers, R. J., Waterman, M. R., and Simpson, E. R. (1986): Cytochromes P-450scc, P-450i7ct, adrenodoxin, and reduced nicotin¬ amide adenine dinucleotide phosphate-cytochrome P-450 reductase in bovine follicles and corpora lutea. Changes in specific contents during the ovarian cycle. Endocrinology, 118:1366-1374. 106. Makris, A., and Ryan, K. J. (1975): Progesterone, androstenedione, testosterone, estrone, and estradiol synthesis in hamster ovarian fol¬ licle cells. Endocrinology, 96:694-701. 107. Evans, G., Dobias, M., King, G. J., and Armstrong, D. T. (1981): Estrogen, androgen, and progesterone biosynthesis by theca and gran¬ ulosa of preovulatory follicles in the pig. Biol. Reprod., 25:673682. 108. McNatty, K. P., Heath, D. A., Lun, S., Fanin, J. M., McDiarmid, J. M., and Henderson, K. M. (1984): Steroidogenesis by bovine theca interna in an in vitro perfusion system. Biol. Reprod., 30:159170. 109. Fortune, J. E., and Hansel, W. (1985): Concentrations of steroids and gonadotropins in follicular fluid from normal heifers and heifers primed for superovulation. Biol. Reprod., 32:1069-1079. 110. McNatty, K. P., Makris, A., Reinhold, V. N., DeGrazia, C., Osathanondh, R., and Ryan, K. J. (1979): Metabolism of andro¬ stenedione by human ovarian tissues in vitro with particular refer¬ ences to reductase and aromatase activity. Steroids, 34:429^443. 111. McNatty, K. P., Makris, A., Osathanondh, R., and Ryan, K. J. (1980): Effects of luteinizing hormone on steroidogenesis by thecal tissue from human ovarian follicles in vitro. Steroids, 36:53-63. 112. Lemer, N., and Eckstein, B. (1976): Identification of two 5a-reduced pregnanes as major metabolites of progesterone in immature rat ova¬ ries (1000 x g supernatant) in vitro. Endocrinology, 98:179-188. 113. Bogovich, K., Scales, L. M., Higginbottom, E., Ewing, L. L., and Richards, J. S. (1986): Short term androgen production by rat ovarian follicles and long term steroidogenesis by thecal explants in culture. Endocrinology, 118:1379-1386. 114. Eckstein, B., Mechoulam, R., and Burstein, S. H. (1970): The identification of 5a-androstane-3a,17(3-diol as a principal metabolite of pregnenolone in rat ovary at the onset of puberty. Nature, 228:866-

868. 115. Suzuki, K., Kawakura, K., and Tamaoki, B. I. (1978): Effect of pregnant mare’s serum gonadotrophin on the activities of 5a-reductase, aromatase, and other enzymes in the ovaries of immature rats. Endocrinology, 102:1595-1605. 116. Eckstein, B., and Ravid, R. (1979): Changes in pathways of steroid production taking place in the rat ovary around the time of the first ovulation. J. Steroid Biochem., 11:593-597. 117. Eckstein, B., Golan, R., and Mishinsky, J. S. (1973): Onset of puberty in the immature female rat induced by 5a-androstane-3f3,17pdiol. Endocrinology, 92:941-945. 118. Eckstein, B., and Springer, C. (1971): Induction of an ovarian epimerase system catalyzing the transformation of 5a-androstane-3a, 17(3diol to 5a-androstane-3p,17|3-diol after treatment of immature rats with gonadotropins exhibiting FSH-like activity. Endocrinology, 89:347-352. 119. Florensa, E., Sommerville, I. F., Harrison, R. F., Johnson, M. W., and Youssefnejadian, E. (1976): Plasma 20a-dihydroprogesterone, progesterone and 17-hydroxyprogesterone: daily and four-hourly variations during the menstrual cycle. J. Steroid Biochem., 7:769777. 120. Holmdahl, T. H., and Johansson, E. D. B. (1972): Peripheral plasma levels of 17a-hydroxyprogesterone, progesterone and oestradiol dur¬ ing normal menstrual cycles in women. Acta Endocrinol. (Copenh.), 71:743-754. 121. Aedo, A. R., Langren, B. M., Cekan, Z., and Diczfalusy, E. (1976): Studies on the pattern of circulating steroids in the normal menstrual cycle. Acta Endocrinol. (Copenh.), 82:600-616.

372

/

Chapter 10

122. Ryan, K. J., Petro, Z., and Kaiser, J. (1968): Steroid formation by isolated and recombined ovarian granulosa and theca cells. J. Clin. Endocrinol. Metab., 28:355-358. 123. Charming, C. P. (1969): Steroidogenesis and morphology of human ovarian cell types in tissue culture. J. Endocrinol., 45:297-308. 124. Batta, S. K., Wentz, A. C., and Channing, C. P. (1980): Steroido¬ genesis by human ovarian cell types in culture: influence of mixing of cell types and effect of added testosterone. J. Clin. Endocrinol. Metab, 50:274-279. 125. Channing, C. P., Wentz, A. C., and Jones, G. (1978): Steroid secretion by monkey and human ovarian cell types in vivo and in vitro. In: Symposium on the Ovary held in Fresnes, France, edited by R. Scholler, pp. 71-86. Editions Sepe, Paris. 126. Ryan, K. J., and Short, R. V. (1965): Formation of estradiol by granulosa and thecal cells of the equine ovarian follicle. Endocri¬ nology, 76:108-114. 127. Channing, C. P., and Grieves, S. A. (1969): Studies on tissue culture of equine ovarian cell types: steroidogenesis. J. Endocrinol., 43:391402. 128. Armstrong, D. T., Weiss, T. J., Selstam, G., and Seamark, R. F. (1981): Hormonal and cellular interactions in follicular steroid bio¬ synthesis by the sheep ovary. J. Reprod. Fertil. (Suppl. 30), 30:143154. 129. Moon, Y. S., Tsang, B. K., Simpson, C., and Armstrong, D. T. (1978): 17p-Estradiol biosynthesis in cultured granulosa and thecal cells of human ovarian follicles: stimulation by follicle-stimulating hormone. J. Clin. Endocrinol. Metab., 47:263-267. 130. Hillier, S. G. (1981): Regulation of follicular oestrogen biosynthesis: a survey of current concepts. J. Endocrinol., 89:3P-18P. 131. Hillier, S. G., van den Boogaard, A. J. M., Reichert Jr., L. E., and van Hall, E. V. (1981): Control of preovulatory follicular estrogen biosynthesis in the human ovary. J. Clin. Endocrinol. Metab., 52:847856. 132. Mossman, H. W., Koering, M. J., and Ferry Jr., D. (1964): Cyclic changes of interstitial gland tissue of the human ovary. Am. J. Anat 115:235-256. 133. Leymarie, P., and Savard, K. (1968): Steroid hormone formation in the human ovary. VI. Evidence for two pathways of synthesis of androgens in the stromal compartment. J. Clin. Endocrinol. Metab 28:1547-1554. 134. Marsh, J. M., Savard, K., and LeMaire, W. J. (1976): Steroidogenic capacities of the different compartments of the human ovary. In: The Endocrine Function of The Human Ovary, edited by V. H. T. James, M. Serio, and G. Giusti, pp. 37-45. Academic, London. 135. McNatty, K. P., Makris, A., De Grazia, C., Osathanondh, R., and Ryan, K. J. (1980): Steroidogenesis by recombined follicular cells from the human ovary in vitro. J. Clin. Endocrinol. Metab., 51 1286— 1292. 136. Dorrington, J. H., Moon, Y. S., and Armstrong, D. T. (1975): Estradiol-17p biosynthesis in cultured granulosa cells from hypophysectomized immature rats; stimulation by follicle-stimulating hor¬ mone. Endocrinology, 97:1328-1331. 137. Gore-Langton, R. E., and Dorrington, J. H. (1981): FSH induction of aromatase in cultured rat granulosa cells measured by a radiometric assay. Mol. Cell. Endocrinol., 22:135-151. 138. Wang, C., Hsueh, A. J. W., and Erickson, G. F. (1982): The role of cyclic AMP in the induction of estrogen and progestin synthesis in cultured granulosa cells. Mol. Cell. Endocrinol., 25:73-83. 139. Mendelson, C. R., Durham, C., Evans, C., and Simpson, E. R. (1985). The induction of aromatase activity in estrogen-producing cells is mediated by the increased synthesis of aromatase cytochrome P^450. 67th Annual Meeting of The Endocrine Society, p. 77 (Abstr. 140. Durham, C. R., Zhu, H., Masters, B. S. S., Simpson, E. R„ and Mendelson, C. R. (1985): Regulation of aromatase activity of rat granulosa cells: induction of synthesis of NADPH-cytochrome P-450 reductase by FSH and dibutyryl cyclic AMP. Mol. Cell. Endocrinol 40:211-219. 141. Dorrington, J. H., and Armstrong, D. T. (1979): Effect of FSH on gonadal functions. Rec. Prog. Horm. Res., 35:301-342. 142. Hillier, S. G., Zeleznik, A. J., and Ross, G. T. (1978): Independ¬ ence of steroidogenic capacity and luteinizing hormone receptor induction in developing granulosa cells. Endocrinology 102 937— 946.

143. Armstrong, D. T., and Dorrington, J. H. (1976): Androgens augment FSH-induced progesterone secretion by cultured rat granulosa cells. Endocrinology, 99:1411-1414. 144. Nimrod, A. (1977): Studies on the synergistic effect of androgen on the stimulation of progestin secretion by FSH in cultured rat granulosa cells: a search for the mechanism of action. Mol. Cell. Endocrinol.,

8:201-211. 145. Nimrod, A., and Lindner, H. R. (1976): A synergistic effect of androgen on the stimulation of progesterone secretion by FSH in cultured rat granulosa cells. Mol. Cell. Endocrinol., 5:315-320. 146. McNatty, K. P., Makris, A., De Grazia, C., Osathanondh, R., and Ryan, K. J. (1979): The production of progesterone, androgens and oestrogens by human granulosa cells in vitro and in vivo. J. Steroid Biochem., 11:775-779. 147. Channing, C. P. (1974): Temporal effects of LH, hCG, FSH and dibutyryl cyclic 3',5'-AMP upon luteinization of rhesus monkey granulosa cells in culture. Endocrinology, 94:1215-1223. 148. Hammond, R. W., Burke, W. H., and Hertelendy, F. (1981): In¬ fluence of follicular maturation of progesterone release in chicken granulosa cells in response to turkey and ovine gonadotropins. Biol. Reprod., 24:1048-1055. 149. Hillensjo, T., Magnusson, C., Svensson, U., and Thelander, H. (1981): Effects of luteinizing hormone and follicle-stimulating hor¬ mone on progesterone synthesis by cultured rat cumulus cells. En¬ docrinology, 108:1920-1924. 150. Wang, C., Hsueh, A. J. W., and Erickson, G. F. (1981): LH stim¬ ulation of estrogen secretion in cultured granulosa cells. Mol. Cell. Endocrinol., 24:17-28. 151. Chang, S. C. S., Jones, J. D., Ellefson, R. D., and Ryan, R. J. (1976): The porcine ovarian follicle. I. Selected chemical analysis of follicular fluid at different developmental stages. Biol. Reprod., 15:321-328. 152. Carr, B. R., MacDonald, P. C., and Simpson, E. R. (1982): The role of lipoproteins in the regulation of progesterone secretion by the human corpus luteum. Fertil. Steril., 38:303-311. 153. Jones, P. B. C., and Hsueh, A. J. W. (1982): Pregnenolone bio¬ synthesis by cultured granulosa cells: modulation by follicle-stimu¬ lating hormone and gonadotropin-releasing hormone. Endocrinology 111:713-721. 154. Toaff, M.E ., Strauss III, J. F., and Hammond, J. M. (1983): Reg¬ ulation of cytochrome P-450Scc in immature porcine granulosa cells by FSH and estradiol. Endocrinology, 112:1156-1158. 155. Boyd, G. S., Arthur, J. R., Beckett, G. J., Mason, J. I., and Trzeciak, W. H. (1975): The role of cholesterol and cytochrome P450 in the cholesterol side-chain cleavage reaction in adrenal cortex and corpora lutea. J. Steroid Biochem., 6:427^136. 156. Robinson, J., Stevenson, P. M., Boyd, G. S., and Armstrong, D. T. (1975): Acute in vivo effects of hCG and LH on ovarian mitochondrial cholesterol utilization. Mol. Cell. Endocrinol 2• 149155. 157. Mori, M., and Marsh, J. M. (1982): The site of luteinizing hormone stimulation of steroidogenesis in mitochondria of the rat corpus lu¬ teum. J. Biol. Chem., 257:6178-6183. 158. Naumoff, P. A., and Stevenson, P. M. (1981): The differential development of mitochondrial cytochrome P-450 and the respiratory cytochromes in rat ovary. Biochim. Biophys. Acta, 673:359-365. 159. Schuler, L. A., Toaff, M. E., and Strauss HI, J. F. (1981): Regulation of ovarian cholesterol metabolism: control of 3-hydroxy-3-methylglutaryl coenzyme A reductase and acyl coenzyme A:cholesterol acyltransferase. Endocrinology, 108:1476-1486. 160. Behrman, H. R., and Armstrong, D. T. (1969): Cholesterol esterase stimulation by luteinizing hormone in luteinized rat ovaries. Endo¬ crinology, 85:474-480. 161. Caffrey, J. L., Fletcher, P. W., Diekman, M. A., O’Callaghan, P. L., and Niswender, G. D. (1979): The activity of ovine luteal cholesterol esterase during several experimental conditions. Biol. Reprod., 21:601-608. 162. Henderson, K. M., Gorban, A. M. S., and Boyd, G. S. (1981): Effect of LH factors regulating ovarian cholesterol metabolism and progesterone synthesis in PMSG-primed immature rats J Reprod Fertil., 61:373-380. 163. Hwang, J., and Menon, J. M. J. (1983): Characterization of low density and high density lipoprotein receptors in the rat corpus luteum and regulation by gonadotropin. J. Biol. Chem., 258:8020-8027.

Follicular Steroidogenesis and Its Control /

164. Jones, P. B. C., Valk, C. A., andHsueh, A. J. W. (1983): Regulation of progestin biosynthetic enzymes in cultured rat granulosa cells: Effects of prolactin, (^-adrenergic agonist, human chorionic gonad¬ otropin and gonadotropin-releasing hormone. Biol. Reprod., 29:572585. 165. Rubin, B. L., Deane, H. W., Hamilton, J. A., and Driks, E. C. (1963): Changes in A5-3(3-hydroxysteroid dehydrogenase activity in the ovaries of maturing rats. Endocrinology, 72:924-930. 166. Koritz, S. B. (1967): On the regulation of pregnenolone synthesis. In: Functions of the Adrenal Cortex, edited by K. McKerns, pp. 2748. Appleton-Century-Crofts, New York. 167. Madej, E. (1980): Effect of exogenous hormones on the activity of A5-3(3-hydroxysteroid dehydrogenase in cultured granulosa cells from proestrous and preovulatory rat ovarian follicles. Acta Histochem., 67:253-260. 168. Neville, A. M., and Engel, L. L. (1968): Steroid A-isomerase of the bovine adrenal gland: kinetics, activation by NAD and attempted solubilization. Endocrinology, 83:864—872. 169. Philpott, J. E., and Peron, F. G. (1971): A microassay procedure for A5-3(3-hydroxysteroid dehydrogenase based on substrate deple¬ tion. Endocrinology, 88:1082-1085. 170. Dorrington, J. H., McKeracher, H. L., Chan, A., and Gore-Langton, R. E. (1984): Luteinizing hormone-releasing hormone independently stimulates cytodifferentiation of granulosa cells. In: Hormonal Con¬ trol of The Hypothalamo-pituitary-gonadal Axis. Biochemical En¬ docrinology Series, edited by K. W. McKerns and Z. Naor, pp. 467478. Plenum, New York. 171. Lischinsky, A., Evans, G., and Armstrong, D. T. (1983): Site of androgen inhibition of FSH-stimulated progesterone production in porcine granulosa cells. Endocrinology, 113:1999-2003. 172. Eckstein, B., and Nimrod, A. (1979): Effect of human chorionic gonadotropin and prolactin on 20a-hydroxysteroid dehydrogenase activity in granulosa cells of immature rat ovary. Endocrinology, 104:711-714. 173. Jones, P. B. C., and Hsueh, A. J. W. (1981): Direct stimulation of ovarian progesterone metabolizing enzyme by gonadotropin-releas¬ ing hormone in cultured granulosa cells. J. Biol. Chem., 256:12481254. 174. Eckstein, B., Raanan, M., Lemer, N., Cohen, S., and Nimrod, A. (1977): The appearance of 20a-hydroxysteroid dehydrogenase activ¬ ity in preovulatory follicles of immature rats treated with pregnant mare serum gonadotropin. J. Steroid Biochem., 8:213-216. 175. Hickman-Smith, D., and Kuhn, N. J. (1976): A proposed sequence of hormones controlling the induction of luteal 20a-hydroxysteroid dehydrogenase and progesterone withdrawal in the late-pregnant rat. Biochem. J., 160:663-670. 176. Loewit, K., and Zambelis, N. (1979): Progesterone and 20a-hydroxysteroid dehydrogenase regulation in the corpus luteum of the pregnant rat. Acta Endocrinol. (Copenh.), 90:176-184. 177. Suzuki, K., and Tamaoki, B. I. (1979): Enzymological studies of rat luteinized ovaries in relation to acute reduction of aromatizable androgen formation and stimulated production of progestins. En¬ docrinology, 104:1317-1323. 178. Wiest, W. G. (1959): Conversion of progesterone to 4-pregn-20aol-3-one by rat ovarian tissue in vitro. J. Biol. Chem., 234:31153121. 179. Wiest, W. G., Kidwell, W. R., and Kirschbaum, T. H. (1963): Induction of rat ovarian 20a-hydroxysteroid dehydrogenase ac¬ tivity by gonadotrophic hormone administration. Steroids, 2.617— 630. 180. Csapo, A. I., and Wiest, W. G. (1969): An examination of the quantitative relationship between progesterone and the maintenance of pregnancy. Endocrinology, 85:735-746. 181. Wiest, W. G. (1968): On the function of 20a-hydroxypregn-4-en-3one during parturition in the rat. Endocrinology, 83:1181-1184. 182. Diaz-Zagoya, J. C., Wiest, W. G., and Arias, F. (1979). 20aHydroxysteroid oxidoreductase activity and 20a-dihydroprogesterone concentration in human placenta before and after parturition. Am. J. Obstet. Gynecol., 133:673-676. 183. Louth, G., Peron, F., and Dorfman, R. I. (1962): Formation of 20ahydroxy-A4-pregnen-3-one and A4-androsten-3,17-dione in rodent ovaries. Endocrinology, 71:983-985. 184. Savard, K., Marsh, J. M., and Rice, B. F. (1965): Gonadotropins and ovarian steroidogenesis. Rec. Prog. Horm. Res., 21.285-356.

373

185. Eckstein, B., and Lemer, N. (1977): Changes in ovarian 5a-steroid reductase and 20a-hydroxysteroid dehydrogenase activity produced by induction of hrst ovulation with gonadotropin. Biochim. Biophys. Acta, 489:143-149. 186. Epstein-Almog, R., and Orly, J. (1985): Inhibition of hormoneinduced steroidogenesis during cell proliferation in serum-free cul¬ tures of rat granulosa cells. Endocrinology, 116:2103-2112. 187. Short, R. V. (1962): Steroids in the follicular fluid and the corpus luteum of the mare: A “two-cell type” theory of ovarian steroid synthesis. J. Endocrinol., 24:59-63. 188. Bjersing, L., and Carstensen, H. (1967): Biosynthesis of steroids by granulosa cells of the porcine ovary in vitro. J. Reprod. Fertil., 14:101-111. 189. Younglai, E. V., and Short, R. V. (1970): Pathways of steroid biosynthesis in the intact Graafian follicle of mares in oestrus. J. Endocrinol., 47:321-331. 190. Fortune, J. E., and Armstrong, D. T. (1978): Hormonal control of 173-estradiol biosynthesis in proestrous rat follicles: estradiol pro¬ duction by isolated theca versus granulosa cells. Endocrinology, 102:227-235. 191. Fowler, R. E., Fox, N. L., Edwards, R. G., Walters, D. E., and Steptoe, P. C. (1978): Steroidogenesis by cultured granulosa cells aspirated from human follicles using pregnenolone and androgen as precursors. J. Endocrinol., 77:171-183. 192. Johnson, D. C., and Hoversland, R. C. (1983): Oestradiol synthesis by granulosa cells from immature rats treated with pregnant mare’s serum gonadotrophin. Acta Endocrinol. (Copenh.), 104:74-79. 193. Bjersing, L., and Carstensen, H. (1964): The role of the granulosa in the biosynthesis of ovarian steroids. Biochim. Biophys. Acta, 86:637639. 194. Moor, R. M. (1977): Sites of steroid production in ovine Graafian follicles in culture. J. Endocrinol., 73:143-150. 195. Nimrod, A., Rosenfield, R. L., and Otto, P. (1980): Relationship of androgen action to androgen metabolism in isolated rat granulosa cells. J. Steroid Biochem., 13:1015-1019. 196. Moon, Y. S., and Duleba, A. J. (1982): Comparative studies of androgen metabolism in theca and granulosa cells of human follicles in vitro. Steroids, 39:419-430. 197. Terakawa, N., Kondo, K., Aono, T., Kurachi, K., and Matsumoto, K. (1978): Hormonal regulation of 4-ene-5a-reductase activity in prepubertal rat ovaries. J. Steroid Biochem., 9:307-311. 198. Allen, E., and Doisy, E. A. (1927): Ovarian and placental hormones. Physiol. Rev., 7:600-650. 199. Mossman, H. W. (1937): The thecal gland and its relation to the reproductive cycle. A study of the cyclic changes in the ovary of the pocket gopher, Geomys Bursarius (Shaw). Am. J. Anat., 61:289319. 200. Comer, G. W. (1983): The sites of formation of estrogenic substances in the animal body. Physiol. Rev., 18:154-172. 201. Hisaw, F. L. (1947): Development of the Graafian follicle and ovu¬ lation. Physiol. Rev., 27:95-119. 202. Allen, E. (1941): Glandular physiology and therapy. JAMA, 116:405413. 203. Falck, B. (1959): Site of production of oestrogen in rat ovary as studied in micro-transplants. Acta Physiol. Scand. Suppl. 47, 163:1101. 204. Greep, R. O., van Dyke, H. B., and Chow, B. F. (1942): Gonad¬ otropins of the swine pituitary. I. Various biological effects of puri¬ fied thylakentrin (FSH) and pure metakentrin (ICSH). Endocrinol¬ ogy, 30:635-649. 205. Hollander, N., and Hollander, V. (1958): The effect of folliclestimulating hormone on the biosynthesis in vitro of estradiol-173 from acetate-l-C14 and testosterone-4-C14. J. Biol. Chem., 233:10971099. 206. Mills, T. M., Davies, P. J. A., and Savard, K. (1971): Stimulation of estrogen synthesis in rabbit follicles by luteinizing hormone. En¬ docrinology, 88:857-862. 207. Moon, Y. S., Dorrington, J. H., and Armstrong, D. T. (1975): Stimulatory action of follicle-stimulating hormone on estradiol-173 secretion by hypophysectomized rat ovaries in organ culture. En¬ docrinology, 97:244-247. 208. Dorrington, J. H., and Armstrong, D. T. (1975): Follicle-stimulating hormone stimulates estradiol-173 synthesis in cultured Sertoli cells. Proc. Natl. Acad. Sci. USA, 72:2677-2681.

374

/ Chapter 10

209. Armstrong, D. T., and Papkoff, H. (1976): Stimulation of aromatization of exogenous and endogenous androgens in ovaries of hypophysectomized rats in vivo by follicle-stimulating hormone. En¬ docrinology, 99:1144-1151. 210. Baird, D. T. (1977): Evidence in vivo for the two-cell hypothesis of ovarian estrogen synthesis by the sheep Graafian follicle. J. Reprod. Fertil., 50:183-185. 211. Fowler, R. E., Fox, N. L., Edwards, R. G., and Steptoe, P. C. (1978): Steroid production from 17a-hydroxypregnenolone and dehydroepiandrosterone by human granulosa cells in vitro. J. Reprod. Fertil., 54:109-117. 212. Fortune, J. E., and Armstrong, D. T. (1977): Androgen production by theca and granulosa isolated from proestrous rat follicles. En¬ docrinology, 100:1341-1347. 213. Short, R. V. (1962): Steroids present in the follicular fluid of the cow. J. Endocrinol., 23:401-411. 214. Fortune, J. E. (1981): Bovine theca and granulosa cells interact to promote androgen and progestin production. Biol. Reprod. (Suppl. 1), 24:39A (abstr. 33). 215. Lischinsky, A., and Armstrong, D. T. (1983): Granulosa cell stim¬ ulation of thecal androgen synthesis. Can. J. Physiol. Pharmacol., 61:472-477. 216. Pencharz, R. I. (1940): Effect of estrogens and androgens alone and in combination with chorionic gonadotropin on the ovary of the hypophysectomized rat. Science, 91:554-555. 217. Williams, P. C. (1940): Effect of stilbestrol on the ovaries of hy¬ pophysectomized rats. Nature, 145:388-389. 218. Simpson, M. E., Evans, H. M., Fraenkel-Conrat, H. L., and Li, C. H. (1941): Synergism of estrogens with pituitary gonadotropins in hypophysectomized rats. Endocrinology, 28:37^11. 219. Williams, P. C. (1944): Ovarian stimulation by oestrogens: effects in immature hypophysectomized rats. Proc. R. Soc. London [Biol.], 132:189-199. 220. Payne, R. W., and Hellbaum, A. A. (1955): The effect of estrogens on the ovary of the hypophysectomized rat. Endocrinology, 57:193199. 221. Payne, R. W., and Runser, R. H. (1958): The influence of estrogen and androgen on the ovarian response of hypophysectomized im¬ mature rats to gonadotropins. Endocrinology, 62:313-321. 222. Bradbury, J. T. (1961): Direct action of estrogen on the ovary of the immature rat. Endocrinology, 68:112-120. 223. Smith, B. D. (1961): The effect of diethylstilbestrol on the immature rat ovary. Endocrinology, 69:238-245. 224. Reiter, E. O., Goldenberg, R. L., Vaitukaitis, J. L., and Ross, G. T. (1972): Evidence for a role of estrogen in ovarian augmentation reaction. Endocrinology, 91:1518-1522. 225. Harman, S. M., Louvet, J. P., and Ross, G. T. (1975): Interaction of estrogen and gonadotrophins on follicular atresia. Endocrinology, 96:1145-1152. 226. Saiduddui, S. (1971): 3H-estradiol uptake by the rat ovary. Proc. Soc. Exp. Biol. Med., 138:651-660. 227. Saiduddui, S., and Milo Jr., G. E. (1974): Effect of hypophysectomy and pretreatment on uptake and retention of estradiol by the ovary. Proc. Soc. Exp. Biol. Med., 146:513-517. 228. Stumpf, W. E. (1969): Nuclear concentration of 3H-estradiol in target issues. Dry-mount autoradiography of vagina, oviduct, ovary, testis, mammary tumor, liver and adrenal. Endocrinology, 85:31-37. 229. Richards, J. S. (1975): Estradiol receptor content of rat granulosa cells during follicular development: modification by estradiol and gonadotropins. Endocrinology, 97:1174-1184. 230. Richards, J. S. (1974): Estradiol binding to rat corpora lutea during pregnancy. Endocrinology, 95:1046-1053. 231. Saidudduin, S., and Zassenhaus, H. P. (1977): Estradiol-173 recep¬ tors in the immature rat ovary. Steroids, 29:197-213. 232. Leung, P. C. K., Goff, A. K., Kennedy, T. G., and Armstrong, D. T. (1978): An intraovarian inhibitory action of estrogen and an¬ drogen production in vivo. Biol. Reprod., 19:641-647. 233. Leung, P. C. K., and Armstrong, D. T. (1979): Estrogen treatment of immature rats inhibits ovarian androgen production in vitro. En¬ docrinology, 104:1411-1417. 234. Leung, P. C. K., and Armstrong, D. T. (1980): Further evidence in support of a short-loop feedback action of estrogen on ovarian an¬ drogen production. Life Sci., 27:415-420.

235. Hunter, M. G., and Armstrong, D. T. (1986): Estrogens inhibit steroid production by dispersed porcine thecal cells. Biol. Reprod. (Suppl. 1), 34:196 (abstr. 293). 236. Magoffin, D. A., and Erickson, G. F. (1981): Mechanism by which 173-estradiol inhibits ovarian androgen production in the rat. En¬ docrinology, 108:962-969. 237. Magoffin, D. A., and Erickson, G. F. (1982): Direct inhibitory effects of estrogen on LH-stimulated androgen synthesis by ovarian cells cultured in defined medium. Mol. Cell. Endocrinol., 28:81— 89. 238. Eckstein, B., and Nimrod, A. (1977): Properties of microsomal A4-3-ketosteroid 5a-reductase in immature rat ovary. Biochim. Biophys. Acta, 499:1-9. 239. Armstrong, D. T. (1979): Alterations of progesterone metabolism in immature rat ovaries by luteinizing hormone. Biol. Reprod., 21:10251033. 240. Evans, G., Leung, P. C. K., Brodie, A. M. H., and Armstrong, D. T. (1981): Effect of an aromatase inhibitor (4-acetoxy-4-androstene-3,17-dione) on the stimulatory action of luteinizing hormone on estradiol-17p synthesis by rat preovulatory follicles in vitro. Biol. Reprod., 25:290-294. 241. Armstrong, D. T., Dorrington, J. H., and Robinson, J. (1976): „ Effects of indomethacin and aminoglutethimide phosphate in vivo on luteinizing-hormone-induced alterations on cyclic adenosine monophosphate, prostaglandin F, and steroid levels in preovulatory rat ovaries. Can. J. Biochem., 54:796-802. 242. Bahr, J. M. (1978): Simultaneous measurement of steroids in fol¬ licular fluid and ovarian venous blood in the rabbit. Biol. Reprod , 18:193-197. 243. Goff, A. K., and Henderson, K. M. (1979): Changes in follicular fluid and serum concentrations of steroids in PMS-treated immature rats following LH administration. Biol. Reprod., 20:1153-1157. 244. Smith, M. S., Freeman, M. E., and Neill, J. D. (1975): The control of progesterone secretion during the estrous cycle and early pseudo¬ pregnancy in the rat: prolactin gonadotropin and steroid levels as¬ sociated with rescue of the corpus luteum of pseudopregnancy. En¬ docrinology, 96:219-226. 245. Kalra, S. P., and Kalra, P. S. (1974): Temporal interrelationships among circulating levels of estradiol, progesterone and LH during the rat estrous cycle: effects of exogenous progesterone. Endocri¬ nology, 95:1711-1718. 246. Saidapur, S. K., and Greenwald, G. S. (1979): Regulation of 1713estradiol synthesis in the proestrous hamster: role of progesterone and luteinizing hormone. Endocrinology, 105:1432-1439. 247. Fortune, J. E., and Hansel, W. (1979): The effects of 173-estradiol on progesterone secretion by bovine theca and granulosa cells. En¬ docrinology, 104:1834-1838. 248. Engels, J. A., Friedlander, R. L., and Eik-Nes, K. B. (1968): An effect in vivo of clomiphene on the rate of conversion of androstenedione-C to estrone-C14 and estradiol-C14 by the canine ovary. Me¬ tabolism, 17:189-198. 249. Zhuang, L.-Z., Adashi, E. Y., and Hsueh, A. J. W. (1982): Direct enhancement of gonadotropin-stimulated ovarian estrogen biosyn¬ thesis by estrogen and clomiphene citrate. Endocrinology, 110 2219-

2221. 250. Adashi, E. Y., and Hsueh, A. J. W. (1982): Estrogens augment the stimulation of ovarian aromatase activity by follicle-stimulating hor¬ mone in cultured rat granulosa cells. J. Biol. Chem., 257:60776083. 251. Welsh Jr., T. H., Jia, X.-C., Jones, P. B. C., Zhuang, L.-Z., and Hsueh, A. J. W. (1984): Disparate effects of triphenylethylene antiestrogens on estrogen and progestin biosyntheses by cultured rat granulosa cells. Endocrinology, 115:1275-1282. 252. Daniel, S. A. J., and Armstrong, D. T. (1983): Involvement of estrogens in the regulation of granulosa cell aromatase activity. Can J. Physiol. Pharmacol., 61:507-511. 253. Thanki, K. H., and Channing, C. P. (1976): Influence of serum, estrogen, and gonadotropins upon growth and progesterone secretion by cultures of granulosa cells from small porcine follicles. Endocr. Res. Commun., 3:319-333. 254. Thanki, K. H., and Channing, C. P. (1978): Effects of folliclestimulating hormone and estradiol upon progesterone secretion by porcine granulosa cells in tissue culture. Endocrinology, 103:74-80.

Follicular Steroidogenesis and Its Control / 255. Schomberg, D. W., Stouffer, R. L., and Tyrey, L. (1976): Modu¬ lation of progestin secretion in ovarian cells by 17(3-hydroxy-5-androstan-3-one (dihydrotestosterone): a direct demonstration in monolayer culture. Biochem. Biophys. Res. Commun., 68:77-81. 256. Haney, A. F., and Schomberg, D. W. (1978): Steroidal modulation of progesterone secretion by granulosa cells from large porcine fol¬ licles: a role for androgens and estrogens in controlling steroidogen¬ esis. Biol. Reprod., 19:242-248. 257. Hillier, S. G., Knazek, R. A., and Ross, G. T. (1977): Androgenic stimulation of progesterone production by granulosa cells from preantral ovarian follicles: further in vitro studies using replicate cell cul¬ tures. Endocrinology, 100:1539-1549. 258. Bieszczad, R. R., McClintock, J. S., Pepe, G. J., and Dimino, M. J. (1982): Progesterone secretion by granulosa cells from different sized follicles of human ovaries after short term incubation. J. Clin. Endocrinol. Metab., 55:181-184. 259. Veldhuis, J. D., Klase, P. A., Sandon, B. A., and Kolp, L. A. (1983): Progesterone secretion by highly differentiated human gran¬ ulosa cells isolated from preovulatory Graafian follicles, induced by endogenous gonadotropins and human chorionic gonadotropin. J. Clin. Endocrinol. Metab., 57:287-291. 260. Shemesh, M., and Ailenberg, M. (1977): The effect of androstenedione on progesterone accumulation in cultures of bovine ovarian follicles. Biol. Reprod., 17:499-505. 261. Veldhuis, J. D. (1985): Bipotential actions of estrogens on proges¬ terone biosynthesis by ovarian cells. II. Relation of estradiol’s stim¬ ulatory actions to cholesterol and progestin metabolism in cultured swine granulosa cells. Endocrinology, 117:1076-1083. 262. Leung, P. C. K., and Armstrong, D. T. (1979): A mechanism for the intraovarian inhibitory action of estrogen on androgen production. Biol. Reprod., 21:1035-1042. 263. Goldenberg, R. L., Bridson, W. E., and Kohler, P. O. (1972): Estrogen stimulation of progesterone synthesis by porcine granulosa cells in culture. Biochem. Biophys. Res. Commun., 48:101-107. 264. Bernard, J. (1975): Effect of follicular fluid and oestradiol on the luteinization of rat granulosa cells in vitro. J. Reprod. Fertil., 45:453460. 265. Welsh Jr., T. H., Zhuang, L.-Z., and Hsueh, A. J. W. (1983): Estrogen augmentation of gonadotropin-stimulated progestin biosyn¬ thesis in cultured rat granulosa cells. Endocrinology, 112:1916-1924. 266. Hudson, K. E., and Hillier, S. G. (1985): Catechol estradiol control of FSH-stimulated granulosa cell steroidogenesis. J. Endocrinol., 106:R1-R4. 267. Khan, M. I., and Gibori, G. (1984): Catechol estrogens and their role in luteal steroidogenesis. Biol. Reprod., 30 (Suppl. 1): 127 (abstr. 194). 268. Zeleznik, A. J., Hillier, S. G., and Ross, G. T. (1979): Folliclestimulating hormone-induced follicular development: an examination of the role of androgens. Biol. Reprod., 21:673-681. 269. Neumann, F., von Berswordt-Wallrabe, R., Eiger, W., Steinbeck, K., Hann, J. D., and Kramer, M. (1970): Aspects of androgendependent events as studied by antiandrogens. Recent Prog. Horm. Res., 26:337-410. 270. Lyon, M. F., and Glenister, P. H. (1974): Evidence from Tfm/O that androgen is inessential for reproduction in female mice. Nature, 247:366-367. 271. Schreiber, J. R., Reid, R., and Ross, G. T. (1976): A receptor-like testosterone binding protein in ovaries from estrogen-stimulated hypophysectomized immature female rats. Endocrinology, 98:1206— 1213. 272. Schreiber, J. R., and Ross, G. T. (1976): Further characterization of rat ovarian testosterone receptor with evidence for nuclear trans¬ location. Endocrinology, 99:590-596. 273. Campo, S. M., Carson, R. S„ and Findlay, J. K. (1984): Distribution and characterisation of specific androgen-binding sites within the ovine follicle. 15th Annual Conference of the Australian Society for Reproductive Biology, Canberra, p. 27 (abstr.). 274. Milwidsky, A., Younes, M. A., Besch, N. F., Besch, P. K., and Kaufman, R. H. (1980): Receptor-like binding proteins for testos¬ terone and progesterone in the human ovary. Am. J. Obstet. Gyne¬ col., 138:93-98. 275. McNatty, K. P., Moore Smith, D., Makris, A., Osathanondh, R., and Ryan, K. J. (1979): The microenvironment of the human antral

276.

277.

278.

279.

280.

281. 282.

283.

284.

285.

286.

287.

288.

289.

290.

291.

292.

293.

294.

295.

375

follicle: interrelationships among the steroid levels in antral fluid, the population of granulosa cells, and the status of the oocyte in vivo and in vitro. J. Clin. Endocrinol. Metab., 49:851-860. Carson, R. S., Findlay, J. K., Clarke, I. J., and Burger, H. G. (1981): Estradiol, testosterone, and androstenedione in ovine follic¬ ular fluid during growth and atresia of ovarian follicles. Biol. Re¬ prod., 24:105-113. Hillier, S. G., and Ross, G. T. (1979): Effects of exogenous tes¬ tosterone on ovarian weight, follicular morphology and intraovarian progesterone concentration in estrogen-primed hypophysectomized immature female rats. Biol. Reprod., 20:261-268. Louvet, J. P., Harman, S. M., Schreiber, J. R., and Ross, G. T. (1975): Evidence for a role of androgens in follicular maturation. Endocrinology, 97:366-372. Bagnell, C. A., Mills, T. M., Costoff, A., andMahesh, V. B. (1982): A model for the study of androgen effects of follicular atresia and ovulation. Biol. Reprod., 27:903-914. Opavsky, M. A., and Armstrong, D. T. (1985): The effectiveness of FSH in inducing superovulation is influenced by LH. Biol. Re¬ prod., 32 (Suppl. 1 ):71 (abstr. 67). Ohno, S., Christian, L., and Attardi, B. (1973): Role of testosterone in normal female function. Nature, 243:119-120. Peluso, J. J., Brown, I., and Steger, R. W. (1979): Effects of cyproterone acetate, a potent antiandrogen, on the preovulatory follicle. Biol. Reprod., 21:929-936. Kumari, G. L., Datta, J. K., Das, R. P., and Roy, S. (1978): Evidence for a role of androgens in the growth and maturation of ovarian follicles in rats. Horm. Res., 9:112-120. Mori, T., Suzuki, A., Nishimura, T., and Kambegawa, A. (1977): Evidence for androgen participation in induced ovulation in immature rats. Endocrinology, 101:623-626. Tsafriri, A., and Braw, R. H. (1984): Experimental approaches to atresia in mammals. In: Oxford Reviews of Reproductive Biol¬ ogy, Vol. 6, edited by R. E. Clarke, pp. 226-265. Clarendon Press, Oxford. Moor, R. M., Hay, M. F., and Seamark, R. F. (1975): The sheep ovary: regulation of steoidogenic, haemodynamic and structural changes in the largest follicle and adjacent tissue before ovulation. J. Reprod. Fertil., 45:595-604. Lucky, A. W., Schreiber, J. R., Hillier, S. G., Schulman, J. D., and Ross, G. T. (1977): Progesterone production by cultured preantral rat granulosa cells: stimulation by androgens. Endocrinology, 100:128-133. Corredor, A., and Flickinger, G. L. (1983): Hormonal regulation of progesterone secretion by cultured mouse granulosa cells. Biol. Re¬ prod, 29:1142-1146 . Leung, P. C. K., Goff, A. K., and Armstrong, D. T. (1979): Stim¬ ulatory action of androgen administration in vivo on ovarian re¬ sponsiveness to gonadotropins. Endocrinology, 104:1119-1123. Schomberg, D. W., Williams, R. F., Tyrey, L., and Ulberg, L. C. (1978): Reduction of granulosa cell progesterone secretion in vitro by intraovarian implants of antiandrogen. Endocrinology, 102:984987. Welsh Jr., J. H., Jones, P. B. C., Ruiz de Galaretta, C. M., Fanjul, L. F., and Hsueh, A. J. W. (1982): Androgen regulation of progestin biosynthetic enzymes in FSH-treated rat granulosa cells in vitro. Steroids, 40:691-700. Hillier, S. G., and deZwart, F. A. (1982): Androgen/antiandrogen modulation of cyclic AMP-induced steroidogenesis during granulosa cell differentiation in tissue culture. Mol. Cell. Endocrinol., 28:347361. Goff, A. K., Leung, P. C. K., and Armstrong, D. T. (1979): Stim¬ ulatory action of follicle-stimulating hormone and androgens on the responsiveness of rat granulosa cells to gonadotropins in vitro. En¬ docrinology, 104:1124-1129. Daniel, S. A. J., and Armstrong, D. T. (1984): Site of action of androgens on follicle-stimulating hormone-induced aromatase activ¬ ity in cultured rat granulosa cells. Endocrinology, 114:1975-1982. Knecht, M., Darbon, J. M., Ranta, T., Baukal, A. J., and Catt, K. J. (1984): Estrogens enhance the adenosine 3',5'-monophosphatemediated induction of follicle-stimulating hormone and luteinizing hormone receptors in rat granulosa cells. Endocrinology, 115:41— 49.

376

/ Chapter 10

296. Duleba, A. J., Takahashi, H., and Moon, Y. S. (1983): Androgenic modulation of progesterone metabolism by rat granulosa cells in culture. Steroids, 42:321-330. 297. Moon, Y. S., Duleba, A. J., and Takahashi, H. (1984): Differential actions of LH and androgens on progesterone catabolism by rat gran¬ ulosa cells. Biochem. Biophys. Res. Commun., 119:694-699. 298. Moon, Y. S., Duleba, A. J., Kuir, K. S., and Yuen, B. H. (1985): Alterations of 20a-hydroxysteroid dehydrogenase activity in cultured rat granulosa cells by follicle-stimulating hormone and testosterone. Biol. Reprod., 32:998-1009. 299. Nimrod, A. (1981): On the synergistic action of androgen and FSH on progestin secretion by cultured rat granulosa cells. Cellular and mitochondrial cholesterol metabolism. Mol. Cell. Endocrinol., 21:5162. 300. Schrieber, J. R., Nakamura, K., and Weinstein, D. B. (1983): An¬ drogen and FSH synergistically stimulate rat ovary granulosa cell utilization of rat and human lipoproteins. In: Factors Regulating Ovarian Function, edited by G. Greenwald and P. F. Terranova, pp. 311-315. Raven Press, New York. 301. Evans, G., Lischinsky, A., Daniel, S. A. J., and Armstrong, D. T. (1984): Androgen-inhibition of FSH-stimulated progesterone pro¬ duction by granulosa cells of pre-pubertal pig. Can. J. Physiol. Pharmacol., 62:840-845. 302. Tan, C. H., and Armstrong, D. T. (1984): FSH-stimulated 3|3-HSD activity of porcine granulosa cells: inhibition by androgens. Pro¬ ceedings of the 3rd Joint Meeting of the British Endocrine Societies, Edinburgh, Abstr. 12. 303. Daniel, S. A. J., and Armstrong, D. T. (1980): Enhancement of follicle-stimulating hormone-induced aromatase by androgens in cul¬ tured rat granulosa cells. Endocrinology, 107:1027-1033. 304. Armstrong, D. T., Daniel, S. A. J., Salhanick, A. R., and Sheela Rani, C. S. (1980): Hormonal interactions in regulation of steroid biosynthesis by the ovarian follicle. In: Functional Correlates of Hormone Receptors in Reproduction, edited by V. B. Makesh, T. G. Muldoon, B. B. Saxena, and W. A. Sadler, pp. 245-260. Elsevier/North-Holland, New York. 305. Hillier, S. G., and deZwart, F. A. (1981): Evidence that granulosa cell induction/activation by follicle-stimulating hormone is an an¬ drogen receptor-regulated process in vitro. Endocrinology, 109:13031305. 306. Hillier, S. G., van den Boogaard, A. M. J., Reichert Jr., L. E., and van Hall, E. V. (1980): Alterations in granulosa cell aromatase ac¬ tivity accompanying preovulatory follicular development in the rat ovary with evidence that 5a-reduced C19 steroids inhibit the aro¬ matase reaction in vitro. J. Endocrinol., 84:409-419. 307. Hillier, S. G., van den Boogaard, A. M. J., Reichert Jr., L. E., and van Hall, E. V. (1980): Intraovarian sex steroid hormone interactions and the regulation of follicular maturation: aromatization of andro¬ gens by human granulosa cells in vitro. J. Clin. Endocrinol. Metab., 50:640-647. 308. Smith, O. W., Ofner, P., and Vena, R. L. (1974): In vitro conversion of testosterone-4-l4C to androgens of the 5a-androstane series by normal human ovary. Steroids, 24:311-315. 309. Siiteri, P. K., and Thompson, E. A. (1975): Studies of human pla¬ cental aromatase. J. Steroid Biochem., 6:317-322. 310. Katz, Y., and Armstrong, D. T. (1976): Inhibition of ovarian estra¬ diol- 173 secretion by luteinizing hormone in prepubertal, pregnant mare serum-treated rats. Endocrinology, 99:1442-1447. 311. Saidudduin, S., and Zassenhaus, H. P. (1978): Effect of testosterone and progesterone on the estradiol receptor in the immature rat ovary. Endocrinology, 102:1069-1076. 312. Smith, B. D., and Bradbury, J. T. (1966): Influence of progestins on ovarian responses to estrogen and gonadotrophins in immature rats. Endocrinology, 78:297-301. 313. Jesel, L. (1970): Donnees nouvelles sur le controle exerce par le corps jaune sur la croissance folliculaire au debut du cycle oestral chez le Cobaye. C. R. Acad. Sci., Ser D, 271:1693-1696. 314. Hori, T., Kato, G., and Miyake, T. (1973): Acute effects of ovarian steroids upon follicular growth in the cycling rat. Endocrinol. Jpn , 20:475-482. 315. Buffler, G., and Roser, S. (1974): New data concerning the role played by progesterone in the control of follicular growth in the rat. Acta Endocrinol., 75:569-578.

316. Beattie, C. W., and Corbin, A. (1975): The differential effects of diestrous progesterone administration on proestrous gonadotrophin levels. Endocrinology, 97:885-890. 317. Goodman, A. L., and Hodgen, G. D. (1977): Systemic versus in¬ traovarian progesterone replacement after luteectomy in rhesus mon¬ keys: differential patterns of gonadotropins and follicle growth. J. Clin. Endocrinol. Metab., 45:837-840. 318. Richards, J. S., and Bogovich, K. (1982): Effect of human chorionic gonadotropin and progesterone on follicular development in the im¬ mature rat. Endocrinology, 111:1429-1438. 319. Bogovich, K., Richards, J. S., and Reichert Jr., L. E. (1981): Oblig¬ atory role of LH in the initiation of preovulatory follicular growth in the pregnant rat: specific effects of human chorionic gonadotropin and follicle-stimulating hormone on LH receptors and steroidogenesis in theca, granulosa and luteal cells. Endocrinology, 109:860-867. 320. Schrieber, J. R., and Hsueh, A. J. W. (1979): Progesterone “recep¬ tor” in rat ovary. Endocrinology, 105:915-919. 321. Schreiber, J. R., Hsueh, A. J. W., and Baulieu, E. E. (1983): Binding of the anti-progestin RU-486 to rat ovary steroid receptors. Contra¬ ception, 28:77-85. 322. Schreiber, J. R., and Erickson, G. F. (1979): Progesterone receptor in the rat ovary: further characterization and localization in the gran¬ ulosa cell. Steroids, 34:459-469. 323. Naess, O. (1981): Characterization of cytoplasmic progesterone re¬ ceptors in rat granulosa cells: evidence for nuclear translocation. Acta Endocrinol., 98:288-294. 324. Philibert, D., Ojasoo, T., and Raynaud, J. P. (1977): Properties of the cytoplasmic progestin-binding protein in the rabbit uterus. En¬ docrinology, 101:1850-1861. 325. Pasqualini, J. R., and Nguyen, B. J. (1980): Progesterone receptors in fetal uterus and ovary of the guinea pig. Evolution during fetal development and induction and stimulation in estradiol-primed ani¬ mals. Endocrinology, 106:1160-1165. 326. Jacobs, B. R., and Smith, R. G. (1980): Evidence for a receptorlike protein for progesterone in bovine ovarian cytosol. Endocrinol¬ ogy, 106:1276-1282. 327. Jacobs, B. R., Suchocki, S., and Smith, R. G. (1980): Evidence for human ovarian progesterone receptor. Am. J. Obstet. Gynecol., 138:332-336. 328. Fanjul, L. F., de Galarreta, R., and Hsueh, A. J. W. (1983): Pro¬ gestin augmentation of gonadotrophin-stimulated progesterone pro¬ duction by cultured rat granulosa cells. Endocrinology, 112:405407. 329. Hsueh, A. J. W., Adashi, E. Y., Jones, P. B. C., and Welsh, T. H. (1984): Hormonal regulation of the differentiation of cultured ovarian granulosa cells. Endocr. Rev., 5:76-127. 330. Schreiber, J. R., Nakamura, K., and Erickson, G. F. (1980): Pro¬ gestins inhibit FSH-stimulated steroidogenesis in cultured rat gran¬ ulosa cells. Mol. Cell. Endocrinol., 19:165-173. 331. Schreiber, J. R., Nakamura, K., and Erickson, G. F. (1981): Pro¬ gestins inhibit FSH-stimulated granulosa estrogen production at a post-cAMP site. Mol. Cell. Endocrinol., 21:161-170. 332. Fortune, J. E., and Vincent, S. E. (1983): Progesterone inhibits the induction of aromatase activity in rat granulosa cells in vitro. Biol Reprod., 28:1078-1089. 333. Greenwald, G. S. (1974): Gonadotropin regulation of follicular de¬ velopment. In: Gonadotropins and Gonadal Function, edited by N. R. Moudgal, pp. 205-212. Academic, New York. 334. Mahajan, D. K., and Samuels, L. T. (1975): Inhibition of 17,20(17hydroxyprogesterone)-lyase by progesterone. Steroids, 25:217-228. 335. Brophy, P. J., and Gower, D. B. (1974): Studies on the inhibition by 5a-pregnane-3,20-dione of the biosynthesis of 16-androstenes and dehydroepiandrosterone in boar testis preparations. Biochim Bio¬ phys. Acta, 360:252-259. 336. Rothchild, I. (1981): The regulation of the mammalian corpus luteum. Rec. Prog. Horm. Res., 37:183-298. 337. Crisp, T. M. (1977): Hormone requirements for early maintenance of rat granulosa cell cultures. Endocrinology, 101:1286-1297. 338. Veldhuis, J. D., Klase, P., and Hammond, J. M. (1980): Divergent effects of prolactin upon steroidogenesis by porcine granulosa cells in vitro: influence of cytodifferentiation. Endocrinology, 107:42-46. 339. Advis, J. P., Richards, J. S., and Ojeda, S. R. (1981): Hyperpro¬ lactinemia-induced precocious puberty: studies on the mechanism(s)

Follicular Steroidogenesis and Its Control

by which prolactin enhances ovarian progesterone responsiveness to gonadotropins in prepubertal rats. Endocrinology, 108:1333-1342. 340. Advis, J. P., and Ojeda, S. R. (1978): Hyperprolactinemia-induced precocious puberty in the female rat: ovarian site of action. Endo¬ crinology, 103:924-935. 341. Gibori, G., and Richards, J. S. (1978): Dissociation of two distinct luteotropic effects of prolactin: regulation of luteinizing hormonereceptor content and progesterone secretion during pregnancy. En¬ docrinology, 102:767-774. 342. Veldhuis, J. D., and Hammond, J. M. (1980): Oestrogens regulate divergent effects of prolactin in the ovary. Nature, 284:262-264. 343. Wang, C., Hsueh, A. J. W., and Erickson, G. F. (1980): Prolactin inhibition of estrogen production by cultured rat granulosa cells. Mol. Cell. Endocrinol., 20:135-144. 344. Wang, C., and Chan, V. (1982): Divergent effects of prolactin on estrogen and progesterone production by granulosa cells of rat Graa¬ fian follicles. Endocrinology, 110:1085-1093. 345. Dorrington, J., and Gore-Langton, R. E. (1981): Prolactin inhibits oestrogen synthesis in the ovary. Nature, 290:600-602. 346. Dorrington, J. H., and Gore-Langton, R. E. (1982): Antigonadal action of prolactin: further studies on the mechanism of inhibition of follicle-stimulatory hormone-induced aromatase activity in rat granulosa cell cultures. Endocrinology, 110:1701-1707. 347. Tsai-Morris, C. H., Ghosh, M., Hirshfield, A. N., Wise, P. M., and Brodie, A. M. H. (1983): Inhibition of ovarian aromatase by prolactin in vivo. Biol. Reprod., 29:342-346. 348. Demura, R., Ono, M., Demura, H., Shizume, K., and Oouchi, H. (1982): Prolactin directly inhibits basal as well as gonadotropinstimulated secretion of progesterone and 17(3-estradiol in the human ovary. J. Clin. Endocrinol. Metab., 54:1246-1250. 349. Advis, J. P., Wiener, S. L., and Ojeda, S. R. (1981): Changes in ovarian 3a-androstanediol response to human chorionic gonadotropin during puberty in the rat: modulatory role of prolactin. Endocrinol¬ ogy, 109:223-228. 350. Magoffin, D. A., and Erickson, G. P. (1982): Prolactin inhibition of luteinizing hormone-stimulated androgen synthesis in ovarian in¬ terstitial cells cultured in defined medium: mechanism of actions. Endocrinology, 111:2001 -2007. 351. Fraser, H. M. (1982): Antifertility effects of GnRH. J. Reprod. Fertil., 64:503-515. 352. de Koning, J., van Dieten, J. A. M. J., and van Rees, G. P. (1978): Refractoriness of the pituitary gland after continuous exposure to luteinizing hormone releasing hormone. J. Endocrinol., 79:311-318. 353. Rippel, R. H., Johnson, E. S., and White, W. F. (1974): Effect of consecutive injections of synthetic gonadotropin-releasing hormone on LH release in anestrous and ovariectomized ewes. J. Anim. Sci., 39:907-914. 354. Sandow, J., van Rechenberg, W., Kuhl, H., Baumann, R., Krauss, B., Jerzabek, G., and Killie, S. (1979): Inhibitory control of the pituitary LH secretion by LHRH in male rats. Horm. Res., 11:303317. 355. Fraser, H. M., Laird, N. C., and Blakeley, D. M. (1980): Decreased pituitary responsiveness and inhibition of the luteinizing hormone surge and ovulation in the stump-tailed monkey (Macaca arctoides) by chronic treatment with an agonist of luteinizing hormone-releasing hormone. Endocrinology, 106:452—457. 356. Conti, M., Harwood, J. P., Hsueh, A. J. W., Dufau, M. L., and Catt, K. J. (1976): Gonadotropin-induced loss of hormone receptors and desensitization of luteal adenylate cyclase in the ovary. J. Biol. Chem., 251:7729-7731. 357. Hunzicker-Dunn, M., and Bimbaumer, L. (1976): Adenylate cyclase activities in ovarian tissues. IV. Gonadotropin-induced desensitiza¬ tion of luteal adenylyl cyclase throughout pregnancy and pseudo¬ pregnancy in the rabbit and rat. Endocrinology, 99.211-222. 358. Jonassen, J. A., and Richards, J. S. (1980): Granulosa cell desen¬ sitization: effects of gonadotropins on antral and preantral follicles. Endocrinology, 106:1786-1794. 359. Jonassen, J. A., Bose, K., and Richards, J. S. (1982): Enhancement and desensitization of hormone-responsive adenylate cyclase in gran¬ ulosa cells of preantral and antral ovarian follicles, effects of estradiol and follicle-stimulating hormone. Endocrinology, 111:74-79. 360. Hseuh, A. J. W„ and Jones, P. B. C. (1981): Extrapituitary actions of gonadotropin-releasing hormone. Endocr. Rev., 2:437^161.

/

377

361. Jones, P. B. C., Conn, P. M., Marian, J., and Hsueh, A. J. W. (1980): Binding of gonadotropin-releasing hormone agonist to rat ovarian granulosa cells. Life Sci., 27:2125-2132. 362. Pieper, D. R., Richards, J. S., and Marshall, J. C. (1981): Ovarian gonadotropin-releasing hormone (GnRH) receptors: characterization, distribution, and induction by GnRH. Endocrinology, 108:11481155. 363. Clayton, R. N., Harwood, J. P., and Catt, K. J. (1979): Gonado¬ tropin-releasing hormone analogue binds to luteal cells and inhibits progesterone production. Nature, 282:90-92. 364. Pelletier, G., Sequin, C., Dube, D., and St.-Amaud, R. (1982): Distribution of LHRH receptors in the rat ovary. Biol. Reprod., 26 (Suppl. 1): 151 (abstr. 230). 365. Marian, J., and Conn, P. M. (1983): Subcellular localization of the receptor for gonadotropin-releasing hormone in pituitary and ovarian tissue. Endocrinology, 112:104-112. 366. Ranta, T., Knecht, M., Kody, M., and Catt, K. J. (1983): GnRH receptors in cultured rat granulosa cells: mediation of the inhibitory and stimulatory actions of GnRH. Mol. Cell. Endocrinol., 27:233240. 367. Hazum, E., and Nimrod, A. (1982): Photoaffinity-labeling and flu¬ orescence-distribution studies of gonadotropin-releasing hormone re¬ ceptors in ovarian granulosa cells. Proc. Natl. Acad. Sci. USA, 79:1747-1750. 368. Hazum, E. (1981): Photoaffinity labeling of luteinizing hormone releasing hormone receptor of rat pituitary membrane preparations. Endocrinology, 109:1281-1283. 369. Clayton, R. N., and Huhtaniemi, I. T. (1982): Absence of gonad¬ otropin-releasing hormone receptors in human gonadal tissue. Na¬ ture, 299:56-59. 370. Popkin, R., Bramley, T. A., Currie, A., Shaw, R. W., Baird, D. T., and Fraser, H. M. (1983): Specific binding of luteinizinghormone releasing hormone to human luteal tissue. Biochem. Biophys. Res. Commun., 114:750-756. 371. Bramley, T. A., Menzies, G. S., and Baird, D. T. (1985): Specific binding of gonadotropin-releasing hormone and an agonist to human corpus-luteum homogenates—characterization, properties, and luteal phase levels. J. Clin. Endocrinol. Metab., 61:834-841. 372. Tureck, R. W., Mastroianni Jr., L., Blasco, L., and Strauss III, J. F. (1982): Inhibition of human granulosa cell progesterone secre¬ tion by a gonadotropin-releasing hormone agonist. J. Clin. Endo¬ crinol. Metab., 54:1078-1080. 373. Casper, R. F., Erickson, G. F., Rebar, R. W., and Yen, S. S. C. (1982): The effect of luteinizing hormone releasing factor and its agonist on cultured human granulosa cells. Fertil. Steril., 37:406409. 374. Koos, R. D., Ahren, K. E. B., Janson, P. O., and LeMaire, W. J. (1982): Effect of a GnRH agonist on the rabbit ovary perfused in vitro. 64th Annual Meeting of the Endocrine Society, p. 178 (abstr. 395). 375. Massicotte, J., Veilleux, R., Lavoie, M., and Labrie, F. (1980): An LHRH agonist inhibits FSH-induced cyclic AMP accumulation and steroidogenesis in porcine granulosa cells in culture. Biochem. Biophys. Res. Commun., 94:1362-1366. 376. Takats, A., and Hertelendy, F. (1982): Adenylate cyclase activity of avian granulosa: effect of gonadotropin-releasing hormone. Gen. Comp. Endocrinol., 48:515-524. 377. Hertelendy, F., Linker, F., Asem, E. K., and Raab, B. (1982): Synergistic effect of gonadotropin releasing hormone on LHstimulated progesterone production in granulosa cells of the domestic fowl (Gallus domesticus). Gen. Comp. Endocrinol., 48: 117-122. 378. Bex, F. J., Corbin, A., and France, E. (1982): Resistance of the mouse to the antifertility effects of LHRH agonists. Life Sci., 30:12631269. 379. Asch, R. H., Eddy, C. A., and Schally, A. V. (1981): Lack of luteolytic effect of D-Trp-6-LH-RH in hypophysectomized rhesus monkeys (Macaca mulatto). Biol. Reprod., 25:963-968. 380. Brown, J. L., and Reeves, J. J. (1983): Absence of specific lutein¬ izing hormone releasing hormone receptors in ovine, bovine and porcine ovaries. Biol. Reprod., 29:1179-1182. 381. Asch, R. H., VanSickle, M., Rettori, V., Balmaceda, J. P., Eddy, C. A., Coy, D. H., and Schally, A. V. (1981): Absence of LHRH

378

/

Chapter

10

binding sites in corpora lutea from rhesus monkeys (Macacca mu¬ latto). J. Clin. Endocrinol. Metab., 53:215-217. 382. Rippel, R. H., and Johnson, E. S. (1976): Inhibition of hCG-induced ovarian and uterine weight augmentation in the immature rat by analogs of GnRH. Proc. Soc. Exp. Biol. Med., 152:432—436. 383. Hsueh, A. J. W., and Erickson, G. F. (1979): Extrapituitary action of gonadotropin-releasing hormone: direct inhibition of ovarian ste¬ roidogenesis. Science, 204:845-855. 384. Hsueh, A. J. W., Wang, C., and Erickson, G. F. (1980): Direct inhibitory effect of gonadotropin-releasing hormone upon folliclestimulating hormone induction of luteinizing hormone receptor and aromatase activity in rat granulosa cells. Endocrinology, 106:16971705. 385. Gore-Langton, R. E., Lacroix, M., and Dorrington, J. H. (1981): Differential effects of luteinizing hormone-releasing hormone on fol¬ licle-stimulating hormone-dependent responses in rat granulosa cells and Sertoli cells in vitro. Endocrinology, 108:812-819. 386. Dorrington, J. H., McKeracher, H. L., Chan, A. K., and GoreLangton, R. E. (1983): Hormonal interactions in the control of gran¬ ulosa cell differentiation. J. Steroid Biochem., 19:17-32. 387. Ying, S.-Y., and Guilleman, R. (1979): (DTrp6-Pro9-NEt)-luteinising hormone-releasing factor inhibits follicular development in hypophysectomised rats. Nature, 280:593-595. 388. Jones, P. B. C., and Hsueh, A. J. W. (1982): Regulation of 3(3hydroxysteroid dehydrogenase by gonadotropin-releasing hormone and follicle-stimulating hormone in cultured rat granulosa cells. En¬ docrinology, 110:1663-1671. 389. Jones, P. B. C., and Hsueh, A. J. W. (1981): Regulation of ovarian 20a-hydroxysteroid dehydrogenase by gonadotropin-releasing hor¬ mone and its antagonist in vitro and in vivo. J. Steroid Biochem., 14:1169-1175. 390. Jones, P. B. C., and Hsueh, A. J. W. (1981): Direct effects of gonadotropin-releasing hormone and its antagonist upon ovarian functions stimulated by FSH, prolactin and LH. Biol. Reprod., 24:747759. 391. Jones, P. B. C., and Hsueh, A. J. W. (1981): Regulation of pro¬ gesterone metabolizing enzyme by adrenergic agents, prolactin and prostaglandins in cultured rat ovarian granulosa cells. Endocrinology, 109:1347-1354. 392. Clark, M. R. (1982): Stimulation of progesterone and prostaglandin E accumulation by luteinizing hormone-releasing hormone (LHRH) and LHRH analogs in rat granulosa cells. Endocrinology, 110:146— 152. 393. Sheela Rani, C. S., Ekholm, C., Billig, H., Magnusson, C., and Hillensjo, T. (1983): Biphasic effect of gonadotropin releasing hor¬ mone on progestin secretion by rat granulosa cells. Biol. Reprod., 28:591-597. 394. Popkin, R., Fraser, H. M., and Jonassen, J. (1983): Stimulation of androstenedione and progesterone release by LHRH and LHRH ag¬ onist from isolated rat preovulatory follicles. Mol. Cell. Endocrinol., 29:169-179. 395. Hsueh, A. J. W., and Ling, N. C. (1979): Effect of an antagonistic analog of gonadotropin-releasing hormone upon ovarian granulosa cell function. Life Sci., 25:1223-1230. 396. St.-Amaud, R., Walker, P., Kelly, P. A., and Labrie, F. (1983): Rat ovarian epidermal growth factor receptors: characterization and hormonal regulation. Mol. Cell. Endocrinol., 31:43-52. 397. Clark, M. R., Thibier, C., Marsh, J. M., and LeMaire, W. J. (1980): Stimulation of prostaglandin accumulation by luteinizing hormone¬ releasing hormone (LHRH) and LHRH analogs in rat granulosa cells in vitro. Endocrinology, 107:17—23. 398. Knecht, M., Katz, M. S., and Catt, K. J. (1981): Gonadotropin¬ releasing hormone inhibits cyclic nucleotide accumulation in cultured rat granulosa cells. J. Biol. Chem., 256:34-36. 399. Knecht, M., and Catt, K. J. (1981): Gonadotropin-releasing hor¬ mone: regulation of adenosine 3',5'-monophosphate in ovarian gran¬ ulosa cells. Science, 214:1346-1348. 400. Ranta, T., Baukal, A., Knecht, M., Korhonen, M., and Catt, K. J. (1983): Inhibitory actions of a gonadotropin-releasing hormone ag¬ onist on ovarian follicle-stimulating hormone receptors and adenylate cyclase in vitro. Endocrinology, 112:956-964. 401. Jones. P. B. C., and Hsueh, A. J. W. (1983): Modulation of steroido¬ genic enzymes by gonadotropin-releasing hormone in cultured gran¬ ulosa cells. In: Factors Regulating Ovarian Function, edited by

G. S. Greenwald and P. F. Terranova, pp. 275-279. Raven Press, New York. 402. Ranta, T., Knecht, M., Darbon, J.-M., Baukal, A. J., and Catt, K. J. (1983): Calcium dependence of the inhibitory effect of go¬ nadotropin-releasing hormone on luteinizing hormone-induced cyclic AMP production in rat granulosa cells. Endocrinology, 113:427429. 403. Hsueh, A. J. W., and Jones, P. B. C. (1982): Direct hormonal modulation of ovarian granulosa cell maturation: effects of gonad¬ otropin-releasing hormone. In: Proceedings IVth Regnier de Graaf Symposium: Follicular Maturation and Ovulation, edited by R. Rolland, E. V. Van Hall, S. G. Hillier, K. P. McNatty, and J. Schoemaker, pp. 19-33. Excerpta Medica, Amsterdam. 404. Hsueh, A. J. W., and Jones, P. B. C. (1982): Regulation of ovarian granulosa and luteal cell functions by gonadotropin-releasing hor¬ mone and its antagonist. Adv. Exp. Med. Biol., 147:223-262. 405. Billig, H., Magnusson, C., Ekholm, C., and Hillensjo, T. (1982): Biphasic effect of a GnRH agonist on glycolysis in cultured rat granulosa cells. Biol. Reprod., 26 (Suppl 1): 152A (abstr. 231). 406. Wang, C. (1983): Luteinizing hormone-releasing hormone stimulates plasminogen activator production by rat granulosa cells. Endocri¬ nology, 112:1130-1132. 407. Dorrington, J. H., and Skinner, M. K. (1986): Cytodifferentiation of granulosa cells induced by gonadotropin-releasing hormone pro¬ motes fibronectin secretion. Endocrinology, 118:2065-2071. 408. Naor, Z., and Yavin, E. (1982): Gonadotropin releasing hormone stimulates phospholipid labeling in cultured granulosa cells. Endo¬ crinology, 111:1615-1619. 409. Leung, P. C. K., Raymond, V., and Labrie, F. (1983): Stimulation of phosphatidic acid and phosphatidylinositol labeling in luteal cells by luteinizing hormone-releasing hormone. Endocrinology, 112: 1138-1140. 410. Davis, J. S., Farese, R. V., and Clark, M. R. (1983): Gonadotropin¬ releasing hormone (GnRH) stimulates phosphatidylinositol metabo¬ lism in rat granulosa cells: mechanism of action of GnRH. Proc. Natl. Acad. Sci. USA, 80:2049-2053. 411. Davis, J. S., West, L. A., and Farese, R. V. (1986): Gonadotropinreleasing hormone (GnRH) rapidly stimulates the formation of in¬ ositol phosphates and diacylglycerol in rat granulosa cells: further evidence for the involvement of Ca2+ and protein kinase C in the action of GnRH. Endocrinology, 118:2561-2571. 412. Minegishi, T., and Leung, P. C. K. (1985): Effects of prostaglandins and luteinizing hormone-releasing hormone on phosphatidic acidphosphatidylinositol labelling in rat granulosa cells. Can. J. Physiol. Pharmacol., 63:320-324. 413. Minegishi, T., and Leung, P. C. K. (1985): Luteinizing hormone¬ releasing hormone stimulates arachidonic acid release in rat granulosa cells. Endocrinology, 117:2001-2007. 414. Berridge, M. J. (1981): Phosphatidylinositol hydrolysis: a multifunctioned transducing mechanism. Mol. Cell. Endocrinol., 24115140. 415. Nishizuka, Y. (1984): The role of protein kinase C in cell surface signal transduction and tumour promotion. Nature, 308:693-698. 416. Zilberstein, M., Sakut, H., Eli, Y., and Naor, Z. (1984): Regulation of prostaglandin E, progesterone, and cyclic adenosine monophos¬ phate production in ovarian granulosa cells by luteinizing hormone and gonadotropin-releasing hormone and gonadotropin-releasing hor¬ mone agonist: comparative studies. Endocrinology, 114:2374-2381. 417. Hillensjo, T., and LeMaire, W. J. (1980): Gonadotropin-releasing hormone agonists stimulate meiotic maturation of follicle-enclosed rat oocytes in vitro. Nature, 287:145-146. 418. Banka, C. L., and Erickson, G. F. (1985): Gonadotropin-releasing hormone induces classical meiotic maturation in subpopulations of atretic preantral follicles. Endocrinology, 117:1500-1507. 419. Dekel, N., Sherizly, I., Phillips, D. M., Nimrod, A., Zilberstein, M., and Naor, Z. (1985): Characterization of the maturational changes induced by a GnRH analogue in the rat ovarian follicle. J. Reprod. Fertil., 75:461^(66. 420. Corbin, A., and Bex, F. J. (1981): Luteinizing hormone releasing hormone agonists induce ovulation in hypophysectomized proestrous rats: direct ovarian effect. Life Sci., 29:185-192. 421. Ekholm, C., Hillensjo, T., and Isaksson, O. (1981): Gonadotropinreleasing hormone agonists stimulate oocyte meiosis and ovulation in hypophysectomized rats. Endocrinology, 108:2022-2024.

Follicular Steroidogenesis and Its Control / 379 422. Magoffin, D. A., Reynolds, D. S., and Erickson, G. F. (1981): Direct inhibitory effect of GnRH on androgen secretion by ovarian interstitial cells. Endocrinology, 109:661-663. 423. Magoffin, D. A., and Erickson, G. F. (1982): Mechanism by which GnRH inhibits androgen synthesis directly in ovarian interstitial cells. Mol. Cell. Endocrinol., 27:191-198. 424. Elkind-Hirsch, K., Ravnikar, V., Schift, I., Tulchinsky, D., and Ryan, K. J. (1982): Determinations of endogenous immunoreactive luteinizing hormone-releasing hormone in human plasma. J. Clin. Endocrinol. Metab., 54:602-607. 425. Ying, S.-Y., Ling, N., Bohlen, P., and Guillemin, R. (1981): Gonadocrinins: peptides in ovarian follicular fluid stimulating the secretion of pituitary gonadotropins. Endocrinology, 108:12061215. 426. Esch, F., Ling, N., Ying, S.-Y., and Guillemin, R. (1982): Peptides of gonadal origin involved in reproductive biology. In: Role of Pep¬ tides and Proteins in Control of Reproduction, edited by S. M. McCann and D. S. Dhindsa, Proceedings of National Institute of Health Workshop, Bethesda, MD. 427. Aten, R. F., Williams, T., and Behrman, H. R. (1986): Ovarian gonadotropin-releasing hormone-like protein(s): demonstration and characterization. Endocrinology, 118:961-967. 428. Bimbaumer, L., Shahabi, N., Rivier, J., and Vale, W. (1985): Evi¬ dence for a physiological role of gonadotropin-releasing hormone (GnRH) or GnRH-like material in the ovary. Endocrinology, 116:13671370. 429. Jarrett, R. J. (1965): Effect and mode of action of adrenocorticotrophic hormone upon the reproductive tract of the female mouse. Endocrinology, 76:434^140. 430. Ramaley, J. A. (1973): Role of the adrenal in PMSG-induced ovu¬ lation before puberty: effect of adrenalectomy. Endocrinology, 92:881— 887. 431. Baldwin, D. M., and Sawyer, C. H. (1974): Effect of dexamethasone on LH release and ovulation in the cyclic rat. Endocrinology, 94:13971403. 432. Yaginuma, T., and Kobayashi, T. (1977): Effect of stress, metyrapone and adrenalectomy on compensatory ovarian hypertrophy. En¬ docrinol. Jpn., 24:403—407. 433. Baldwin, D. M. (1979): The effect of glucocorticoids on estrogendependent luteinizing hormone release in the ovariectomized rat and on gonadotropin secretion in the intact female. Endocrinology, 105:120— 128. 434. Cortes-Gallegos, V., Gallegos, A. J., BedollaTovar, N., Cervantes, C., and Parra, A. (1975): Effect of paramethasone acetate on ovarian steroids and gonadotropins. I. Normal menstrual cycle. J. Clin. En¬ docrinol. Metab., 41:215-220. 435. Cunningham, G. R., Goldzieher, J. W., de la Pena, A., and Oliver, M. (1978): The mechanism of ovulation inhibition by triamcinolone acetonide. J. Clin. Endocrinol. Metab., 46:8-14. 436. Hagino, N., Watanabe, M., and Goldzieher, J. W. (1969): Inhibition by adrenocorticotrophin of gonadotrophin-induced ovulation in im¬ mature female rats. Endocrinology, 84:308-314. 437. Schreiber, J. R., Nakamura, K., and Erickson, G. F. (1982): Rat ovary glucocorticoid receptor: identification and characterization. Steroids, 39:569-584. 438. Louvet, J. P., Baislic, M., Bayard, F., and Boulard, C. (1977): Glucocorticoid receptors in rat ovarian granulosa cell cytosol. 59th Annual Meeting of the Endocrine Society, p. 363 (abstr. 601). 439. Hsueh, A. J. W., and Erickson, G. F. (1978): Glucocorticoid in¬ hibition of FSH-induced estrogen production in cultured rat granulosa cells. Steroids, 32:639-648. 440. Schoonmaker, J. N., and Erickson, G. F. (1983): Glucocorticoid modulation of follicle-stimulating hormone-mediated granulosa cell differentiation. Endocrinology, 113:1356-1363. 441. Adashi, E. Y., Jones, P. B. C., and Hsueh, A. J. W. (1981): Syn¬ ergistic effect of glucocorticoids on the stimulation of progesterone production by follicle-stimulating hormone in cultured rat granulosa cells. Endocrinology, 109:1888-1894. 442. Channing, C. P., Tsai, V., and Sachs, D. (1976): Role of insulin, thyroxin and cortisol in luteinization of porcine granulosa cells grown in chemically defined media. Biol. Reprod., 15:235-247. 443. Mahajan, D. K., and Little, A. B. (1978): Specific cortisol binding protein in porcine follicular fluid. Biol. Reprod., 17.834-842.

444. Mahajan, D. K., Billiar, R. B., and Little, A. B. (1980): Isolation of cortisol binding globulin (CBG) from porcine follicular fluid by affinity chromatography. J. Steroid Biochem., 13:67-71. 445. Adashi, E. Y., Resnick, C. E., D’ercole, A. J., Svoboda, M. E., and Van Wyk, J. J. (1985): Insulin-like growth factors as intraovarian regulators of granulosa cell growth and function. Endocr. Rev., 6:400420. 446. Jones, P. B. C., Welsh Jr., T. H., and Hsueh, A. J. W. (1982): Regulation of ovarian progestin production by epidermal growth factor in cultured rat granulosa cells. J. Biol. Chem., 257:1126811273. 447. Hsueh, A. J. W., Welsh Jr., T. H., and Jones, P. B. C. (1981): Inhibition of ovarian and testicular steroidogenesis by epidermal growth factor. Endocrinology, 108:2002-2004. 448. Knecht, M., and Catt, K. J. (1983): Modulation of cAMP-mediated differentiation in ovarian granulosa cells by epidermal growth factor and platelet-derived growth factor. J. Biol. Chem., 258: 2789-2794. 449. Schomberg, D. W., May, J. V., and Mondschein, J. S. (1983): Interactions between hormones and growth factors in the regulation of granulosa cell differentiation in vitro. J. Steroid Biochem., 19:291— 295. 450. Pulley, D. D., and Marrone, B. L. (1986): Inhibitory action of epidermal growth factor on progesterone biosynthesis in hen gran¬ ulosa cells during short term culture: two sites of action. Endocri¬ nology, 118:2284-2291. 451. Erickson, G. F., and Case, E. (1983): Epidermal growth factor an¬ tagonizes ovarian theca-interstitial cytodifferentiation. Mol. Cell. Endocrinol., 31:71-76. 452. Lintem-Moore, S., Moore, G. P. M., Panaretto, B. A., and Rob¬ ertson, D. (1981): Follicular development in the neonatal mouse ovary; effect of epidermal growth factor. Acta Endocrinol. (Copenh.), 96:123—126. 453. Byyny, R. L., Orth, D. N., Cohen, S., and Doyne, E. S. (1974): Epidermal growth factor: effects of androgens and adrenergic agents. Endocrinology, 95:776-782. 454. Savage, C. R., and Cohen, S. (1972): Epidermal growth factor and a new derivative: rapid isolation procedures and biological and chem¬ ical characterization. J. Biol. Chem., 247:7609-7611. 455. Moore Jr., J. B. (1978): Purification and partial characterization of epidermal growth factor isolated from the male rat submaxillary gland. Arch. Biochem. Biophys., 189:1-7. 456. Cohen, S., and Carpenter, G. (1975): Human epidermal growth factor: isolation and chemical and biological properties. Proc. Natl. Acad. Sci. USA, 72:1317-1321. 457. Byyny, R. L., Orth, D. N., Cohen, S., and Island, D. P. (1971): Epidermal growth factor radioimmunoassay: effects of age, androgen and adrenergic agents on EGF storage and release. 53rd Annual Meeting of the Endocrine Society, p. A45 (abstr. 6). 458. Nexo, E., Hollenberg, M. D., Figueroa, A., and Pratt, R. M. (1980): Detection of epidermal growth factor-urogastrone and its receptor during fetal mouse development. Proc. Natl. Acad. Sci. USA, 77:27822785. 459. Spom, M. B., Roberts, A. B., Shull, J. H., Smith, J. M., Ward, J. M., and Sodek, J. (1983): Polypeptide transforming growth factors isolated from bovine sources and used for wound healing in vivo. Science, 219:1329-1331. 460. Twardzik, D. R., Ranchalis, J. E., and Todaro, G. J. (1982): Mouse embryonic transforming growth factors related to those isolated from tumour cells. Cancer Res., 42:590-593. 461. Kudlow, J. E., and Korbin, M. S. (1984): Secretion of epidermal growth factor-like mitogens by cultured cells from bovine anterior pituitary glands. Endocrinology, 115:911-917. 462. Baranao, J. L. S., and Hammond, J. M. (1984): Comparative effects of insulin and insulin-like growth factors on DNA synthesis and differentiation of porcine granulosa cells. Biochem. Biophys. Res. Commun., 124:484-490. 463. Veldhuis, J. D., and Furlanetto, R. W. (1985): Trophic actions of human somatomedin C/insulin-like growth factor I on ovarian cells: in vitro studies with swine granulosa cells. Endocrinology, 116:12351242. 464. Ladenheim, R. G., Tesone, M., and Charreau, E. H. (1984): Insulin action and characterization of insulin receptors in rat luteal cells. Endocrinology, 115:752-756.

380

/ Chapter 10

465. Rein, M. S., and Schomberg, D. W. (1982): Characterization of insulin receptors on porcine granulosa cells. Biol. Reprod., 26 (Suppl. 1): 113A (abstr. 154). 466. Otani, T., Mauro, T., Yukimur, N., and Mochizuki, M. (1985): Effect of insulin on porcine granulosa cells: implications of a possible receptor mediated action. Acta Endocrinol. (Copenh.), 108:104-110. 467. Veldhuis, J. D., Tamura, S., Kolp, L., Furlanetto, R. W., and Lamer, J. (1984): Mechanisms subserving insulin action in the gonad: evidence that insulin induces specific phosphorylation of its immunoprecipitable receptor on ovarian cells. Biochem. Biophys. Res. Commun., 120:144-149. 468. Poretsky, L., Grigorescu, F., and Flier, J. S. (1985): Insulin but not IGF-I receptors are widely distributed in normal human ovary. 67th Annual Meeting of the Endocrine Society, p. 204 (abstr. 814). 469. Hammond, J. M., Knight, A. P., and Rechler, M. M. (1984): So¬ matomedin secretion by porcine granulosa cells: a potential mech¬ anism for regulating ovarian follicular growth. Clin. Res., 32:485A (abstr.). 470. Hammond, J. M., Baranao, J. L. F., Skaleris, D. A., Rechler, M. M., and Knight, A. P. (1984): Somatomedin (Sm) production by cultured porcine granulosa cells (GC). J. Steroid Biochem., 20:1597 (abstr. 128). 471. Hammond, J. M. (1981): Peptide regulators in the ovarian follicle. Aust. J. Biol. Sci., 34:491-504. 472. Hammond, J. M., Yoshida, K., Veldhuis, J. D., Rechler, M. M., and Knight, A. P. (1983): Intrafollicular role of somatomedins: com¬ parison with effect of insulin. In: Factors Regulating Ovarian Func¬ tion, edited by G. S. Greenwald and P. F. Terranova, pp. 197-201. Raven Press, New York. 473. Davoren, J. B., and Hsueh, A. J. W. (1986): Growth hormone increases ovarian levels of immunoreactive somatomedin C/insulin¬ like growth factor I in vivo. Endocrinology, 118:888-898. 474. Diamond, M. P., Webster, B. W., Carr, R. K., Wentz, A. C., and Osteen, K. G. (1985): Human follicular-fluid insulin concentrations. J. Clin. Endocrinol. Metab., 61:990-992. 475. Adashi, E. Y., Resnick, C. E., Svoboda, M. E., and Van Wyk, J. J. (1984): A novel role for somatomedin-C in the cytodifferentiation of the ovarian granulosa cell. Endocrinology, 115:1227-1229. 476. Adashi, E. Y., Resnick, C. E., Svoboda, M. E., and Van Wyk, J. J. (1985): Somatomedin-C synergizes with follicle-stimulating hor¬ mone in the acquisition of progesterone biosynthetic capacity by cultured rat granulosa cells. Endocrinology, 116:2135-2142. 477. Adashi, E. Y., Resnick, C. E., Brodie, A. M. H., Svoboda, M. E., and Van Wyk, J. J. (1985): Somatomedin-C-mediated potentiation of follicle-stimulating hormone-induced aromatase activity of cul¬ tured rat granulosa cells. Endocrinology, 117:2313-2320. 478. Veldhuis, J. D., Kolp, L. A., Toaff, M. E., Strauss III, J. F., and Demers, L. M. (1983): Mechanisms subserving the trophic actions of insulin on ovarian cells: in vitro studies using swine granulosa cells. J. Clin. Invest., 72:1046-1057. 479. Veldhuis, J. D., Nestler, J. E., Strauss III, J. F., and Gwynne, J T. (1986): Insulin regulates low density lipoprotein metabolism by swine granulosa cells. Endocrinology, 118:2242-2253. 480. Jia, X.-C., Kalmijn, J., and Hsueh, A. J. W. (1986): Growth hor¬ mone enhances follicle-stimulating hormone-induced differentiation of cultured rat granulosa cells. Endocrinology, 118:1401-1409. 481. Davoren, J. B., and Hsueh, A. J. W. (1984): Insulin enhances FSHstimulated steroidogenesis by cultured rat granulosa cells. Mol. Cell. Endocrinol., 35:97-105. 482. Veldhuis, J. D., and Kolp, L. A. (1985): Mechanisms subserving insulin’s differentiating actions on progestin biosynthesis by ovarian cells: studies with cultured swine granulosa cells. Endocrinology 116:651-659. 483. Ciancio, M. J., and LaBarbera, A. R. (1984): Insulin stimulates granulosa cells: increased progesterone and cAMP production in vi¬ tro. Am. J. Physiol., 10:E468-E474. 484. Garzo, V. G., and Dorrington, J. H. (1984): Aromatase activity in human granulosa cells during follicular development and the mod¬ ulation by follicle-stimulating hormone and insulin. Am. J. Obstet. Gynecol., 148:657-662. 485. May, J. V., and Schomberg, D. W. (1981): Granulosa cell differ¬ entiation in vitro: effect of insulin on growth and functional integrity. Biol. Reprod., 25:421-431. 486. Burden, H. W. (1972): Adrenergic innervation in ovaries of the rat and guinea pig. Am. J. Anat., 133:455^162.

487. Bahr, J. M., and Ben-Jonathan, N. (1981): Pre-ovulatory depletion of ovarian catecholamine. Endocrinology, 108:1815-1820. 488. Ben-Jonathan, N., Brown, R. H., Laufer, N., Reich, R., and Bahr, J. M. (1982): Norepinephrine in Graafian follicles is depleted by follicle-stimulating hormone. Endocrinology, 110:457-461. 489. Bahr, J. M., and Ben-Jonathan, N. (1985): Elevated catecholamine in porcine follicular fluid before ovulation. Endocrinology, 117:620— 623. 490. Burden, H. W., and Lawrence, I. E. (1977): The effects of de¬ nervation on the localization of A5-3(3-hydroxysteroid dehydro¬ genase activity in the rat ovary during pregnancy. Acta Anat., 97: 286-290. 491. Capps, M. L., Lawrence, I. E., and Burden, H. W. (1978): Ultra¬ structure of the cells of the ovarian interstitial gland in hypophysectomized rats. The effects of stimulation of the ovarian plexus and of denervation. Cell Tissue Res., 193:433^442. 492. Bahr, J., Kao, L., and Nalbandov, A. V. (1974): The role of cate¬ cholamine and nerves in ovulation. Biol. Reprod., 10:273-290. 493. Condon, W. A., and Black, D. L. (1976): Catecholamine-induced stimulation of progesterone by the bovine corpus luteum in vitro. Biol. Reprod., 15:573-578. 494. Jordan III, A. W., Caffrey, J. L., and Niswender, G. D. (1978): Catecholamine-induced stimulation of progesterone and adenosine 3',5'-monophosphate production by dispersed ovine luteal cells. En¬ docrinology, 103:385-392. 495. Kliachko, S., and Zor, U. (1981): Increase in catecholamine-stim¬ ulated cyclic AMP and progesterone synthesis in rat granulosa cells during culture. Mol. Cell Endocrinol., 23:23-32. 496. Adashi, E. Y., and Hsueh, A. J. W. (1981): Stimulation of 3adrenergic responsiveness by follicle-stimulating hormone in rat granulosa cells in vitro and in vivo. Endocrinology, 108:2170-2178. 497. Aguado, L. I., Petrovic, S. L., and Ojeda, S. R. (1982): Ovarian 3-adrenergic receptors during the onset of puberty: characterization, distribution and coupling to steroidogenic responses. Endocrinology, 110:1124-1132. 498. Sheela Rani, C. S., Nordenstrom, K. Norjavaara, E., and Ahren, K. (1983): Development of catecholamine responsiveness in gran¬ ulosa cells from preovulatory rat follicles—dependence on preovu¬ latory luteinizing hormone surge. Biol. Reprod., 28:1021-1031. 499. Dyer, C. A., and Erickson, G. F. (1985): Norepinephrine amplifies human chorionic gonadotropin-stimulated androgen biosynthesis by ovarian theca-interstitial cells. Endocrinology, 116:1645-1652. 500. Burden, H. W., and Lawrence Jr., I. E. (1978): Experimental studies on the acetylcholinesterase-positive nerves in the ovary of the rat Anat. Rec., 190:233-242. 501. Farrar, J. A., Handeberg, G. M., Hartley, M. L., and Pennefather, J. N. (1980): Catecholamine levels in the guinea pig ovary, myo¬ metrium and costo-uterine muscle during the estrous cycle and in the ovary remaining after unilateral ovariectomy. Biol. Reprod., 22:473479. 502. Ojeda, S. R., Costa, M. E., Katz, K. H., and Hersh, L. B. (1985): Evidence for the existence of substance P in the prepubertal rat ovary. I. Biochemical and physiological studies. Biol. Reprod., 33 286295. 503. Dees, W. L., Kozlowski, G. P., Dey, R., and Ojeda, S. R. (1985): Evidence for the existence of substance P in the prepubertal rat ovary. II. Immunocytochemical localization. Biol. Reprod 33471-476. 504. Larson, L. I., Fahrenkrug, J., and Schaffalitsky de Misckadell, O. B. (1977): Vasoactive intestinal polypeptide occurs in nerves of the female genitourinary tract. Science, 197:1374-1375. 505. Davoren, J. B., and Hsueh, A. J. W. (1985): Vasoactive intestinal peptide: a novel stimulator of steroidogenesis by cultured rat gran¬ ulosa cells. Biol. Reprod., 33:37-52. 506. Pedersen, T., and Peters, H. (1968): Proposal for a classification of oocytes and follicles in the mouse ovary. J. Reprod. Fertil., 17:555— 507. Peters, H., Byskov, A. G., and Grinsted, J. (1978): Follicular growth in fetal and prepubertal ovaries in humans and other primates. In: Clinics in Endocrinology and Metabolism. Reproductive Endocri¬ nology, edited by G. T. Ross and M. B. Lipsett, pp. 469^185. Saunders, London. 508. Noumura, T., Weisz, J., and Lloyd, C. W. (1966): In vitro con¬ version of 7-H3 progesterone to androgens by the rat testis during the second half of fetal life. Endocrinology, 78:245-253.

Follicular Steroidogenesis and Its Control / 509. Arai, H. (1920): On the postnatal development of the ovary (albino rat) with special reference to the number of ova. Am. J. Anat., 27:405-462. 510. Beaumont, H., and Mandl, A. M. (1962): A quantitative and cytological study of oogonia and oocytes in the foetal and neonatal rat. Proc. R. Soc. London [Biol.] 155:557-579. 511. Levina, S. E., Gyevai, A., and Horvath, E. (1975): Responsive¬ ness of the ovary to gonadotrophins in pre- and perinatal life: estrogen secretion in tissue and organ cultures. J. Endocrinol., 65: 219-223. 512. Schlegel, R. J., Farias, E., Russo, N. C., Moore, J. R., and Gardner, L. I. (1967): Structural changes in the fetal gonads and gonaducts during maturation of an enzyme, steroid 3(3-ol-dehydrogenase, in the gonads, adrenal cortex and placenta of fetal rats. Endocrinology, 81:565-572. 51.3. Brambell, F. W. R. (1927): The development and morphology of the gonads of the mouse. 1. The morphogenesis of the indifferent gonad and of the ovary. Proc. R. Soc. London [Biol.], 101:391408. 514. Borum, K. (1961): Oogenesis in the mouse. A study of the meiotic prophase. Exp. Cell Res., 24:495-507. 515. Peters, H. (1969): The development of the mouse ovary from birth to maturity. Acta Endocrinol. (Copenh.), 62:98-116. 516. Milewich, L., George, F. W., and Wilson, J. D. (1977): Estrogen formation by the ovary of the rabbit embryo. Endocrinology, 100:187196. 517. Mauleon, P., Bezard, J., and Terqui, M. (1977): Very early and transient 17(3-estradiol secretion by fetal sheep ovary—in vitro study. Ann. Biol. Anim. Biochim. Biophys., 17:339-401. 518. Shemesh, M., Ailenberg, M., Milaguir, F. Ayalon, N., and Hansel, W. (1978): Hormone secretion by cultured bovine pre- and postim¬ plantation gonads. Biol. Reprod., 19:761-767. 519. Erickson, G. F., Challis, J. R. G., and Ryan, K. J. (1974): A developmental study on the capacity of rabbit granulosa cells to respond to trophic hormones and secrete progesterone in vitro. Dev. Biol., 40:208-224. 520. Mauleon, P. (1961): Utilization de la colchicine dans l’etude des divisions goniales de l’ovaire d’embryons de brebis et analyse de quelques resultats. Ann. Biol. Anim. Biochem. Biophys., 1:70-73. 521. Teplitz, R., and Ohno, S. (1963): Postnatal induction of ovogenesis in the rabbit (Oryctolagus cuniculus). Exp. Cell Res., 31:183-189. 522. Peters, H., Levy, E., and Crone, M. (1965): Oogenesis in rabbits. J. Exp. Zool., 158:169-180. 523. Mauleon, P. (1967): Cinetique de l’ovogenese chez les mammiferes. Arch. Anat. Microsc. Morphol. Exp. (Suppl).3/4:125-150. 524. Shemesh, M. (1980): Estradiol-17(3 biosynthesis by the early bovine fetal ovary during the active and refractory phases. Biol. Reprod., 23:577-582. 525. Roberts, J. D., and Warren, J. C. (1964): Steroid biosynthesis in the fetal ovary. Endocrinology, 74:846-852. 526. Goldman, A. A., Yakovac, W. C., and Bongiovanni, A. M. (1966): Development of activity of 3 P-hydroxysteroid dehydrogenase in hu¬ man fetal tissues and in two anencephalic newborns. J. Clin. En¬ docrinol. Metab., 26:14—22. 527. Cavallero, C., and Magrini, U. (1966) Histochemical studies on 3phydroxysteroid dehydrogenase and other enzymes in the steroid-se¬ creting structures of human foetus. In: Second International Congress on Hormonal Steroids, International Congress Ser. 132, edited by L. Martini, F. Fraschini, and M. Motta, pp. 667-674. Excerpta Medica, Amsterdam. 528. Brandau, H., and Lehmann, V. (1970): Histoenzymatische untersuchungen an menschlichen gonaden wahrend der intrauterinen entwicklung. Z. Geburtshilfe Gynaekol., 173:233—249. 529. Singh, R. F., and Carr, D. H. (1966): The anatomy and histology of XO human embryos and fetuses. Anat. Rec., 155:369-383. 530. Morishima, A., and Grumbach, M. M. (1968): The interrelationship of sex chromosome constitution and phenotype in the syndrome of gonadal dysgenesis and its variants. Ann NY Acad. Sci., 155.695715. 531. Weiss, L. (1971): Additional evidence of gradual loss of germ cells in the pathogenesis of streak ovaries in Turner s syndrome. J. Med. Genet., 8:540-544. 532. Lyon, M. F., and Hawker, S. G. (1973): Reproductive lifespan in irradiated and unirradiated chromosomally XO mice. Genet. Res., 21:185-194.

381

533. Burgoyne, P. S., and Baker, T. G. (1981): The XO ovary—devel¬ opment and function. In: Development and Function of Reproductive Organs, International Congress Series No. 559, edited by A. G. Byskov and H. Peters, pp. 122-128. Excerpta Medica, Amsterdam. 534. Vanhems, E., and Bousquet, J. (1971): Influence du misulban sur le developpement de l’ovaire du rat. Ann. Endocrinol. (Paris), 33:119128. 535. Merchant Larios, H. (1976): The role of germ cells in the morpho¬ genesis and cytodifferentiation of the rat ovary. In: Progress in Dif¬ ferentiation Research, edited by N. Muller-Berat, pp. 453^462. NorthHolland, Amsterdam. 536. Beaumont, H. M. (1961): Radiosensitivity of oogonia and oocytes in the foetal rat. Int. J. Rad. Biol., 3:59-72. 537. Reddoch, R. B., Pelletier, R. M., Barbe, G. J., and Armstrong, D. T. (1986): Lack of ovarian responsiveness to gonadotropic hor¬ mones in infantile rats sterilized with Busulfan. Endocrinology, 119:879-886. 538. Edwards, R. G., Fowler, R. E., Gore-Langton, R. E., et al. (1977): Normal and abnormal follicular growth in mouse, rat and human ovaries. J. Reprod. Fertil., 51:237-263. 539. Nakano, R., Mizuno, T., Katayama, K., and Tojo, S. (1975): Growth of ovarian follicles in the absence of gonadotrophins. J. Reprod. Fertil., 45:545-546. 540. Roy, S. K., and Greenwald, G. S. (1985): An enzymatic method for dissociation of intact follicles from the hamster ovary: histological and quantitative aspects. Biol. Reprod., 32:203-215. 541. Brambell, F. W. R. (1928): The development and morphology of the gonads of the mouse. 3. The growth of the follicles. Proc. R. Soc. London [Biol.], 102:258-272. 542. Harrison, R. J., and Weir, B. J. (1977): Structure of the mammalian ovary. In: The Ovary, Vol. 1, edited by S. Zuckerman and B. J. Weir, pp. 113-217, Academic, New York. 543. Presl, J., Pospisil, J., Figarova, V., and Krabec, Z. (1974): Stage dependent changes in binding of iodinated FSH during ovar¬ ian follicle maturation in rats. Endocrinol. Exp. (Bratisl.), 8:291— 298. 544. Nimrod, A., Erickson, G. F., and Ryan, K. J. (1976): A specific FSH receptor in rat granulosa cells: properties of binding in vitro. Endocrinology, 98:56-64. 545. Quattropani, S. L., and Weisz, J. (1973): Conversion of progesterone to estrone and estradiol in vitro by the ovary of the infantile rat in relation to the development of its interstitial tissue. Endocrinology, 53:1269-1276. 546. Peluso, J. J., Steger, R. W., and Hafez, E. S. E. (1976): Devel¬ opment of gonadotropin-binding sites in the immature rat ovary. J. Reprod. Fertil., 47:55-58. 547. Reddoch, R. B., and Armstrong, D. T. (1984): Interactions of a phosphodiesterase inhibitor, 3-isobutyl-1-methyl xanthine, with pros¬ taglandin E2, follicle-stimulating hormone, luteinizing hormone, and dibutyryl cyclic 3',5'-adenosine monophosphate (cAMP) in cAMP and steroid production by neonatal rat ovaries in vitro. Endocrinol¬ ogy, 115:11-18. 548. Funkenstein, B., Nimrod, A., and Lindner, H. R. (1980): The de¬ velopment of steroidogenic capability and responsiveness to gonad¬ otropins in cultured neonatal rat ovaries. Endocrinology, 106:98106. 549. Kolena, J. (1976): Reversal of the unresponsiveness of neonatal rat ovary to LH in cAMP synthesis by estrogen. Horm. Res., 7:152157. 550. Dohler, K. D., and Wuttke, W. (1975): Changes with age in levels of serum gonadotropins, prolactin and gonadal steroids in prepubertal male and female rats. Endocrinology, 97:898-907. 551. Meijs-Roelofs, H. M. A., deGreef, W. J., and Uilenbroek, J. T. J. (1975): Plasma progesterone and its relationship to serum goadotropins in immature female rats. J. Endocrinol., 64:329-334. 552. Ramaley, J. A. (1979): Development of gonadotropin regulation in the prepubertal mammal. Biol. Reprod., 20:1-31. 553. van Wagenen, G., and Simpson, M. E. (1965): Embryology of The Ovary and Testis. Homo Sapiens and Macaca Mulatto. Yale Uni¬ versity Press, New Haven, CT. 554. Gondos, B., and Hobel, C. J. (1973): Interstitial cells in the human fetal ovary. Endocrinology, 93:736-739. 555. George, F. W., and Wilson, J. D. (1978): Conversion of androgen to estrogen by the human fetal ovary. J. Clin. Endocrinol. Metab., 47:550-555.

382

/ Chapter 10

556. Taylor, T., Coutles, J. R. T., and MacNaughton, M. C. (1974): Human foetal synthesis of testosterone from perfused progesterone. J. Endocrinol, 60:321-326. 557. Jungmann, R. A., and Schweppe, J. S. (1968): Biosynthesis of sterols and steroids from acetate l4C by human fetal ovaries. J. Clin. Endocrinol. Metab., 28:1599-1604. 558. Schindler, A. E., and Friedrich, E. (1975): Steroid metabolism of foetal tissues. I. Metabolism of pregnenolone-4-l4C by human foetal ovaries. Endokrinologie, 65:72-79. 559. Bloch, E. (1964): Metabolism of [4-14C]-progesterone by human fetal testis and ovaries Endocrinology, 74:833-845. 560. Wilson, E. A., and Jawad, M. J. (1979): The effects of trophic agents on fetal ovarian steroidogenesis in organ culture. Fertil. Steril., 32:73-79. 561. Payne, A. H., and Jaffe, R. B. (1974): Androgen formation from pregnenolone sulfate by the human fetal ovary. J. Clin. Endocrinol. Metab., 39:300-304. 562. Huhtaniemi, I., and Vihko, R. (1970): Determination of unconju¬ gated and sulfated neutral steroids in fetal blood of early and mid¬ pregnancy. Steroids, 16:197-206. 563. Ellinwood, W. M., McClellan, M. C., Brenner, R. M., and Resko, J. A. (1983): Estradiol synthesis by fetal monkey ovaries correlates with antral follicle formation. Biol. Reprod., 28:505-516. 564. Grob, H. S. (1969): Growth and endocrine function of isolated ovar¬ ian follicles cultivated in vivo. Biol. Reprod., 1:320-323. 565. Nicosia, S. V., Evangelista, I., and Batta, S. K. (1975): Rabbit ovarian follicles. I. Isolation technique and characterization at dif¬ ferent stages of development. Biol. Reprod., 13:423-447. 566. Terranova, P. F., and Garza, F. (1983): Relationship between the preovulatory luteinizing hormone (LH) surge and androstenedione synthesis of preantral follicles in the cyclic hamster: Detection by in vitro responses to LH. Biol. Reprod., 29:630-636. 567. H^yer, P. E., and Anderson, H. (1977): Histochemistry of 33-hydroxysteroid dehydrogenase in rat ovary. Histochemistry, 51:167193. 568. Richards, J. S. (1980): Maturation of ovarian follicles: actions and interactions of pituitary and ovarian hormones on follicular cell dif¬ ferentiation. Physiol. Rev., 60:51-89. 569. McNatty, K. P. (1978): Follicular fluid. In: The Vertebrate Ovary, edited by R. E. Jones, pp. 215-259. Plenum Press, New York. 570. Richards, J. S., Jonasson, J. A., Rolfes, A. I., Kersey, K., and Reichert Jr., L. E. (1979): Adenosine 3',5'-monophosphate, lutein¬ izing hormone receptor, and progesterone during granulosa cell dif¬ ferentiation: effects of estradiol and follicle-stimulating hormone. Endocrinology, 104:765-773. 571. Sheela Rani, C. S., Salhanick, A. R., and Armstrong, D. T. (1981): Follicle-stimulating hormone induction of luteinizing hormone re¬ ceptor in cultured rat granulosa cells: an examination of the need for steroids in the induction process. Endocrinology, 108:1379-1385. 572. Wang, C., Hsueh, A. J. W., and Erickson, G. F. (1979): Induction of functional prolactin receptors by follicle-stimulating hormone in rat granulosa cells in vivo and in vitro. J. Biol. Chem., 254:11330— 11336. 573. Edwards, R. G. (1974): Follicular fluid. J. Reprod. Fertil., 37:189219. 574. Mossman, H. W., and Duke, K. L. (1973): Comparative Morphology of the Mammalian Ovary. University of Wisconsin Press, Madison. 575. Bjersing, L. (1978): Maturation, morphology, and endocrine function of the follicular wall in mammals. In: The Vertebrate Ovary, edited by R. E. Jones, pp. 181-214. Plenum, New York. 576. Baird, D. T., Swanston, I., and Scaramuzzi, R. J. (1976): Pulsatile release of LH and secretion of ovarian steroids in sheep during the luteal phase of the estrous cycle. Endocrinology, 98:1490-1496. 577. Giorgi, E. P., Addis, M., and Columbo, G. (1969): The fate of free and conjugated oestrogens injected into the Graafian follicles of equines. J. Endocrinol., 43:37-50. 578. Martin, B., Rotten, D., Jolivet, A., and Gautray, J. P. (1981): Binding of steroids by proteins in follicular fluid of the human ovary. J. Clin. Endocrinol. Metab., 53:443^447. 579. Cook, B., Hunter, R. H. F., and Kelly, A. S. L. (1977): Steroid¬ binding proteins in follicular fluid and peripheral plasma from pigs, cows and sheep. J. Reprod. Fertil., 51:65-71. 580. Baird, D. T. (1976): Ovarian steroid secretion and metabolism in women. In: The Endocrine Function of the Human Ovary, edited by

V. H. T. James, M. Seiro, and G. Giusti, pp. 125-133. Academic, London. 581. McNatty, K. P., Baird, D. T., Bolton, A., Chambers, P., Corker, C. S., and McLean, H. (1976): Concentration of oestrogens and androgens in human ovarian venous plasma and follicular fluid throughout the menstrual cycle. J. Endocrinol., 71:77-85. 582. McNatty, K. P., Hunter, W. M., McNeilly, A. S., and Sawers, R. S. (1975): Changes in the concentration of pituitary and steroid hormones in the follicular fluid of human Graafian follicles through¬ out the menstrual cycle. J. Endocrinol., 64:555-571. 583. Kemeter, P., Salzer, H., Breitenecker, G., and Friedrich, F. (1975): Progesterone, oestradiol-17(3, and testosterone levels in the follicular fluid of tertiary follicles and Graafian follicles of human ovaries. Acta Endocrinol. (Copenh.), 80:686-704. 584. McNatty, K. P., and Baird, D. T. (1978): Relationship between follicle-stimulating hormone, androstenedione and oestradiol in hu¬ man follicular fluid. J. Endocrinol., 76:527-531. 585. Breitenecker, G., Friedrich, F., and Kemeter, P. (1978): Further investigations on the maturation and degeneration of human ovarian follicles and their oocytes. Fertil. Steril., 29:336-341. 586. McNatty, K. P. (1982): Ovarian follicular development from the onset of luteal regression in humans and sheep. In: Proceedings IVth ~ Regnier de Graaf Symposium: Follicular Maturation and Ovulation, edited by R. Rolland, E. V. van Hall, S. G. Hillier, K. P. McNatty, and J. Schoemaker, pp. 1-18. Excerpta Medica, Amsterdam. 587. Hillier, S. G., van Hall, E. V., van den Boogaard, A. J. M., de Zwart, F. A., and Keyzer, R. (1982): Activation and modulation of the granulosa cell aromatase system: experimental studies with rat and human ovaries. In: Proceedings IVth Regnier de Graaf Sym¬ posium: Follicular Maturation and Ovulation, edited by R. Rolland, E. V. van Hall, S. G. Hillier, K. P. McNatty, and J. Schoemaker, pp. 51-70. Excerpta Medica, Amsterdam. 588. Bomsel-Helmreich, O., Gougeon, A., Thebault, A., et al. (1979): Healthy and atretic human follicles in the preovulatory phase: dif¬ ferences in evolution of follicular morphology and steroid content of follicular fluid. J. Clin. Endocrinol. Metab., 48:686—694. 589. Westergaard, L., McNatty, K. P., Christensen, I., Larsen, J. K., and Byskov, A. G. (1982): Flow cytometric deoxyribonucleic acid analysis of granulosa cells aspirated from human ovarian follicles. A new method to distinguish healthy and atretic ovarian follicles. J. Clin. Endocrinol. Metab., 55:693-698. 590. McNatty, K. P., Hillier, S. G., van den Boogaard, A. M. J., Trimbos-Kemper, T. C. M., Reichert Jr., L. E., and van Hall, E. V. (1983): Follicular development during the luteal phase of the human menstrual cycle. J. Clin. Endocrinol. Metab., 56:1022-1031. 591. McNatty, K. M., Gibb, M., Dobson, C., Thurley, D. C., and Find¬ lay, J. K. (1981): Changes in the concentration of gonadotrophic and steroidal hormones in the antral fluid of ovarian follicles throughout the estrous cycle of the sheep. Aust. J. Biol. Sci., 34:67-80. 592. Eiler, H., and Nalbandov, A. V. (1977): Sex steroids in follicular fluid and blood plasma during the estrous cycle of pigs. Endocri¬ nology, 100:331-338. 593. Patwardhan, V. V., and Lanthier, A. (1976): Effect of an ovulatory dose of luteinizing hormone on the concentration of oestrone, oes¬ tradiol and progesterone in the rabbit ovarian follicles. Acta Endo¬ crinol. (Copenh.), 82:792-800. 594. Fujii, T., Hoover, D. J., and Channing, C. P. (1983): Changes in inhibin activity, and progesterone, oestrogen and androstenedione concentrations in rat follicular fluid throughout the oestrous cycle. J. Reprod. Fertil., 69:307-314. 595. Moor, R. M., Hay, M. F., Dott, H. M., and Cran, D. G. (1978): Macroscopic identification and steroidogenic function of atretic fol¬ licles in sheep. J. Endocrinol., 77:309-318. 596. Tsuji, K., Sowa, M., and Nakano, R. (1983): Relationship among the status of the human oocyte, the 173-estradiol concentration in the antral fluid and the follicular size. Endocrinol. Jpn 30 251254. 597. Baird, D. T., and Fraser, I. S. (1975): Concentration of oestrone and oestradiol-173 in follicular fluid and ovarian venous blood of women. Clin. Endocrinol., 4:259-266. 598. Baird, D. T. (1983): Factors regulating the growth of the preovulatory follicle in the sheep and human. J. Reprod. Fertil., 69:343-352. 599. Smith, O. W. (1960): Estrogens in the ovarian fluids of normally menstruating women. Endocrinology, 67:698-707.

Follicular Steroidogenesis and Its Control /

600. Short, R. V., and London, D. R. (1961): Defective biosynthesis of ovarian steroids in the Stein-Leventhal syndrome. Br. Med. J., 1:1764— 1727. 601. deJong, F. H., Baird, D. T., and van der Molen, H. J. (1974): Ovarian secretion rates of oestrogens, androgens and progesterone in normal women and in women with persistent ovarian follicles. Acta Endocrinol. (Copenh.), 77:575-587. 602. Sanyal, M. K., Berger, M. J., Thompson, I. E., Taymor, M. L., and Home Jr., H. W. (1974): Development of Graafian follicles in adult human ovary. I. Correlation of estrogen and progesterone con¬ centration in antral fluid with growth of follicles. J. Clin. Endocrinol. Metab., 38:828-835. 603. Short, R. V. (1964): Steroid concentrations in the fluid from nor¬ mal and polycystic (Stein-Leventhal) ovaries. In: Proceedings of the Second International Congress of Endocrinology. Interna¬ tional Congress Ser. No. 83, pp. 940-943. Excerpta Medica, Amsterdam. 604. Edwards, R. G., Steptoe, P. C., Abraham, G. E., Walters, E., Purdy, J. M., and Fotherby, K. (1972): Steroid assays and preovulatory follicular development in human ovaries primed with gonadotro¬ phins. Lancet, 2:611-615. 605. Knudsen, O., and Velle, W. (1961): Ovarian oestrogen levels in the nonpregnant mare: relationship to histological appearance of the uterus and to clinical status. J. Reprod. Eertil., 2:130—137. 606. Short, R. V. (1962): Steroid concentrations in normal follicular fluid and ovarian cyst fluid from cows. J. Reprod. Fertil., 4:27—45. 607. Channing, C. P., and Coudert, S. P. (1976): Contribution of gran¬ ulosa cells and follicular fluid to ovarian estrogen secretion in the rhesus monkey in vivo. Endocrinology, 98:590-597. 608. Aedo, A. R., Pedersen, P. H., Pedersen, S. C., and Diczfalusy, E. (1980): Ovarian steroid secretion in normally menstruating women. I. The contribution of the developing follicle. Acta Endocrinol. (Co¬ penh.), 95:212-221. 609. Uilenbroek, J. T. J., van der Schoot, P., den Besten, D., and Woutersen, P. J. A. (1982): Control of steroidogenesis during growth and early atresia of preovulatory rat follicles. In: Proceedings of the IVth Regnier De Graaf Symposium. Follicular Maturation and Ovu¬ lation, edited by R. Rolland, E. V. van Hall, S. G. Hillier, K. P. McNatty, and J. Schoemaker, pp. 71-82. Excerpta Medica, Am¬ sterdam. 610. diZerega, G. S., Marrs, R. P., Roche, P. C., Campeau, J. D., and Kling, O. R. (1983): Identification of proteins in pooled human follicular fluid which suppress follicular response to gonadotropins. J. Clin. Endocrinol. Metab., 56:35—41. 611. diZerega, G. S., Marrs, R. P., Campeau, J. D., and Kling, O. R. (1983): Human granulosa cell secretion of protein(s) which suppress follicular response to gonadotropins. J. Clin. Endocrinol. Metab., 56:147-155. 612. Kling, O. R., Roche, P. C., Campeau, J. D., Nishimura, K., Nak¬ amura, R. M., and diZerega, G. S. (1984): Identification of protein(s) in porcine follicular fluid which suppress follicular response to go¬ nadotropins. Biol. Reprod., 30:564-572. 613. diZerega, G. S., Campeau, J. D., Nakamura, R. M., Ujita, E. L., Lobo, R., and Marrs, R. P. (1983): Activity of a human follicular fluid protein(s) from spontaneous and induced ovarian cycles. J. Clin. Endocrinol. Metab., 57:838-846. 614. diZerega, G. S., Goebelsman, U., and Nakamura, R. M. (1982): Identification of protein(s) secreted by the preovulatory ovary which suppresses the follicle response to gonadotropins. J. Clin. Endocri¬ nol. Metab., 54:1091-1096. 615. Brailly, D., Gougeon, A., Milgrom, E., Bomsel-Helmreich, O., and Papiemik, E. (1981): Androgen and progestins in the human ovarian follicle: differences in the evolution of preovulatory, healthy non¬ ovulatory and atretic follicles. J. Clin. Endocrinol. Metab., 53:128134. 616. Veldhuis, J. D., Klase, P. A., and Hammond, J. M. (1981): Sex steroids modulate prolactin action in spontaneously lutein¬ izing porcine granulosa cells in vitro. Endocrinology, 108.14631468. 617. Schaar, H. (1976): Funktionelle morphologie der theca interna lm blaschenfollikel des menschlichen ovars. Acta Anat., 94:283-298. 618. Leung, P. C. K., and Armstrong, D. T. (1980): Interactions of steroids and gonadotropins in the control of steroidogenesis in the ovarian follicle. Annu. Rev. Physiol., 42.71-82.

383

619. Tsang, B. K., Leung, P. C. K., and Armstrong, D. T. (1979): Inhibition by estradiol-17(3 of porcine thecal androgen production in vitro. Mol. Cell. Endocrinol., 14:131—139. 620. Henderson, K. M., McNeilly, A. S., and Swanston, I. A. (1982): Gonadotrophin and steroid concentrations in bovine follicular fluid and their relationship to follicle size. J. Reprod. Fertil., 65:467473. 621. Henderson, K. M., and Swanston, I. A. (1978): Androgen aromatization by luteinized bovine granulosa cells in tissue culture. J. Reprod. Fertil., 52:131-134. 622. Henderson, K. M., and Moon, Y. S. (1979): Luteinization of bovine granulosa cells and corpus luteum formation associated with loss of androgen aromatizing ability. J. Reprod. Fertil., 56:89-97. 623. Matson, P. L., Tyler, J. P. P., and Collins, W. P. (1981): Follicular steroid content and oocyte meiotic status after PMSG stimulation of immature hamsters. J .'Reprod. Fertil., 61:443-452. 624. Khalil, M. W., and Snow, K. (1985): 19-norandrostenedione (4estren-3,17-dione) levels in follicular fluid during ovarian follicular development in gilts. Biol. Reprod., 32 (Suppl. 1): 122 (abstr. 170). 625. Daniel, S. A. J., Khalil, M. W., and Armstrong, D. T. (1986): 19Norandrostenedione (4-estrene-3,17-dione) inhibits porcine oocyte maturation in vitro. Gamete Res., 13:173-184. 626. Axelrod, L. R., and Goldzieher, J. W. (1970): The effect of cofactors on steroid biosynthesis in normal ovarian tissue. Biochim. Biophys. Acta, 202:349-353. 627. Swanston, I., McNatty, K. P., and Baird, D. T. (1977): The con¬ centration of prostaglandin p2a and steroids in the human corpus luteum. J. Endocrinol., 73:115-122. 628. Yen, S. S. C. (1978): The human menstrual cycle (integrative func¬ tion of the hypothalamic-pituitary-ovarian-endometrial axis). In: Reproductive Endocrinology, edited by S. S. C. Yen and R. B. Jaffe, pp. 126-151. Saunders, Philadelphia. 629. McNatty, K. P., and Sawers, R. S. (1975): Relationship between the endocrine environment within the Graafian follicle and the sub¬ sequent rate of progesterone secretion by human granulosa cells in vitro. J. Endocrinol., 66:391-400. 630. Laborde, N., Carril, M., Cheviakoff, S., Croxatto, H. D., Pedroza, E., and Rosner, J. M. (1976): The secretion of progesterone during the periovulatory period in women with certified ovulation. J. Clin. Endocrinol. Metab., 43:1157-1163. 631. Fowler, R. E., Chan, S. T. H., Walters, D. E., Edwards, R. G., and Steptoe, P. C. (1977): Steroidogenesis in human follicles ap¬ proaching ovulation as judged from assays of follicular fluid. J. Endocrinol., 72:259-271. 632. Fortune, J. E. (1981): Bovine theca and granulosa cells interact to promote androgen and progestin production. Biol. Reprod., 24 (Suppl. 1): 39A (abstr.). 633. Peters, H., and McNatty, K. P. (1980): The Ovary. Granada, Lon¬ don. 634. Mestwerdt, W. (1977): Die follikel-granulosazellen in beziehung zur steroid-biosynthese in der periovulationsphase. Fortschr. Med., 95:361365. 635. Mestwerdt, W., Muller, O., and Brandau, H. (1977): Die differenzierte struktur und funktion der granulosa und theka in verschiedenen follikelstadien menschlicher ovarien. 2. Mittleilung: der reifende, reife, sprungreife und frisch geplatzte follikel. Arch. Gynaekol., 222:115-136. 636. Mestwerdt, W., Muller, O., and Brandau, H. (1977): Die differenzierte struktur und funktion der granulosa und theka in verschiedenen follikelstadien menschlicher ovarien. 1. mitteilung: der pri¬ mordial-, primar-, sekundar- und ruhende tertiarfollikel. Arch. Gynakol., 222:45-71. 637. Albertini, D. F., and Anderson, E. (1974): The appearance and structure of intercellular connections during the ontogeny of the rabbit ovarian follicle with particular reference to gap junctions. J. Cell Biol., 63:234-250. 638. Bjersing, L., and Cajander, S. (1974): Ovulation and the mechanism of follicle rupture. VI. Ultrastructure of theca interna and the inner vascular network surrounding rabbit Graafian follicles prior to in¬ duced ovulation. Cell Tissue Res., 153:31—44. 639. Delforge, J. P., Thomas, K., Roux, F. Cameiro de Siqueira, J., and Ferin, J. (1972): Time relationships between granulosa cell growth and luteinization, and plasma luteinizing hormone discharge in hu¬ man. I. A morphometric analysis. Fertil. Steril., 23:1-11.

384

/ Chapter 10

640. Norman, R. L., and Greenwald, G. S. (1972): Follicular histology and physiological correlates in the preovulatory hamster. Anat. Rec., 173:95-108. 641. Hermreck, A. S., and Greenwald, G. S. (1964): The effects of unilateral ovariectomy on follicular maturation in the guinea pig. Anat. Rec., 148:171-176. 642. Boucek, R. J., Telegdy, G., and Savard, K. (1967): Influence of gonadotropin on histochemical properties of the rabbit ovary. Acta Endocrinol. (Copenh.), 54:295-310. 643. Klinken, S. P., and Stevenson, P. M. (1977): Changes in enzymatic activities during the artificially stimulated transition from follicular to luteal cell types in rat ovary. Eur. J. Biochem., 81:327-332. 644. Pupkin, M., Bratt, H., Weisz, J., Lloyd, C. W., and Balogh Jr., K. (1966): Dehydrogenases in the rat ovary. I. A histochemical study of A5-3(3- and 20a-hydroxysteroid dehydrogenases and enzymes of carbohydrate oxidation during the estrous cycle. Endocrinology, 79:316-327. 645. Bjersing, L. (1977): Ovarian histochemistry. In: The Ovary, Vol. 1, edited by S. Zuckerman and B. J. Wier, pp. 303-391. Academic, New York. 646. Wingate, A. L. (1970): A histochemical study of the hamster ovary. Anat. Rec., 166:399 (abstr.). 647. Blaha, G. C., and Leavitt, W. W. (1970): The distribution of A53 3-hydroxy steroid activity in the golden hamster during the estrous cycle, pregnancy, and lactation. Biol. Reprod., 3:362-368. 648. Norman, R. L., and Greenwald, G. S. (1971): Effect of phenobarbital, hypophysectomy, and X-irradiation on preovulatory proges¬ terone levels in the cyclic hamster. Endocrinology, 89:598-605. 649. Friedrich, F., Breitenecker, G., Salzer, H., and Holzner, J. H. (1974): The progesterone content of the fluid and the activity of the steroid33-ol-dehydrogenase within the wall of the ovarian follicles. Acta Endocrinol. (Copenh.), 76:343-352. 650. Hillensjo, T., Bauminger, S., and Ahren, K. (1976): Effect of lu¬ teinizing hormone on the pattern of steroid production by preovu¬ latory follicles of pregnant mare’s serum gonadotropin-injected im¬ mature rats. Endocrinology, 99:996-1002. 651. LeMaire, W. J., and Marsh, J. M. (1975): Interrelationships between prostaglandins, cyclic AMP and steroids in ovulation. J. Reprod. Fertil., 22 (Suppl.):53-74. 652. Younglai, E. V. (1977): Steroid production by isolated rabbit ovarian follicles: effects of luteinizing hormone from mating to implantation. J. Endocrinol., 73:59-65. 653. Hori, T., Ide, M., and Miyake, T. (1969): Pituitary regulation of preovulatory oestrogen secretion in the rat. Endocrinol. Jpn., 16:351— 360. 654. Hubbard, C. J., and Greenwald, G. S. (1982): Cyclic nucleotides, DNA, and steroid levels in ovarian follicles and corpora lutea of the cyclic hamster. Biol. Reprod., 26:230-240. 655. Murdoch, W. J., and Dunn, T. G. (1982): Alterations in follicular steroid hormones during the preovulatory period in the ewe. Biol. Reprod., 24:1171-1181. 656. Dieleman, S. J., Bevers, M. M., Poortman, J., and van Tol, H. T. M. (1983): Steroid and pituitary hormone concentrations in the fluid of preovulatory bovine follicles relative to the peak of LH in the peripheral blood. J. Reprod. Fertil., 69:641-649. 657. Testart, J. Castanier, M., Feinstein, M.-C., and Frydman, R. (1982): Pituitary and steroid hormones in the preovulatory follicle during spontaneous or stimulated cycles. In: Follicular Maturation and Ovu¬ lation, edited by R. Rolland, E. V. van Hall, S. G. Hillier, K. P. McNatty, and J. Schoemaker, pp. 193-201. Excerpta Medica, Am¬ sterdam. 658. Van Look, P. F. A., Templeton, A. A., Swantson, I. A., et al. (1984). The effect of hCG on steroid levels in human Graafian fol¬ licles. Proceedings of the 3rd Joint Meeting of the British Endocrine Societies, Edinburgh, Abstract. 659. Moor, R. M. (1974): The ovarian follicle of the sheep: inhibition of oestrogen secretion by luteinizing hormone. J. Endocrinol., 61:455463. 660. Bockaert, J., Hunzicker-Dunn, M., and Bimbaumer, L. (1976): Hor¬ mone-stimulated desensitization of hormone-dependent adenylyl cy¬ clase: dual action of luteinizing hormone on pig Graafian follicle membranes. J. Biol. Chem., 251:2653-2663. 661. Mori, T., Fujita, Y., Suzuki, A., Kinoshita, Y., Nishimura, T., and Kambegawa, A. (1978): Functional and structural relationships in

662.

663.

664.

665.

666.

667.

668.

669.

670.

671.

672.

673.

674.

675.

676.

677.

678.

679.

680.

681.

682.

steroidogenesis in vitro by human ovarian follicles during maturation and ovulation. J. Clin. Endocrinol. Metab., 47:955-966. Edwards, R. G., Steptoe, P. C., Fowler, R. E., and Bailie, J. (1980): Observations «pn preovulatory human ovarian follicles and their as¬ pirates. Br. J. tfbstet. Gynaecol., 87:769-779. Landgren, B.-M., Aedo, A.-R., Nunez, M., Cekan, S. Z., and Diczfalusy, E. (1977): Studies on the pattern of circulating steroids in the normal menstrual cycle. Acta Endocrinol. (Copenh.), 84:620632. Kirchick, H. J., and Bimbaumer, L. (1983): Luteal adenylyl cyclase does not develop sensitivity to desensitization by human chorionic gonadotropin in the absence of nonluteal ovarian tissue. Endocri¬ nology, 113:2052-2058. Richards, J. S., Ireland, J. J., Rao, M. C., Bemath, G. A., Midgley Jr., A. R., and Reichert Jr., L. E. (1976): Ovarian follicular devel¬ opment in the rat: hormone receptor regulation by estradiol, folliclestimulating hormone and luteinizing hormone. Endocrinology, 99:1562-1570. Rao, M. C., Richards, J. S., Midgley Jr., A. R., and Reichert Jr., L. E. (1977): Regulation of gonadotropin receptors by luteinizing hormone in granulosa cells. Endocrinology, 101:512-523. Suzuki, K., and Tamaoki, B. I. (1980): Postovulatory decrease in estrogen production is caused by the diminished supply of aromatizable androgen to ovarian aromatase. Endocrinology, 107:2115— 2116. Hillensjo, T., Hamberger, L., and Ahren, K. (1977): Effect of an¬ drogens on the biosynthesis of estradiol-173 by isolated preovulatory follicles. Mol. Cell. Endocrinol., 9:183-193. Suzuki, K., and Tamaoki, B. I. (1983): Acute decrease by human chorionic gonadotropin of the activity of preovulatory ovarian Hahydroxylase and C-17-20-lyase is due to decrease of microsomal cytochrome P-450 through de novo synthesis of ribonucleic acid and protein. Endocrinology, 113:1985-1991. Eckstein, B., and Tsafriri, A. (1986): The steroid C-17,20-lyase complex in isolated Graafian follicles: effects of human chorionic gonadotropin. Endocrinology, 118:1266-1270. Bjersing, L., Hay, M. F., Kann, G., et al. (1972): Changes in gonadotrophins, ovarian steroids and follicular morphology in sheep at oestrus. J. Endocrinol., 52:465-479. Watzka, M. (1957): Weibliche genitalorgane. Das ovarium. In: Handbuch der Mikroskopischen Anatomies des Menschen, Vol. 7, edited by M. V. Mollendorf and W. Bargmann, pp. 1-178. Springer, Berlin. LeMaire, W. J., Yang, N. S. T., Behrman, H. H., and Marsh, J. M. (1973): Preovulatory changes in the concentration of prosta¬ glandins in rabbit Graafian follicles. Prostaglandins, 3:367-376. Yang, N. S. T., Marsh, J. M., and LeMaire, W. J. (1974): Post¬ ovulatory changes in the concentrations of prostaglandins in rabbit Graafian follicles. Prostaglandins, 6:37^44. Armstrong, D. T., Moon, Y. S., and Zamecnik, J. (1974): Evidence for a role of prostaglandins in ovulation. In: Gonadotropins and Gonadal Function, edited by N. R. Moudgal, pp. 345-356. Aca¬ demic, New York. Armstrong, D. T., and Zamecnik, J. (1975): Pre-ovulatory elevation of rat ovarian prostaglandin F, and its blockade by indomethacin. Mol. Cell. Endocrinol., 2:125-131. Bauminger, S., and Lindner, H. R. (1975): Periovulatory changes in ovarian prostaglandin formation and their hormonal control in the rat. Prostaglandins, 9:737-751. Ainsworth, L. Baker, R. D., and Armstrong, D. T. (1975): Pre¬ ovulatory changes in follicular fluid prostaglandin F levels in swine. Prostaglandins, 9:915-925. Tsang, B. K., Ainsworth, L., Downey, B. R., and Armstrong, D. T. (1979): Pre-ovulatory changes in cyclic AMP and prostaglandin concentrations in follicular fluid of gilts. Prostaglandins, 17:141148. Plunkett, E. R., Moon, Y. S., Zamecnik, J., and Armstrong, D. T. (1975): Preliminary evidence of a role for prostaglandin F in human follicular function. Am. J. Obstet. Gynecol., 123:391-397. Henderson, K. M., and McNatty, K. P. (1975): A biochemical hy¬ pothesis to explain the mechanism of luteal regression. Prostaglan¬ dins, 9:779-798. Armstrong, D. T. (1981): Prostaglandins and follicular functions. J. Reprod. Fertil., 62:283-291.

Follicular Steroidogenesis and Its Control

683. Janson, P. O. (1975): Effects of luteinizing hormone on blood flow in the follicular rabbit ovary as measured by radioactive microspheres. Acta Endocrinol. (Copenh.), 79:122-133. 684. Armstrong, D. T., and Grinwich, D. L. (1972): Blockade of spon¬ taneous and LH-induced ovulation in rats by indomethacin, an in¬ hibitor of prostaglandin biosynthesis. Prostaglandins, 1:21-28.

/

385

685. Grinwich, D. L., Kennedy, T. G., and Armstrong, D. T. (1972): Dissociation of ovulatory and steroidogenic actions of luteinizing hormone in rabbits with indomethacin, an inhibitor of prostaglandin synthesis. Prostaglandins, 1:89-95. 686. Lau, I. F., Saksena, S. K., and Chang, M. C. (1974): Prostaglandins F and ovulation in mice. J. Reprod. Fertil., 40:467^469.

*

The Physiology of Reproduction, edited by E. Knobil and J. Neill et al. Raven Press, Ltd., New York © 1988.

CHAPTER 11

Follicular Selection and Its Control Gilbert S. Greenwald and Paul F. Terranova Early Stages of Follicular Development, 387 Rat, 389 • Mouse, 390 • Hamster, 390 • Guinea Pig, 391 • Sheep, 391 • Human, 391

Follicular Development 411 • Diestrus, 412

During

the

Cycle:

Estrus,

Structure and Function of the Antral Follicle, 391

Follicular Development During the Estrous Cycles of Sheep, Cow, and Sow, 414

Thecal Structure and Function, 394

Follicular Development in the Cow, 417

Follicular Atresia, 396

Follicular Development During the Porcine Estrous Cycle, 419

The Normal Process of Atresia: Membrana Granulosa, 397 • Experimental Induction of Follicular Atresia, 400

Follicular Development and Superovulation, 404

Follicular Development During the Menstrual Cycle of Macaca and Human, 419 Follicular Development in the Human, 421

Effects of Unilateral Ovariectomy on Follicular Development, 407

Follicular Development During Pregnancy, 424

Short-Term Effects of ULO, 408 • Long-Term Effects of ULO, 409

Follicular Development During the Estrous Cycle of Rat and Hamster, 410

“One of the most intriguing mysteries in ovarian physi¬ ology is what factors determine whether one follicle remains quiescent, another begins to develop but later becomes atretic, while still a third matures and ovulates” (1). The above description of the fate of three primordial follicles sets the theme for this chapter. In view of the paucity of information about primordial follicles, emphasis is upon follicles once they have entered into the growing pool. We hope to provide a balanced account of follicu¬ lar development in the cyclic, pregnant, and lactating animal. A comparative approach that will point out sig¬ nificant species similarities and differences is stressed. Moreover, an attempt will be made to cover all aspects of folliculogenesis. Folliculogenesis has attracted a great deal of attention and, where feasible, this chapter focuses on recent findings. Table 1 lists a series of relevant reviews and chap¬ ters—which is by no means complete. Further entree to the literature on follicular development can be gained by con¬ sulting the volumes published during the past 10 years (Ta¬ ble 2).

387

Effects of Lactation on Follicular Development, 426 Summary and Conclusions, 429 Acknowledgments, 432 References, 432

EARLY STAGES OF FOLLICULAR DEVELOPMENT The majority of mammals restrict oogonial proliferation to prenatal development or to the early postnatal period (29); the rare exceptions are lemurs, in which mitotic activity of germ cells is demonstrable even in the adult (30). Thus, in most mammals oogonia are transformed before or soon after birth into primary oocytes that are characterized by a pro¬ longed meiotic prophase and surrounded by a squamous layer of pregranulosa cells. These primordial follicles con¬ stitute the resting stockpile of nongrowing follicles that are progressively depleted during the reproductive life span. Primordial follicles continuously (presumably) leave the nongrowing pool by being converted into primary follicles in which the oocyte is surrounded by a unilaminar layer of cuboidal granulosa cells—the descendents of the pregran¬ ulosa cells. The follicle is then launched on its career as a growing or developing follicle, culminating in either ovu¬ lation or, more likely, atresia at some stage in its subsequent development. According to Peters (31), the initiation of

388

/ Chapter 11 TABLE 1. Reviews and chapters relevant to follicular selection

Year

Author

1947 1956 1959 1974 1977 1977 1977 1980 1981 1983 1984 1984 1985

Hisaw (2) Brambell (3) Falck (4) Greenwald (5) Channing and Tsafriri (6) Lindner et al. (7) Armstrong and Dorrington (8) Richards (9) diZerega and Hodgen (10) Erickson (11) Hsueh et al. (12) Tsafriri and Braw (13) Hillier (14)

follicle growth is not dependent on gonadotropins since unilateral ovariectomy of 2-day-old mice does not change the number of normal developing follicles present 12 days later. On the other hand, continuous daily injection from birth of an antiserum to gonadotropins leads to signs of altered follicular development by 5 days of age and definite effects from 7 days onwards (32). Follicular development in gonadotropin-deprived mice rarely progresses beyond the 40-cell stage, corresponding to a type 3b follicle: the small¬ est medium-sized follicle with an oocyte slightly larger than 20 p.m in diameter (33). Transplantation of ovaries of 1day-old rats to ovariectomized or ovariectomized-hypophysectomized adult female rats results 15 days later in the same number of small and medium follicles (up to two layers of granulosa cells), but the transplants in the gonadotropinrich environment contain many more secondary follicles (multilaminar granulosa layers) and large follicles with in¬ cipient formation of an antral cavity. The oocytes are also

considerably larger in the transplants in the ovariectomized hosts (34). The conclusion is that gonadotropins, especially follicle-stimulating hormone (FSH), enhance early follicle cell development and early oocyte growth. In the immature mouse (33) and rat (35), more follicles start to grow per day in young animals (7-day-old mice, 16-day-old rats) than in older ones. This perhaps can be explained on the basis of high levels of gonadotropins in the prepubertal rodent (rat: 36; hamster: 37). The reduced number of growing follicles recruited per day in 3-week-old mice has been attributed to a factor produced by atretic follicles that re¬ duces growth initiation (31). The fact that large follicles have already differentiated by 21 days of age (33) raises the possibility that production of steroids and/or inhibin by the enlarging follicles can act via the hypothalamic-pituitary axis to account for fewer follicles entering into the growing pool. . With age the number of primordial follicles declines in parallel with the number of growing follicles (mouse: 38,39; rat: 40; human: 41,42). The size of the pool of primordial follicles determines the fraction that is stimulated to grow. A reduction in the size of the pool reduces the number recruited into the growing pool. In the mouse, this rela¬ tionship holds true whether the nongrowing pool is reduced by age or artificially by injection of dimethyl-benzanthra¬ cene (38) or by early androgenization (31). There is con¬ siderable individual variation in the number of primordial follicles in man (41) and rhesus monkey (43). The number of primordial follicles shows significant differences among three strains of rats (44). Interestingly, the strain with the greatest number of primordial follicles also has the greatest number of growing follicles and is most responsive to ex¬ ogenous FSH. It is disconcerting, however, that this strain is the less fecund, as judged by a higher incidence of sterile matings and smaller litter sizes; whether there are differ¬ ences in ovulation rate between the strains is unknown (45).

TABLE 2. Relevant books on the ovarian follicle, 1977-1985 Year

Title

Author(s)

1977 1978 1978 1979 1979 1980 1980 1981 1982 1983 1982-84 1984

The Ovary, 2nd ed. The Vertebrate Ovary Control of Ovulation Ovarian Follicular Development and Function Ovarian Follicular and Corpus Luteum Function Conception in the Human Female Biology of the Ovary Dynamics of Ovarian Function Intraovarian Control Mechanisms Factors Regulating Ovarian Function Reproduction in Mammals, 2nd ed. Marshall’s Physiology of Reproduction/Reproductive Cycles of Vertebrates, 4th ed. Biology of Ovarian Follicles in Mammals Proceedings of the Fifth Ovarian Workshop

Zuckerman and Weir (15) Jones (16) Crighton et al. (17) Midgley and Sadler (18) Channing et al. (19) Edwards (20) Motta and Hafez (21) Schwartz and Hunzicker-Dunn (22) Channing and Segal (23) Greenwald and Terranova (24) Austin and Short (25) Lamming (26)

1985 1985

Guraya (27) Toft and Ryan (28)

Follicular Selection / One of the most critical steps in folliculogenesis is the transformation of primordial into primary follicles. Intui¬ tively, it seems likely that the conversion of the flattened pregranulosa cells into a cuboidal epithelium depends on cues provided by the oocytes, but the nature of the signal is unclear. Several lines of evidence indicate that the pre¬ granulosa cells are unable to form follicles in the absence of the oocyte. Injection of busulfan (an alkylating agent) into pregnant rats destroys all primordial germ cells, and consequently, a sterile gonad devoid of steroidogenic tissue is formed (46). Similarly, a mutant mouse strain may have 2 to 20 oocytes at birth, but their disappearance by 3 months prevents any further follicular development (47). By a series of mitotic divisions, the unilaminar primary follicle is converted into a multilaminated preantral stage designated a secondary follicle; at various times in its life history the secondary follicle becomes invested with thecal cells. Finally, with the appearance of an antral cavity, the secondary follicle is converted into a tertiary follicle. The morphological and biochemical changes associated with these changes will be discussed later. At this point, we are con¬ cerned with a very basic issue: Is the growth and differ¬ entiation of primary and secondary follicles under the in¬ fluence of gonadotropins, or is it only with the development of an antral cavity that the tertiary follicle becomes depen¬ dent on FSH and luteinizing hormone (LH)? The relatively constant number of preantral follicles throughout the estrous cycle has often been cited as evidence that they are unaffected—or at most slightly affected—by changes in gonadotropins. Thus, throughout the cycle of the mouse, no cyclic changes in numbers exist for follicles of types 3a to 6 [type 6 = incipient formation of antral cavity (48)]. Based on large sample sizes (11-18 rats/day of cycle), at least 21 follicles/ovary are recruited from less than 260 p.m into greater than 260 p.m between proestrus and estrus (49). Moreover, the number of smaller follicles (70-110 p,m) significantly increases at diestrus; a definite antrum (i.e., a tertiary follicle) is present in follicles 120 to 130 |xm in diameter. Hence, the number of preantral follicles does change during the estrous cycle of the rat. It is note¬ worthy that rat follicles less than 70 |xm in diameter are never atretic (49). Similarly, in the periovulatory period of the rhesus monkey, there is a significant increase in the percentage of small preantral follicles ranging from 100 to 200 p.m in diameter; the transition to an antral follicle occurs between 200 and 250 |xm (43). Based on the terminology of Pedersen and Peters (50), there are significantly more type 3b and 4 rat follicles at estrus and metestrus than on other days of the cycle (51). Of perhaps greater significance, the duration of the DNA synthesis phase is shorter for all primary and secondary follicles at estrus than at other stages of the cycle (51), and this is also true for the cyclic mouse (48). The number of preantral follicles with two to five layers of granulosa cells does not vary during the hamster cycle (52). However, preantral hamster follicles, with one

389

to five layers of granulosa cells show significant increases in [3H]thymidine incorporation on proestrus and estrus as evaluated by autoradiography (53) or after in vitro incu¬ bation with [3H]thymidine (54). Thus, the dogma of the unchanging number and responsiveness of preantral follicles is refuted in a number of species. Still another way of demonstrating that preantral follicles can be stimulated in intact animals by gonadotropins is by evaluation of the effects of superovulation induced by preg¬ nant mare serum (PMS). For example, 26- to-28-day-old mice ovulate 60 ova in response to 10 IU PMS followed 56 hr later by 5 IU human chorionic gonadotropin (hCG): rats 24 to 30 days old ovulate 50 eggs after 30 IU PMS and an ovulating dose of 10 IU hCG (55). Adult rats injected with 50 IU PMS and then with 15 IU hCG 3 days later ovulate 43 ova per animal (56). The number of ova ovulated hardly makes it feasible that reduced atresia of tertiary fol¬ licles can solely account for these results; rather, recruitment of preantral follicles seems to be an essential component. Unequivocal evidence that preantral follicles are recruited by PMS treatment is provided by the cyclic hamster. In¬ jection of 30 IU PMS on estrus (day 1 of cycle) results in the ovulation of 54 ova by the next cycle, and this is as¬ sociated by day 2 with a doubling in the number of follicles greater than 267 |xm in diameter (57). Within 4 hr after the injection of 30 IU PMS on estrus, hamster follicles with four to five layers of granulosa cells respond by a significant reduction in their numbers and a concomitant increase in follicles with incipient formation of an antral cavity (58). Collectively, these results point to significant effects of go¬ nadotropins on secondary (and even primary) follicles in intact animals. It is frequently stated that gonadotropic hormones are unessential for early growth of follicles and that FSH and LH become indispensable for further development only at the transformation of the secondary to a tertiary follicle. Although species differences may exist, a careful perusal of the literature indicates that early follicular development is influenced by gonadotropins, but the picture is confusing because of the frequent subjective, anecdotal evaluations and the way even quantitative data have been presented. What follows, then, is a species-by-species account of the effects of hypophysectomy on follicular development that attempts to provide a balanced view on this important sub¬ ject.

Rat Rats hypophysectomized at 28 days of age were killed at various postoperative intervals, and healthy follicles were classified as primary or vesicular; the former category in¬ cludes preantral follicles with two or more layers of gran¬ ulosa cells (59). By 10 and 38 days after hypophysectomy, the average number of primary follicles was 102 and 20,

390

/ Chapter

11

respectively, compared with 213 on the day after operation; similarly the number of vesicular follicles at the same time intervals were 99 and 3 compared with 160 on day 1. The maximal diameter of follicles maintained posthypophysectomy was 250 |xm, but the number obviously falls sharply with time. An often-cited, but somewhat confusing study by Paesi (60) dealt with hypophysectomy of young rats weighing 61 to 72 g. A week after operation, there was a 71% increase in the smallest follicles (23-32 pm in di¬ ameter). Since primordial follicles were not counted, pre¬ sumably these were primary follicles with one layer of cuboidal granulosa cells. Approximately 20% of the small follicles showed signs of beginning atresia. One week after hypophysectomy, the largest follicles present were 360 pm compared preoperatively to 576 pm. The accumulation of small follicles might represent a decrease in the number of follicles developing into larger stages per unit of time (60). An excellent, thorough study by De Reviers (61) dealt with follicular development in the immature rat, with follicles classified by the volume of granulosa cells. Volume ranged from 1,259 pm2 for primary follicles to 25,200 pm2 for large tertiary follicles. Ten days after hypophysectomy at 27 days of age, there is a 43% reduction in the number of smallest follicles (types 3b and 4) and a 37% reduction in intermediate follicles (type 5a and 6) and no larger stages (type 6-8) present compared to intact 28-day-old rats. In hypophysectomized rats killed 105 or 135 days later, the total number of small follicles varied from 8 to 48, and only sporadic numbers of intermediate follicles were present. De Reviers pointed out that long-term hypophysectomy of male rats leads to the development of gonadotropin-like cells in the pars tuberalis as evidenced by immunocytology (cited in 61). Even the long-term hypophysectomized rat may therefore not be completely devoid of some gonadotropin reserve. Ovine FSH is capable of stimulating follicular growth in rats hypophysectomized for as long as 25 days, and this effect is manifested on all stages of folliculogenesis. The action of FSH—depending on dose—involves diminished atresia but also increased recruitment. The recruitment may even affect primordial follicles, as demonstrated by injecting [3H]thymidine 1 hr before hypophysectomy with the rats killed 72 hr later. There was approximately a threefold greater labeling index in type 3b follicles after FSH treatment (61). An unpublished study has considered follicular devel¬ opment in adult hypophysectomized rats with the end¬ point—21 days later—expressed as numerical density of follicles based on their diameter (cited in 62). Follicles smaller than 60 p,m were not measured. The results indicate a significant decline in follicles larger than 175 |xm by 3 weeks after hypophysectomy. A single injection of Armour FSH (4 mg) resulted in a significant increase in the number of follicles 75 to 125 |xm in diameter 24 hr later. A recent study reconfirms a number of the above findings: Rats hy¬ pophysectomized at 26 days of age showed significant de¬ clines in secondary and tertiary follicles 3 days later; e.g.,

the number of follicles with two to three layers of granulosa cells were reduced to 107 ± 12 per ovary from the normal value of 175 ± 22 (63). The number of healthy tertiary follicles was even more affected by 3 days: 4.5 ±1.2 ver¬ sus 39 ± 3 in intact controls. The above results are so striking that one wonders why there is any question about the dependency of preantral rat follicles on gonadotropin support. In part, this is because some investigators have been more impressed by the ability of at least some follicles in the hypophysectomized rat to maintain some semblance of normal function. Thus, rats hypophysectomized on day 22 and injected 10 days later with [3H]thymidine show a degree of labeling of granulosa cells in small follicles similar to that in intact animals (64). The authors state that “although many follicles were atretic, a few follicles with more than one layer of granulosa cells persisted.” In another study, hypophysectomized rats were implanted 5 days postsurgery with pumps containing [3H]thymidine and were then killed 8 days later (65). Fol¬ licles were heavily labeled over the granulosa cells: “Some of the labeled follicles were healthy,” but “many labeled follicles were atretic.” Again, the author was more im¬ pressed with the qualitative aspects—not quantitative—of follicular growth after hypophysectomy.

Mouse Hypophysectomy of 12 postpubertal mice resulted in few follicles developing beyond the two-layered granulosa stage, but the authors recognized the tentativeness of the conclu¬ sion because of the heterogeneous ages of the mice and different survival times (66). It is interesting, however, that in intact cyclic mice follicles are committed to either normal development or atresia by the time the third layer of gran¬ ulosa cells has differentiated; a clearly established thecal layer is evident at this stage (67).

Hamster In hamsters hypophysectomized on estrus, 1 to 28 days postsurgery, 99% of the follicles have five or fewer layers of granulosa cells, which is considerably below the size of the largest preantral follicle (68). With time, a greater per¬ centage of follicles than normal accumulates in the group with two to three layers of granulosa cells. After a hiatus of a week, daily injection of 200 jxg ovine FSH (NIH-S7) for 4 days, followed by hCG, resulted in the ovulation of an average of 32 ova. Interestingly, 200 |xg FSH on the first day of treatment followed by 50 fxg/day thereafter led to ovulation of 9 ova—a regimen and number of ovulations simulating the pattern in the intact cyclic hamster. Three days after immature hamsters were hypophysectomized, the number of healthy secondary and tertiary follicles was re¬ duced from 405 to 251 per ovary and there was already a

Follicular Selection

significant reduction in follicles with two to eight layers of granulosa cells and larger follicles had vanished (63). Sim¬ ilar results have also been observed in the adult hypophysectomized hamster (69). Four days after hypophysectomy of adult hamsters, considerable amounts of FSH can still be eluted from the nonluteal ovary and with significantly greater FSH receptor affinity than from follicles of intact hamsters (70). This finding has two important implications: Small preantral follicles have a greater affinity for FSH than large ones (there is no appreciable FSH binding by the interstitium), and it may take considerable time, depending on the species, before FSH completely disappears from the ovary of the hypophysectomized animal and a truly anhormonal environment is established.

Guinea Pig An often-cited paper (71) reported on three hypophysec¬ tomized animals that were killed 12 days after surgery: “There was no reduction in the number of nonvesicular follicles.” This is strictly an anecdotal account. Subse¬ quently, 3- to 4-week-old guinea pigs were hypophysec¬ tomized and groups of three or four animals killed over the next 4 days (72). Follicles in the range of 140 to 800 |xm in diameter were counted. The percentage of normal follicles from 140 to 356 |xm did not appreciably differ between controls and hypophysectomized guinea pigs over the next 14 days. Only one of three animals killed on day 4 had any normal vesicular follicles, and in the other animals, all fol¬ licles beyond the primordial stage were atretic. Paradoxi¬ cally, by 6 and 14 days all ovaries contained vesicular fol¬ licles and a greater percentage were healthy: 63% versus 14% on day 4. No explanation was offered for these unusual results. Is it a matter of small sample size or with time does another source of gonadotropins develop in the hypophy¬ sectomized guinea pig (pars tuberalis)?

Sheep Ewes were hypophysectomized at estrus, with one ovary removed 4 days later and the other removed 70 days after treatment (73). All follicles with three or more layers of granulosa cells were counted. Four days after hypophysec¬ tomy the number of preantral follicles (60% granulosa via¬ bility were associated with healthy oocytes; e.g., at proes¬ trus, of 7.3 large antral follicles per ovary, only 4 were normal. As rat follicles become atretic, ultrastructurally, intercellular spaces between granulosa cells increase (168); this most likely represents a loss of gap junctions, which are reduced in atretic follicles (134). In medium-sized prean¬ tral follicles in the rat, incipient atretic changes in granulosa cells have a patchy distribution in that some cells show nuclear condensation (pyknosis) and marked dilatation of cytoplasmic organelles, with the cytoplasm and microvilli of the oocyte still well preserved (169). Adjacent normal granulosa cells are found to undergo mitosis. The granulosa cells also become flattened and lose cytoplasmic evaginations during atresia, whereas normal granulosa cells are spherical and have irregular cytoplasmic projections. These cytoplasmic projections normally correlate with the devel¬ opment of LH binding to granulosa cells (170). Thus, the loss of microvilli corresponds to loss of LH binding and the onset of atresia (171). With the loss of gap junctions and the loss of LH binding to granulosa cells, it is highly prob-

398

/

Chapter

11

able that the intercellular communication and trophic actions of LH needed to maintain an intact, well-structured mem-

tradiol. Thus, atresia of human follicles was associated with differentiation of granulosa cells into androgen-producing

brana granulosa are lost. An unusual variant of granulosa degeneration is present in the sheep follicle (172). The process again begins in granulosa cells bordering on the antral cavity. Phagocytic cells representing transformed granulosa cells increase in number as atresia progresses, akin to the process in non¬ mammalian vertebrates (155). During secondary atresia, atretic bodies develop from the fusion of many nuclei; these

cells.

multinucleated structures vary in diameter from 15 to 400 |xm. Atretic sheep follicles (2-6 mm in diameter) have been reported to “regenerate” in vitro within 3 days by a two- to fourfold increase in the thickness of the granulosa layer (173). However, no mitotic figures are observed and his¬ tologically the cells are definitely not normal granulosa cells. It is possible to make a gross identification of atretic and healthy follicles in sheep (174), pig (175), and cow (176) based primarily on the vascularization of the theca, integrity of the membrana granulosa, and translucency of the follicle. In fact, in the ewe, normal and advanced atretic follicles can be correctly identified by histological criteria in over 95% of the cases (174). During atresia the granulosa cells from atretic follicles of the rabbit take up less 35S (177), suggesting a decrease in the synthesis of sulfate-containing mucopolysaccharides. In contrast, atretic bovine follicles have low levels of estrogen but high levels of progesterone and chondroitin sulfate in follicular fluid (178). This relationship is significant when chondroitin sulfate concentration is plotted against estrogen or the estrogen:progesterone ratio but not when histology is used as the variable. Histochemical and biochemical alterations in granulosa cells often precede definite morphologic changes in atretic granulosa cells (for citations, see ref. 179). These include an increase in lysosomal enzymes such as acid phosphatase and aminopeptidase (180), and their role in the induction of atresia has been recently evaluated (181). Using recom¬ binant DNA technology it should be possible to determine the gene regulating these granulosa cell enzymes and then, by turning on the gene, induce widespread atresia. In atretic follicles of a number of species (e.g., humans), the histo¬ chemical appearance of 3(3-HSD and lipid droplets in gran¬ ulosa cells is characteristic of atresia (182). Granulosa cells from healthy human follicles maintained in vitro for 48 hr produced large amounts of estradiol and progesterone and small amounts of androstenedione without added steroid precursors (183). The follicles were judged to be normal, based on their possessing at least 75% of the maximal number of granulosa cells for their diameters. Fol¬ licles with 1 (209). As atre¬ sia progresses the ratio shifts in favor of androgen primarily because of a fall in estradiol. In large (2=3.5 mm) atretic follicles, as judged by morphological criteria, follicles with signs of early degeneration have reduced aromatase activity, consistent with the idea that, in the ewe, lack of androgen substrate is not the limiting factor (210). In atretic ovine follicles (>4 mm), binding of [125I]FSH and LH does not differ from that in healthy follicles until the most advanced stage of atresia (211). However, for intermediate-sized an¬ tral follicles (2^4 mm), the loss of FSH binding by granulosa cells precedes the decline in hCG binding. It was concluded that the decline in binding of the labeled gonadotropins as atresia progresses is more likely a consequence than a cause of follicular regression. Bovine follicles from days 3 to 13 of the estrous cycle were assigned to healthy or atretic status based on follicular concentration of estradiol (estrogen active or inactive) and histology (212). From days 3 to 7, a single large estrogen active follicle is present. During this time span the estrogeninactive follicles exhibit significantly less binding of hCG to theca and granulosa and reduced FSH binding to gran¬ ulosa. Analysis of the data is complicated by the few es¬ trogen-inactive follicles, their significantly fewer granulosa cells on days 5 and 7, and a lack of agreement between histological classification and follicular fluid ratios of es¬ trogen to progesterone. There is no difference in [125I]hCG binding to bovine theca interna from healthy and atretic antral follicles (176). When perfused in vitro, the output of androstenedione by theca interna of normal bovine follicles

is considerably enhanced by equimolar concentrations of LH or hCG, but theca of atretic follicles are unresponsive. Lack of available receptors, therefore, does not seem to account for the* failure of androgen production by large atretic bovine follicles. Rather, distal events seem to be affected. A subsequent study (213) classified the theca of large bovine follicles (>8 mm) into three types based on the number and steroidogenic capacities of the associated granulosa cells. Type I theca (normal follicles) perfused in vitro with LH respond by secreting increased cAMP, an¬ drostenedione, and testosterone; Type II theca (from pos¬ sibly early atretic follicles) secrete increased cAMP and progesterone but not androgens; and Type III theca (defi¬ nitely atretic) do not increase cAMP or steroids in response to LH. The theca of all three types contain LH receptors. These results are strikingly similar to findings with some of the rodent models (see below).

Experimental Induction of Follicular Atresia During the past 15 years several models involving rodents have been developed to study atresia, with a number of the experimental designs based on previously described pro¬ cedures (Table 3). Despite disparities in species and ex¬ perimental design, the models have yielded remarkably sim¬ ilar conclusions but differ in key aspects from spontaneous atresia in sheep and human (see above). With one exception (198), the models focus on tertiary follicles, but they could just as well be utilized to study atresia of preantral follicles. As expected, the models using hypophysectomized animals show more rapid onset of morphological and steroidogenic

TABLE 3. Rodent models of follicular atresia Model 1. Hypophysectomy of proestrous rat 2. Hypophysectomy + PMS followed by anti-PMS (hamster) 3. Pentobarbital delay of ovulation in rat 4. Phenobarbital delay of ovulation in hamster 5. PMS to immature mouse 6. PMS to immature rat 7. PMS to immature hamster 8. Hypophysectomy of prepubertal rat, treatment with E2 or DES, then hormone withdrawal 9. Androgen treatment of hypophysectomized rat

Reference(s) Braw et al., 1981 (214) Bill and Greenwald, 1981 (215)

Braw and Tsafriri, 1980 (216) Uilenbroek et al., 1980 (217) Terranova, 1980 (218)

Peters et al., 1975 (219) Peluso and Steger, 1978 (220) Braw and Tsafriri, 1980b (221) Matson et al., 1984 (222) Schwall and Erickson, 1981 (198)

Bagnell et al., 1982 (223)

Follicular Selection / TABLE 4. Methods of inducing superovulation 1. Administration of pregnant mare serum (domestic species and laboratory rodents) 2. Effects of FSH and LH 3. Active immunization with steroids (sheep) 4. Passive immunization with steroid antisera (sheep) 5. Bovine follicular fluid (sheep) 6. Pulsatile administration of GNRH 7. Clomiphene citrate, human menopausal gonadotropin (human) 8. Effect of anti-LH changes indicative of atresia than those using intact animals. Thus, within 12 hr after hypophysectomy of proestrous rats in vitro accumulation of progesterone is significantly in¬ creased by explanted follicles; conversely, estradiol is ap¬ proximately halved (214). Morphological signs of early atre¬ sia are evident by 24 hr, with about 10% of the granulosa cells pyknotic; mitotic figures are still present, however. By 2 days after hypophysectomy, germinal vesicle breakdown and polar body exclusion occur in most oocytes and the resumption of meiosis is associated with a 95% increase in oxygen consumption by the oocyte (224). The onset of atresia is even more dramatic when PMS is neutralized by anti-PMS in the hypophysectomized hamster: Within 1 hr, serum estradiol is reduced 55% (215), and the earliest histological signs of atresia are discernible within 4 hr by a significant increase in pyknotic cells in the cumulus oophorus, from 0.3% to 23.4% (225). At 2 hr after antiPMS, cAMP increases 108% above control levels, while cGMP rises 117% at 4 hr; beginning at 12 hr, cAMP steadily declines (226). Pyknotic cells begin to appear in the mural granulosa by 8 hr, when there are still 65 mitotic figures in the largest cross section of the follicle. By 48 hr, the number of pyknotic granulosa nuclei is maximal and mitoses are absent. By 72 hr, the granulosa layer is virtually eliminated, and DNA values indicate that only 20% of the original number of cells is left in the follicle (226). The thickness of the theca shows a transitory increase at 12 hr (225), which does not persist, unlike the situation observed during spon¬ taneous atresia in the hamster. Steroid production by isolated hamster granulosa and theca parallels the structural demise of the follicle. Necrotic changes in the granulosa cells are too widespread beyond 24 hr to warrant further steroid deTABLE 5. Actions of FSH on follicular steroids e Action Increase in 3p-hydroxysteroid dehydrogenase, induction of LH (hCG) receptors Stimulates estradiol secretion by granulosa cells when incubated with testosterone Adenyl cyclase stimulation and progesterone secretion Stimulates aromatase activity Stimulates prolactin receptor formation Stimulates epidermal growth factor (EGF) and FSH receptor formation

401

terminations (200). The salient observations are that thecal shells exposed to LH produce large amounts of androstenedione and 17-hydroxyprogesterone for the first 24 hr after anti-PMS but continue to produce appreciable quantities of progesterone for at least 72 hr. This confirms previous ob¬ servations with this model. When intact follicles are incu¬ bated with 200 ng LH, cAMP also increases for up to 72 hr, presumably a response of the theca (227). Thus, the loss of C-17,20 lyase in theca of the atretic hamster follicle is a critical event in atresia, an observation consistent with several other rodent models (see below). Twenty-four hours after administration of anti-PMS, grain counts of 1 ^-la¬ beled FSH and LH on granulosa cells are reduced by 69% and 53%, respectively, of control values and receptor bind¬ ing declines to 5% and 24%, respectively, at 72 hr (228). It is noteworthy that hCG binding to thecal cells and the interstitium is maintained at the same levels throughout the 72 hr after the induction of atresia, which points to the extreme resistance of these tissues to hormone withdrawal. Another series of models deal with the administration of barbiturates at proestrus to block preovulatory surges of LH and FSH and hence extend the life span of antral follicles beyond their normal 2 to 3 days duration (Table 3; models 3-4). With these models, atresia unfolds at a slower pace. After 3 days of ovulatory delay by repeated injections of phenobarbital, early signs of atresia in the cyclic hamster are manifested by pyknotic nuclei in the membrana gran¬ ulosa cells bordering on the antral cavity and in granulosa cells of the cumulus oophorus plus oocyte changes (218). During the period of ovulatory delay, the tertiary follicles continue to enlarge, expanding from 561 p.m in diameter at proestrus to 680 |xm on the next day. A new set of follicles is recruited by 3 days of ovulatory delay, possibly in re¬ sponse to elevated serum levels of FSH. After 2 days of phenobarbital treatment, spontaneous ovulation results in about 18 ova being shed, most likely representing a com¬ posite of both delayed and new follicles. On days 2 and 3 of delay, follicles incubated in the presence of LH produce more progesterone and less androstenedione and estradiol than proestrous follicles (229). However, when explanted follicles were provided with androstenedione as a precursor, estrogen accumulation was maintained at high levels, in¬ dicating that impaired estrogen secretion by the follicle is receptors during the early stages of development Reference(s) Zeleznik et al. 1974 (351) Dorrington et al. 1975 (369) Hillier et al. 1978, 1980 (370,371) Erickson and Hsueh, 1978 (347) Wang et al., 1979 (372) EGF: Jones et al., 1982 (373) FSH: Richards et al., 1976 (335), Ireland and Richards, 1978 (374)

402

/ Chapter

11 TABLE 6. Follicular development in the human ovarya

Class of follicle

Description and time of entry into class

1

Preantral with theca: Cycle 1 Luteal Beginning antrum: Cycle 2 End follicular phase Antral: Cycle 3 Follicular phase Antrum: Cycle 3 Midcycle phase Antrum: Cycle 3 Late luteal phase Antrum: Cycle 4 Early follicular phase Antrum: Cycle 4 Midfollicular phase Preovulatory: Cycle 4 Late follicular phase

2

3

4 5 6 7 8

Mitptic index of granulosa cells

Time in each class (days)

Atretic follicles'3 (%)

190-240 |xm 3-5 x 103

3.8

25

23.6

400 (xtn 15 x 103

4.1

20

35.4

1 mm 75 x 103

5.3

15

15.3

2 mm 375 x 103 5 mm 1.9 x 106 10 mm 9.4 x 106 16 mm 47 x 106 20 mm 60 x 106

8.5

10

24.2

10.1

5

58.0

10.6

5

76.8

10.7

5

50

5.2C 0.5"

5

0

Diameter and number of granulosa cells

aAfter refs. 42, 493. ^Throughout cycle. cBefore LH surge. "After LH surge.

not attributable to loss of aromatase activity but rather to a deficiency of androgen precursor. The in vitro ability of the hamster theca to respond to LH by producing androstenedione decreased, but with a concomitant increase in pro¬ gesterone accumulation, as ovulatory delay was lengthened (199). Determination of FSH and LH (hCG) receptors in delayed follicles revealed that LH binding increased slightly or re¬ mained essentially unchanged during 3 days of delay (230); FSH binding decreased steadily throughout delay. Binding favored LH, and therefore, an increase in receptors may account for the increase in progesterone secretion in delayed follicles. Similar results were obtained in the proestrous rat injected with pentobarbital (model 3). After 3 or 4 days of Nembutal treatment, most of the oocytes were in the dictyate stage, but pyknotic nuclei, as well as mitotic figures, characterized the membrana granulosa. Meiosis-like changes in the oocyte were observed by 4 days of ovulatory delay (216). Stage I atresia existed by day 3 and stages I and II by day 4. By day 4 of treatment, injection of hCG resulted in the ovulation of 3.4 ova per animal compared with 11.0 on proestrus. A more drastic regimen involved Nembutal injected at proes¬ trus (day 0) along with 4 mg progesterone (231). Preovu¬ latory follicles regressed 2 days later and were being re¬ placed by a new set of follicles, capable of a full ovulatory response to hCG (10.8 ova) by the next day. The reason for the rapidity of replacement of the preovulatory follicles

was a drastic curtailment in LH secretion and a second surge of FSH on day 2. Before morphological signs of atresia are apparent in the pentobarbital-treated rat, steroid secretion is modified within a day, as evidenced by estradiol and androgen accumulations of about 20% to 25% of proestrous values (216,217). On the other hand, progesterone accumulation over a 4-hr pe¬ riod is unaffected (217). One day after ovulation is blocked, specific binding of hCG to follicles is significantly increased and FSH binding is comparable to proestrous values. Changes in receptor numbers therefore occur as a secondary event in this model of atresia as well as in others. In vitro ste¬ roidogenic activity of the delayed ovulating rat follicles duplicates the pattern in the hamster: Accumulation of an¬ drogen and estradiol is drastically decreased by 1 day of Nembutal treatment, and addition of testosterone to the me¬ dium leads to a fivefold increase in estradiol (217). Mea¬ surement of steroidogenic enzymes by 3H-exchange assays showed that 1 day of Nembutal treatment significantly re¬ duced C-17,20 lyase and 17a-hydroxylase, with a fall in aromatase activity by day 2 (232). Again, the unavailability of androgen substrate, presumably thecal in origin, was one of the first biochemical markers of atresia. When bromocryptine—a dopaminergic agonist—is injected daily along with pentobarbital, the ability to ovulate in response to hCG is prolonged for as long as 3 days, but normal follicular structure and the in vitro ability to secrete high levels of estrogen are maintained (233). It is presumed that bromo-

Follicular Selection / cryptine acts by decreasing PRL secretion, which may have direct inhibitory effects on follicular secretion of estrogen. As enumerated several times in this chapter, the secretion of high levels of estrogen is sine qua non for normal tertiary follicles. The pattern of atresia in rat follicles after pento¬ barbital administration parallels the sequence observed in spontaneous atresia with a delay of about 2 days in all events (234). During the cycle the life span of antral follicles in the rat can be extended to a total of only 5 to 6 days before regressive changes become evident. Another group of models of induced follicular atresia involves the injection of immature animals with PMS, which leads to the waxing and waning of preovulatory antral fol¬ licles (Table 3; models 5-7). For example, 24 hr after in¬ jection of 22-day-old mice with 5 IU PMS there is no change in the total number of “large” follicles, but the balance between healthy and atretic follicles is shifted so that only 33% of the population is atretic compared to 76% in controls (219). Similarly, administration of 15 IU PMS to 26-dayold rats does not alter the total number of preantral and antral follicles present 24 hr later but decreases the pro¬ portion between nonatretic and atretic follicles (221). Both of these studies emphasize that PMS “rescues” follicles from atresia in mice and rats, but species differences and the dose of PMS administered suggest recruitment of follicles as another important role for the hormone in inducing super¬ ovulation (see below). In 26-day-old rats, follicles capable of ovulating to hCG are present 48 hr after IP injection of 5 IU PMS, and they then rapidly degenerate by 60 to 72 hr (235). The follicles can no longer ovulate 72 hr after a challenging dose of hCG; by 60 hr, acid phosphatase begins to build up in the membrana granulosa. By 60 hr after PMS, LH binding is reduced by 46% (235), whereas FSH binding is unaffected until 56 hr (220). Incorporation of [3H]thymidine into the total ovary did not vary between 48 and 96 hr after PMS, but the labeling index of antral follicles fell significantly at 96 hr; this marked the onset of early atresia as judged by the appearance of pyknotic granulosa cells (220). The critical role of FSH in maintaining an optimal environment for the granulosa cells is evident. Ultrastructurally, the earliest signs of atresia are evident by 72 hr; 28% of tertiary follicles are atretic, as evidenced by focal areas of degeneration in gran¬ ulosa cells, while others are normal (168). By 96 hr all antral follicles contained at least two pyknotic nuclei and microvilli were diminishing. Thus, cell-to-cell communi¬ cation is disrupted. Ovarian concentrations of testosterone and estradiol are significantly lower by 72 and 96 hr com¬ pared to 48 hr after PMS (168). By 72 to 78 hr, immature hamsters (25 days old) injected with 40 IU PMS—a superovulatory dose—show normal in vitro follicular outputs of progesterone, 17-hydroxyprogesterone, and estradiol in response to LH (222). After 96 to 102 hr, advanced atresia (established histologically) had affected antral follicles in some ovaries, correlating with significantly higher accumulations of progesterone and greatly

403

elevated plasma levels of the hormone or use in vivo. An estrogen-progesterone shift is thus demonstrable compa¬ rable to the changes observed in healthy tertiary follicles on proestrus after the preovulatory release of gonadotropins. The above models concentrated on atresia induced in antral follicles. Another one deals with preantral follicles and emphasizes thecal changes. The model involves hypophysectomized 21-day-old rats with Silastic implants of diethylstilbestrol (DES), which are removed from half the animals on day 24; necropsy is performed 2 and 4 days later (198). In the animals in which estrogen levels are main¬ tained, numerous preantral follicles with fibroblastic thecal cells are present. In contrast, after DES withdrawal the thecal cells hypertrophy and develop the ultrastructural fea¬ tures of steroidogenic tissues. It is fascinating that these thecal changes occur in a presumably anhormonal environ¬ ment. After DES withdrawal, hCG binding and the number of 3P-hydroxysteroid-reactive cells increase, presumably re¬ flecting thecal and interstitial cells. Four days after estrogen withdrawal, when follicular activity is maximal, in vitro steroidogenesis in the presence of hCG is greatly enhanced for progesterone, 20-dihydroprogesterone, and androstenedione. In both control and estrogen-withdrawn cells, tes¬ tosterone, DHT, and estrogen are undetectable. The increase in androgen production by the theca-interstitium is therefore apparently a secondary event in atresia of preantral follicles. Considerable attention has been devoted to a possible atretogenic role of androgens, as opposed to the antiatretic ef¬ fects of estrogens (236-238). For example, in PMS-hCGtreated immature rats, pretreatment with the antiandrogens cyproterone acetate or flutamide reduces the ovulation rate to hCG by 50% (239). Ovarian concentrations of estradiol, testosterone, and progesterone, however, are unaffected by flutamide; a better endpoint would be in vitro accumulation of steroids by the antral follicles. In control rats, 76% of the follicles >500 (Jim were healthy 48 hr after PMS; in¬ jection of either flutamide or Cl 628 (an anti-estrogen) de¬ creases the number of nonatretic follicles to 14% and 9%, respectively (239). Granulosa cell viability, assessed by try¬ pan blue exclusion, is reduced and is 20 times more sensitive to the anti-estrogen than the anti-androgen. Since granulosa cells have specific androgen receptors (240,241), it is likely that the atretogenic actions of andro¬ gens are exerted directly on these cells. Interestingly, tes¬ tosterone reduces the availability of estrogen receptors (242). A role for DHT in inducing atresia has been the subject of several studies. Thus, DHT administered to hypophysectomized PMS treated rats causes a significant reduction in primary, secondary, and tertiary follicles, and coadminis¬ tration of estrogen reverses this effect (223). However, using the same animal model and the same protocol a recent study failed to demonstrate a direct atretogenic effect of DHT (243). In the immature rat DHT reduces the formation of LH receptors induced by FSH (244,245). Since DHT does not alter FSH-stimulable adenylate cyclase and estradiol production, it is postulated that it acts as an anti-estrogen

404

/ Chapter 11

by blocking the action of estradiol on the estradiol receptor, thus promoting atresia. Estradiol is required for FSH action in rat granulosa cells (246). Another possible mechanism of action of DHT is to block aromatase activity in rat gran¬ ulosa cells (247). It has been suggested that if androgen production exceeds the ability of granulosa cells to aro¬ matize it to estrogen, then DHT will be formed and perhaps lead to atresia (8). Excessive LH stimulation of the theca of small rat follicles increases DHT significantly, whereas the granulosa cell population is great enough in large fol¬ licles to convert most of the androgen to estrogen (247). At least for the rat, a proper balance between LH stimulation, androgen production, and aromatization are necessary to promote estrogen formation and the prevention of atresia. In other species, such as human and sheep, the presence of high levels of androgens in follicular fluid of atretic follicles may not be causative but merely represent the accumulation of large amounts of precursors because of insufficient aro¬ matase activity. Collectively, the rodent models of atresia show a con¬ sistent pattern. The morphological changes duplicate those observed during spontaneous atresia. The steroid profiles demonstrate a fall in estrogen as the primary steroidogenic effect, attributable to a shut-down in thecal androgen se¬ cretion. The same changes prevail in the normal preovu¬ latory period of rat and hamster (for literature, see ref. 248). In contrast, comparable to the changes during atresia, in the proestrous ewe the abrupt fall in serum levels of estradiol is not associated with a concomitant decline in androgens (249). Species differences may therefore exist regarding the hormonal basis of estrogen withdrawal. A charge frequently leveled against models of atresia is that they may not accurately reflect events in normal, spon¬ taneous atresia. However, an unpublished study by Greenwald involving the hamster indicates excellent agreement between the morphological and hormonal changes encoun¬ tered during induced and spontaneous atresia. The experi¬ mental design consisted of dissecting the 10 largest follicles from one ovary of intact cyclic hamsters for each day of the cycle. The follicles were incubated for a baseline 1-hr period followed by the addition of LH for another hour. The media were saved for determinations of steroids and the follicles then prepared for histological examination to assess whether they were healthy or atretic. Follicles with the earliest signs of atresia have steroid profiles comparable to healthy follicles and normal vascularity, as judged by the number of red blood cells present in thecal capillaries. When approximately a third to a half of the membrana granulosa cells are pyknotic and degenerating, a concurrent fall in both androstenedione and estradiol accumulation is apparent in the baseline and LH-stimulated incubations. Thecal vas¬ cularity is then also drastically reduced. All of the aforementioned models of atresia have dealt with polytocous species. A model for inducing atresia in rhesus monkeys is based on exposure to elevated levels of estradiol-17 (3 in Silastic implants for 24 hr (250). Aftertreatment at day 5, the contents of the single dominant

follicle are aspirated on day 10. Follicular fluid concentra¬ tions of estrogen and progesterone were reduced three- and sevenfold, respectively, from control follicles. In light of the previous discussion on spontaneous atresia in the human, it would have been worthwhile to also measure androgens. The viability of granulosa cells had already diminished in the estrogen-treated follicles, but treatment with human FSH restored progesterone to control levels. At the end of 3 days of culture, the percentage of granulosa cells binding [125I]hFSH did not differ in control and treated animals.

FOLLICULAR DEVELOPMENT AND SUPEROVULATION An experimental increase in ovulation rate is an important procedure for analyzing follicular regulation in laboratory species, farm animals, and humans for in vitro fertilization programs. For our purposes, superovulation is defined as an approximate doubling in the normal ovulation rate. Table 4 (p. 401) lists various methods that have been used, all of which act by affecting either exogenous or endogenous lev¬ els of gonadotropins. Three mechanisms of action have been suggested for the effects of gonadotropins on follicular de¬ velopment: (a) Follicles already undergoing early atresia are presumably “rescued” as a result of vigorous mitotic activity in granulosa and/or thecal compartments; (b) smaller healthy follicles are recruited into a more active growth phase; and (c) the rate of follicular atresia is reduced. The last two mechanisms are not mutually exclusive and, indeed, are the most likely combination accounting for superovulation. Since its isolation some 50 years ago, PMS has been the most extensively utilized gonadotropin for inducing super¬ ovulation in laboratory animals and large domestic species (251) . This unique molecule is structurally akin to hCG (252) and functions almost exclusively as an LH-like hor¬ mone in the mare and stallion (253,254). In other species, however, PMS serves in a dual capacity as an FSH and LH molecule (253-256). In one study using radioreceptor assays for FSH and LH, the molar ratio of FSH:LH was 0.20 in pig, 0.25 in rat tissue, and 0.0 in the horse (256). Rat, cow, and pig gonadal tissues bind as much labeled PMS as LH on a molar basis, whereas equine tissues bind only 4% as much PMS as LH, or less (253). For this reason, several investigators have substituted “equine chorionic gonadotro¬ pin” for PMS, but the latter designation may be too firmly entrenched to be displaced. The FSH:LH bioactivity ratio of PMS is almost impos¬ sible to evaluate because of the various endpoints that have been used in various studies. Equine pituitary LH also has significant FSH activity in rat and pig (254,256). Two other features of PMS contribute to its efficacy in stimulating follicular growth: its long half-life, attributable to its high sialic acid content (252), and its ability to increase choles¬ terol side-chain cleavage, cholesterol esterase activity, and cytochrome P-450 when injected into immature rats (257), and therefore overall increased steroidogenesis in rabbits (258), hamsters (259), and other species. At the same time,

Follicular Selection / in large domestic species, increased steroid levels in re¬ sponse to PMS can be detrimental because of disturbances to the hypothalamic-pituitary axis. The effects of PMS on follicular development have been established for several species. Administration of PMS on day 2 or 3 of the ovine cycle results in the formation of numerous large follicular cysts (S320 mm), which ultimately become luteinized without ovulating (260). If, however, PMS is injected on days 5 to 7, an average of 7.3 luteinized follicles with stigmata (indicative of ovulation) are present 9 to 10 days later and, concomitantly, the number of cystic follicles is reduced. For a number of species, including sheep, the follicular population of the two ovaries of one animal show considerably less variation than between-animal variation (sheep: 261; hamster: 262; heifer: 263), and a number of studies have taken advantage of this feature in their experimental design. In one investigation, sheep were either unilaterally ovariectomized on day 12 of the cycle (controls) or were injected with PMS and the other ovary removed 24 or 40 hr later (264). In the control group, 74% of the follicles were between 2 and 2.9 mm in diameter and 7% were between 3 and 3.9 mm. Twenty-four hours after PMS, only 50% of the follicles were in the smallest size category, and 24% were in the 3- to 3.9-mm range. More¬ over, between 24 and 48 hr after PMS, the number of healthy follicles >2 mm was substantially increased, but the number of atretic follicles was unchanged. It was concluded that either recruitment and/or reduced atresia account for the increased number of large follicles after PMS. Similar conclusions were reached for heifers that were unilaterally ovariectomized on day 7, were immediately in¬ jected with PMS, and then had the remaining ovary removed 148 ± 23 hr later—after the onset of the preovulatory surge of LH (263). Follicles >70 p.m were counted and assessed for normality or early to late atresia. An antral cavity began to form between 115 to 280 fxm. Unilateral ovariectomy alone did not affect the number of normal preantral or antral follicles or the number of atretic follicles. After PMS, the remaining ovary significantly increased the number of preantral follicles and follicles with incipient formation of an antral cavity; the number of antral follicles was not affected. For the first two categories of follicles, PMS approximately doubled the mitotic index without changing that for normal or early atretic antral follicles. PMS treatment delayed an¬ trum formation in follicles 5 mm contained significantly fewer granulosa cells than control ovaries, suggesting that the antrum was correspond¬ ingly larger in the former group. The investigators believed that some follicles were “rescued” from early atresia because abundant pyknotic cells were sometimes present in fresh corpora lutea and especially in luteinized follicles. The above findings on increased numbers of preantral and early antral follicles after PMS lend more credence to recruitment and reduced atresia as the factors responsible for increased num¬ bers of healthy follicles. Similar conclusions were reached in another study in which four cows were injected with PMS and the entire antral population of follicles (>3 mm) ana¬

405

lyzed 48 hr later (265). In the treated animals, 70% of the follicles were healthy or very early atretic compared to 35% in the untreated controls; counting only healthy follicles, the percentages were 38.4% and 16.2%, respectively. Re¬ peated PMS-induced superovulations in cows did not affect the ovulation rate or viable embryos recovered (265), and similar results have been obtained in the hamster (266). Hence, antibody formation to repeated exposure to PMS is not a problem. The ovulatory response in PMS-treated cows is enhanced by injection of PMS antiserum on the day of standing heat (267). The number of corpora lutea increased to 15.7 compared to 9.4 in PMS-treated controls, and the number of large unruptured follicles decreased from 6.5 to

2.8. The hamster is the rodent species par excellence for eval¬ uating the effects of PMS on ovulation and follicular de¬ velopment (268). This stems from the precision of its 4-day cycle and the ease with which spontaneous superovulation can be induced, in contrast to the rat (see below). On day 1 of the cycle (estrus), each ovary of the hamster has nor¬ mally recruited 10 developing follicles that are large prean¬ tral stages. In the ensuing days, the preantral follicles mature into antral stages, and the number per ovary is reduced between days 3 and 4 (morning of proestrus) to approxi¬ mately 5, thus accounting for the normal ovulation of about 10 ova. When a small dose of PMS (5 IU) is injected on day 1, the hamsters ovulate 20 eggs by preventing atresia of the developing follicles, normally eliminated between days 3 and 4. In the light of recent findings on the dual FSH and LH actions of PMS on the rodent ovary, the ques¬ tion arises of whether one or both of these gonadotropin activities save the developing follicles. With administration of increasing doses of PMS on day 1, a plateau of 70 ovu¬ lations is reached with 30 or 60 IU PMS. This results from the recruitment of smaller reserve follicles that would nor¬ mally have taken several cycles to reach large preovulatory stages. Within 24 hr of the injection of 30 IU PMS, the hamster ovary contains 27 ± 4 follicles >267 p,m, as com¬ pared with 14 ± 3 follicles in controls, and the combined developing and reserve follicles are maintained for the rest of the cycle (57). In the hamster, clear-cut recruitment and subsequent reduced atresia represents the follicular re¬ sponses to PMS. This has been further substantiated: As early as 4 hr after PMS, preantral follicles with four or more layers of granulosa cells (these are the reserve follicles) are mobilized and begin to develop antral cavities (58). This also illustrates the rapidity with which follicles in the rodent ovary can be recruited by various perturbations, in contrast to the large domestic species in which follicular kinetics operate at a much slower pace. The combined effect of the developing and reserve follicles ultimately results in a threeto fourfold increase in the number of antral follicles and, concomitantly, in enormous increases in estrogen. For ex¬ ample, after 30 IU PMS on day 1 of the hamster cycle, serum estradiol on the afternoon of day 3 is 929 pg/ml, which, however, does not interfere with the normal oper¬ ation of the hypothalamic-pituitary axis and ovulation of

406

/ Chapter

11

an average of 63 ova at the end of a normal 4-day cycle (269). The effects of PMS administration to the cyclic rat are quite different. After the injection of 5 IU PMS during estrus, follicular development is accelerated, and there is a prompt appearance on the same day of approximately three times as many follicles larger than 55 x 106 p,m3 than in untreated animals; however, only the normal number of ova are spontaneously ovulated (56). Higher doses of PMS (1050 IU) recruit an even greater number of healthy follicles; even so, there is no spontaneous ovulation, although hCG treatment on day 3 causes the ovulation of 29 to 43 ova. It is presumed that rising and excessive titers of estrogens impair the proestrous release of LH and, consequently, pre¬ vent ovulation. The salient point of this study (56) is that higher doses of PMS can increase the rate of recruitment of follicles and reduce atresia in the cyclic rat. Injection of 7.5 IU PMS on day 24 to immature rats results 24 hr later in high mitotic indices in granulosa cells of all sizes of follicles, and this is already manifested in the smallest fol¬ licles studied, those with 90 cells in the largest cross section (65). This corresponds to follicles that are about 120 p,m in diameter (270), and they are therefore very small preantral stages. In addition to PMS, superovulation has been induced in several species with “purified” preparations of FSH. The most provocative finding, though, is the ability of LH to induce superovulation in cyclic hamsters and guinea pigs (69). High tonic levels of the hormone are maintained in hamsters by implanting an osmotic minipump on day 1 of the cycle. When 400 p.g of ovine LH was infused, approx¬ imately 32 ova were shed at the next estrus. Several ex¬ periments eliminated the possibility that FSH contamination accounted for these results. Even more striking, hamsters hypophysectomized on day 1 and implanted with LH main¬ tained large antral follicles through day 4. Infusion of LH into guinea pigs doubled the ovulation rate, whereas similar treatment of cyclic mice and rats was ineffective. Possibly excessive levels of estrogen may be acting as a negativefeedback influence in the mouse and rat. Continuous infusion of LH increases blood flow to the hamster ovary on day 3 and induces a depletion of ovarian histamine (271). Injection of an antihistamine reduces the number of ova shed in LH-implanted hamsters but not in controls. It is possible that increased ovarian blood flow may enhance delivery of FSH to the ovary, which seems to be necessary as a synergist for LH-induced superovulation. In the hamster, thousands of mast cells are located in the hilum of the ovary surrounding the blood vessels that enter and exit from the ovary (272). Since the LH surge on proestrus causes mast cell degranulation in the hamster (272) and LH causes ovarian histamine discharge (271,273,274), it seems plausible that mast cells and histamine are mediators in part of LH-induced superovulation in the hamster. Indeed, antihistamines block ovulation in several species by pre¬ venting follicular rupture (275-277). Several questions arise

at this point: What factors are involved in the LH-induced increase in ovarian blood flow? Do PMS and FSH induce ovulation by the same mechanism(s)? Are mast cells es¬ sential for LH-induced superovulation? Other methods to induce superovulation rely upon the experimental manipulation of endogenous gonadotropin lev¬ els (Table 4). Active immunization of female sheep against estrogens increases basal levels of LH and FSH and in¬ creases pulsatile LH release to levels encountered in ovariectomized animals (for literature, see ref. 278). The ovu¬ lation rate is increased from 1.5 to 3 by this treatment. Active immunization against androstenedione increases the ovulation rate from 1.50 ± 0.25 to 2.00 ± 0 and the num¬ ber of surface follicles >3 mm in diameter from 1 to 3, respectively (279). It has been proposed that active im¬ munization against steroids increases the ovulation rate by disrupting the normal negative-feedback effects of estradiol anti/or by a reduction in follicular atresia. A variation of this technique involves passive immunization with antisera to steroids given as a single intravenous injection on the first day of estrus (280). Controls ovulated 1.3 ova compared to 2.1 ova after administration of the estradiol antisera, and the mean number of lambs bom alive was 1.3 versus 1. This was attributable to a higher incidence of twinning in the group treated with the antisera. A mixture of antisera to estradiol, estrone, androstenedione, and testosterone was the most efficacious, increasing the ovulation rate to 2.1 and the lambing rate to 1.5. Intravenous injection of bovine follicular fluid to ewes on days 1 to 11 of the cycle increased the ovulation rate to 3.4 ± 0.3, as compared with 2.3 ± 0.3 in controls (281). The treatment significantly lowered FSH levels over the first 7 days, the levels returning to control values thereafter. Throughout the luteal phase, daily LH concentrations and pulse frequency and amplitude were significantly increased in the treated group. With the onset of induced luteolysis, the ewes injected with follicular fluid showed fourfold greater levels of FSH and about a twofold increase in LH compared with controls. It was pointed out that the treatment prevented the postestrous surge of FSH and that the hypersecretion of FSH at the onset of the follicular phase presumably accounts for the increased ovulation rate. However, since the onset of estrus was significantly delayed (89 vs. 41 hr), the return to control levels of FSH on day 8 and thereafter may con¬ stitute the time when “privileged follicles” may have been exposed to the amounts of FSH required for their ultimate selection after luteal regression. The pulsatile infusion of gonadotropin releasing hormone (GNRH) using Alza osmotic minipumps has been used to increase ovulation rate and estrus in zoo-maintained animals (282). The advantages are that synthetic GNRH seems to be universally capable of stimulating the species’ own FSH and LH; consequently, physiological stimulation of the ova¬ ries is simulated and “natural” mating behavior is elicited. Various regimens have been used to induce superovula¬ tion in humans (283), including clomiphene citrate followed

Follicular Selection / by hCG, clomiphene plus human menopausal gonadotropin (hMG) followed by hCG and hMG followed by hCG. There is still room for improvement in the methodology. A new approach involves intravenous infusion of pulsatile LHRH or FSH with a controlled LH surge, similar to the system described for zoo-maintained species. A detailed consid¬ eration of superovulation in the human is beyond the scope of this chapter. A chance discovery led to the surprising finding that a potent equine antiserum to bovine LH (anti-LH) is able to induce superovulation in several species. A single injection of anti-LH interrupted pregnancy in hamsters and rats, and at subsequent estrus, the hamster superovulated 29 ova, whereas the rat ovulated the normal number of 13 eggs (284) . Cyclic hamsters injected subcutaneously with 100 p.1 anti-LH at estrus (day 1) spontaneously ovulated 31.5 eggs after a cycle lengthened to 5 days from the normal 4 days (285) . The major effect of the anti-LH was to induce atresia of the larger preantral follicles by day 2 followed by a rebound in follicular recruitment by the next day, so that a greater than normal number of antral and intermediate fol¬ licles repopulated the ovaries. Quite distinct from the su¬ perovulatory effects of PMS, with anti-LH, serum levels of estradiol throughout the cycle are within the normal limits of control animals; hence aromatase activity is not increased. The anti-LH was only effective in inducing superovulation in the cyclic hamster when it was administered on estrus or proestrus, i.e., at times when serum levels of FSH are nor¬ mally elevated. After removal of the interfering LH anti¬ bodies from serum by Sephadex G-200 chromatography, radioimmunoassays for FSH and LH revealed normal levels of FSH throughout the cycle, but LH levels were nondetectable on day 2 and in most hamsters on day 3 (286). We had anticipated that anti-LH treatment would elicit a cas¬ tration response and consequent hypersecretion of FSH and LH; indeed, the possibility cannot be discounted that more frequent samples between days 2 and 3 might detect a re¬ bound release of elevated levels of FSH and/or LH. How¬ ever, if the results are taken at face value, the sustained secretion of progesterone for 2 days by the autonomous corpora lutea might act as a sufficient negative-feedback influence to prevent hypersecretion of gonadotropins. Our hypothesis concerning the mechanism of anti-LH’s ability to induce superovulation in the hamster is that by tempo¬ rarily eliminating LH action on the ovary, a “pure” FSH effect is manifested, thus increasing its mitogenic effects on granulosa cells (see ref. 286 for pertinent references on FSH:LH ratios). Several recent papers also point to al¬ tered FSH:LH ratios affecting follicular development, with higher amounts of LH reducing the mitogenic action of FSH. Thus, administration to immature rats of PMS with dif¬ ferent FSH:LH ratios, followed by an ovulating dose of hCG, causes reduced ovulation rates at lower ratios (255). Similarly, immature rats with mini-osmotic pumps delivering 240 p-g porcine FSH/day ovulated 69 ± 10 eggs. When FSH was held constant and in¬

407

creasing amounts of LH concurrently infused, the ovula¬ tion rate progressively declined (287). On the other hand, in hypophysectomized proestrous hamsters injected daily with 5 p-g/day ovine FSH plus 5, 10, or 20 pg LH, the number of follicles that matured or ovulated in response to hCG did not decline (288). The ability of anti-LH to induce superovulation in the cyclic hamster is restricted to its first administration; a sec¬ ond injection—even 3 months later—results in the ovulation of only 6 ova at the end of a 4-day cycle (266). Injection of normal horse serum at estrus followed 14 days later by anti-LH results in the ovulation of only 18 ova. Evidently, after the initial exposure the hamster rapidly forms anti¬ bodies to equine immunoglobulins. Injection of guinea pigs on day 12 of the cycle with 0.8 ml anti-LH prolonged the estrous cycle by 3 days and increased the ovulation rate from 2.9 to 5.6 ova (289). To our knowledge this is the only treatment, other than the continuous infusion of LH, that has increased ovulation rate in the guinea pig (69). Equine anti-bovine LH injected on day 10 of the cycle also increases the number of ova shed in cyclic ewes from 2.1 ±0.1 in control animals to 2.7 ± 0.2, with estrus being delayed by 0.6 days (290). It has likewise been suc¬ cessful in increasing the ovulation rate in pregnant mice, injected with anti-LH on day 4; on day 8 the animals (from a control line) ovulated 16 eggs compared with the normal number of 8.8 at estrus. A high ovulating strain that nor¬ mally ovulates 16 eggs ovulated 30 eggs when similarly treated (Barkley and Greenwald, unpublished data). How¬ ever, anti-LH does not increase the ovulation rate of preg¬ nant (284) or cyclic rats (Terranova and Greenwald, un¬ published data). This intriguing model deserves further study.

EFFECTS OF UNILATERAL OVARIECTOMY ON FOLLICULAR DEVELOPMENT Unilateral ovariectomy (ULO) is a time-honored proce¬ dure that has been useful in elucidating follicular kinetics in species as disparate as pigs, chickens, Drosophila (for literature, see ref. 291), geckos (292), and the California leaf-nosed bat (293). The latter species normally always ovulates from the right ovary, but following its removal the left ovary becomes active. The effects of ULO in mammals can be analyzed in terms of compensatory hypertrophy of the contralateral ovary (i.e., increased weight), representing persistence of increased numbers of corpora lutea (e.g., the rat) and enhanced fol¬ licular activity. For our purposes, the effects of ULO on follicular development within the cycle in which the pro¬ cedure was performed is a more meaningful endpoint, pro¬ viding information on how late in the cycle successful fol¬ licular recruitment and hence increased ovulation rate can be elicited. This can be contrasted with the long-term effects of ULO on follicular compensation, which involves differ¬ ent adjustments in pituitary-ovarian function.

408

/ Chapter

11

The first experiment involving ULO dates back to an often-quoted study of Hunter (294), who followed the far¬ rowing records of two sows, one of which had been semispayed. Over the first eight litters, the intact sow produced 87 young and the semispayed sow 76; thereafter, however, the control animal delivered 5 more litters consisting of 75 young. A pioneering study of Arai (295) established ovarian compensation in the rat after ULO and demonstrated that the surviving ovary had about twice as many corpora lutea and a greater number of mature follicles than the intact animal. Working with the rabbit, Lipschutz (296) proposed that ovarian hypertrophy following ULO was caused by an increased follicular development that depended on some general body factor.” He proposed one of the basic tenets of follicular selection, the law of follicular constancy; “The number of ova entering into follicular development, the rhythm of follicular development and the degree which is attained by follicular development are constant and are con¬ trolled by somatic factors outside the ovary. Ovarian hy¬ pertrophy means those integrative processes which take place in the ovarian fragment after partial castration which are not to be characterized by the increase in weight, but only by those processes which are dictated by the Law of Follicular Constancy” (296).

Short-Term Effects of ULO The first species in which the immediate effects of ULO on compensatory ovulation were established was the ham¬ ster (262). Removal of one ovary at 9:00 a.m. for the first 3 days of the 4-day estrous cycle was followed by a doubling in the number of ovulations from the remaining ovary. It was initially believed that a reduction in atresia of larger follicles between days 3 and 4 spared follicles and therefore resulted in compensatory ovulation (262). Recently, how¬ ever, it was shown that ULO on day 3 within 4 hr mobilizes preantral follicles with six to seven layers of granulosa cells and converts them into small antral follicles (297). This puts the hamster in line with other species (see below) and also points to the rapidity with which follicles can respond in species with short estrous cycles. Compensatory ovulation in cyclic rats after ULO depends on the length of the estrous cycle: in rats with 4-day cycles, ULO as late as day 3 results in a doubling of the number of ovulations from the remaining ovary. Slightly more lee¬ way exists in rats with 5-day cycles, in which removal of one ovary as late as 2 a.m. of diestrus 2 results in follicular compensation (298). Increased proliferation of smaller fol¬ licles results in a doubling of the number of large follicles following ULO and consequently maintains the normal ovu¬ lation rate characteristic of the rat (299). Compensatory ovulation does not occur after ULO of 4-day cyclic rats at 1700 or 2000 hr of diestrus, although a surge in FSH begins 6 hr later (300). However, if a day is added to the cycle by treatment with pentobarbital on proestrus, the animals do compensate by reduced follicular atresia, as well as by re¬

cruitment of smaller follicles. Mice unilaterally ovariectomized at random times during the cycle invariably double the number of ova shed within 3 days after the procedure (301). As an induced ovulator, the rabbit’s response to ULO is of interest. Within 48 hr of ULO, the number of follicles S=1 mm increases two- to threefold in the remaining ovary, and injection of hCG 4 days later results in ovulation of 11.4 ova (302). What are the effects of ULO in species with long estrous cycles? The guinea pig can compensate as late as day 10 of the cycle by doubling the ovulation rate from the re¬ maining ovary; an increased rate of transformation of smallersized follicles into larger ones accounts for the results (303). In contrast, ULO of Finn-Dorset sheep on day 2, 8, or 14 does not affect the number of ova shed (measured by number of corpora lutea) at the very next estrus (304). On the other hand, unilateral ovariectomy of Leicester-Merino ewes at day 14 results in compensatory ovulation by the next cycle (ULO, 2 corpora lutea; intact, 1.70) but is only partially effective when hemicastration is deferred to day 16 (305). In a breed of sheep with high ovulation rate (2.4 corpora lutea), ULO on day 10 of the cycle results in an overall ovulation of 2.2 from the remaining ovary (73). Four days after ULO there is no significant increase in any size range of preantral follicles and antral follicles (73). Unilateral ovariectomy on day 2 of the porcine estrous cycle results by day 13 in a compensatory increase in the number of large follicles (5-12.9 mm in diameter) (306). Sows unilaterally ovariectomized on day 2 of the estrous cycle ovulate as many ova at the next estrus as intact animals: 18.1 and 16.0 corpora lutea, respectively (307). There are obvious species differences between the sow and the ewe. The ewe after ULO ultimately does increase its ovulation rate per ovary, but the 2-year lamb production is significantly lower than that of intact ewe: 1.35 versus 1.61 (308). Evidently, fol¬ licles cannot be recruited as readily in the ewe as in other species. In contrast, within 6 hr after ULO on day 3 of the hamster cycle, there is already a 66% depletion of stage 4 follicles (with four layers of granulosa cells) into stage 5 (five-layered follicles) (141). Moreover, by 6 hr after ULO, incubation of follicles with [3H]thymidine shows a signifi¬ cantly lower rate of incorporation than comparable follicles from intact animals (309). This points to the rapidity with which follicles can be mobilized in animals with short es¬ trous cycles. What are the effects of ULO on species that are normally monovular? In the cow, ULO on day 8 of the cycle results, by the next estrus, in a redistribution of follicular size, there being a greater representation of the 9- to 16-mm class than in controls. However, in the one-ovary group, the largest follicle is smaller than in the control, although the next largest follicle is consistently larger in the ULO group (310). The authors raise the interesting question of whether the potential for twin ovulation is enhanced in the ULO group, but to our knowledge this possibility has never been tested. Seven days after ULO of the heifer there is a significant

Follicular Selection / increase in the number of follicles 5 to 6 mm and >9 mm in diameter; this increase can be blocked by daily injection of bovine follicular fluid (311). Rhesus monkeys hemicastrated for 3 months show ovar¬ ian hypertrophy of the remaining ovary and, histologically, numerous “large follicles and corpora lutea in various stages of development” (312). Hemiovariectomized cynomolgus monkeys have more ovulatory cycles per 13 months than intact animals: 9.8 versus 5.8. Also, in 49% of the cycles, serum progesterone was >9 ng/ml, compared with 26% in the intact group (313). Ovulation was verified by laparos¬ copy on days 11 and 21 of each cycle, and there evidently was no increase in ovulation rate, which would surely have been noted. As expected, there is even more of a dearth of information about the effects of ULO in the human female. The only concrete evidence, albeit incomplete, is provided in a study by Speert and colleagues of a series of 16 patients who required a second laparotomy within 2 years after re¬ moval of one ovary for complications of tubal pregnancy (314). In 6 patients the ovary was enlarged because of luteal or follicular cysts, but the remaining ovary appeared normal in the other patients. A long-term unilaterally ovariectomized woman had only one corpus luteum in the remaining ovary (315). The availability of sonographic techniques to visualize temporal changes in large follicles now makes it feasible to explore the long-term effects of ULO in primates. It is apparent from the literature discussed so far that compensatory follicular development (and therefore ulti¬ mately maintenance of the number of ovulations character¬ istic of the species) depends on rapid recruitment of small follicles as an acute response to ULO. What hormonal changes account for such a prompt response? Before the advent of radioimmunoassays, it was proposed that ULO did not result in an increase in pituitary gonadotropins, but rather had a “sparing effect,” so that the amount of hormone normally available to two ovaries now exerted its actions on the re¬ maining ovary. What is now established beyond doubt is that the acute effect of ULO or ablation of the dominant follicle is manifested in a sharp transient increase in serum levels of FSH, with variable and less striking changes in LH. This has now been observed in hamster (316), rat (for literature, see ref. 317), rabbit (302), gilt (318), cow (311), ewe (305), and human (319). The increase in FSH after ULO is analogous to the periovulatory increase that occurs in most mammals and that is believed to recruit the next cohort of follicles. In the immediate period following ULO there is no change in serum levels of estradiol in the hamster (320) or rat (317). Hence, removal of negative steroid feed¬ back does not seem to be the major factor accounting for the transient increase in FSH. The effects of ULO on rats hemicastrated on diestrus 2 (when antral follicles are pres¬ ent) are reversed by administration of inhibin, suggesting that this may be the key ingredient removed by semispaying (317). Similarly, compensatory ovarian hypertrophy is pre¬ vented in unilaterally ovariectomized gilts by daily treatment with porcine follicular fluid (318). The authors caution that

409

their results do not rule out a direct inhibitory effect on follicular growth, although serum FSH was significantly increased in semispayed gilts injected with porcine serum. At various times it has been suggested that ovarian nerves play a role in compensatory follicular development, but ovarian sympathectomy by injecting 6-hydroxydopamine into an artificially closed ovarian bursa fails to prevent the in¬ crease in follicular number after ULO of the contralateral ovary (reviewed in ref. 321).

Long-Term Effects of ULO Following ULO of the cyclic hamster, compensatory ovu¬ lation occurs in the next 14 cycles without deviating from the normally rigid 4-day cycle (322). The hemicastrated rat, on the other hand, compensates completely for only 10 cycles and thereafter ovulates 7 to 9 ova from the remaining ovary. Comparable results for the rat were obtained by Pep¬ pier (323), who observed that rats hemiovariectomized for 3 months shed 10.9 ova from the remaining ovary, whereas by 6 months only 4.9 ova were shed. In long-term hemi¬ castrated rats (3 months), the remaining ovary when ex¬ amined at metestrus contained a similar number of follicles (>357 |xm) as intact animals (323), suggesting that follic¬ ular selection took place later in the cycle. In rats hemi¬ castrated for 20 to 30 days the total number of large antral follicles (>400 p,m) by metestrus was the same in ULO and intact rats, and it appears that this was accomplished by a significant reduction in atresia of follicles in the 350to 399-p.m category (324). A subsequent paper by Hirshfield explored the problem in greater detail (325). Semispayed rats had twice as many follicles >450 p,m at proestrus than did intact rats, thus accounting for compensatory ovulation. By metestrus and diestrus there were already twice as many follicles in the semispayed group, and this was associated with decreased atresia. Seventy days after ULO of sheep, there is a significant increase in the number of small preantral follicles (4 mm on the ovary bearing the corpus luteum than on the contralateral ovary (423). A very thor¬ ough paper (424) documents changes in follicular dynamics in Merino ewes by estimating the doubling time of granulosa cells utilizing mitotic index and an estimate of 0.43 hr for mitotic time (obtained by using colchicine). The growth rate of follicles up to 0.7 mm was slow (estimated times in classes 1 to 6, 37 days) and accelerated in larger stages: approximately 8 days to reach 5 mm. An important point is that a follicle ovulating at the end of any cycle would have been about 0.5 mm in diameter on day 6 or 7 of the cycle. Early atresia was rare in follicles < 1 mm in diameter. Following subcutaneous injection of PMS, there were twice as many follicles (>0.5 mm) 48 hr later as in untreated ewes, with the greatest increase in follicles >2.5 mm. This was associated with significantly higher mitotic indices and a concomitant reduction in atresia in the larger classes of follicles. A similar study compared follicular kinetics for two strains of ewes with different ovulation rates: Romanov and IleDe-France (average ovulations 3.0 and 1.6, respectively) (425). Only follicles with three or more layers of granulosa cells were counted, and antrum formation began at a fol¬ licular diameter of about 0.2 mm in both strains. There was

415

no difference between breeds in follicular growth rates, although there are 1.5 to 2.0 times as many follicles (>0.09 mm) in the Romanov than in Ile-De-France ewes (261). After pooling the data from the two breeds, it was estimated that the mean time for a follicle to pass from three layers of granulosa cells (—200 cells) to the start of antrum for¬ mation (5,000 cells) was 130 days and to the preovulatory stage another 45 days, for a total of about 6 months. Another study (426) compared follicular growth in a high ovulating breed, D’Man (mean ovulation rate, 2.9), and a low ovulating breed, Timahite (rate, 1.1). The major dif¬ ferences between the two are that the D’Man ewes have a greater number of follicles >1,285 p,m (large follicles with an antrum) and a lower rate of atresia (54.9% vs. 66.7%) for follicles 1,084 to 2,141 p.m in diameter. In addition, there are significantly more follicles 118 to 462 p,m (follicles beginning to develop an antrum)—a critical phase in fol¬ licular growth—in the D’Man ewe. A breed that has attracted considerable and justifiable attention is the Booroola ewe: Some animals ovulate as many as 10 or 11 eggs. There is a genetic basis for this high spontaneous ovulation rate (for literature, see ref. 427). One possible factor accounting for the prolificacy of the breed is that on day 3 of the cycle, pituitary content of FSH is approximately twice that in control ewes; however, electrofocusing reveals no qualitative difference in FSH (428). There are obvious problems in extrapolating pituitary con¬ tent to peripheral levels of FSH. However, the inhibin con¬ tent of ovaries of Booroola ewes collected at the midluteal phase is only a third the value of control animals (427), which is consistent with the possibility of higher circulating levels of FSH. The latter, however, is still to be demon¬ strated. Alternatively, the increased ovulation rate may be attributable to increased sensitivity to normal levels of go¬ nadotropins. This possibility has been explored by com¬ paring Booroola x Romney ewes ovulating 3.3 ova (F + ) to animals ovulating 1.1 (+ +) (429). The animals were injected with cloprostenol on day 10, blood samples col¬ lected 0 to 48 hr later, and all follicles >2 mm in diameter were dissected. The main finding was that in F+ ewes, follicles mature at a smaller diameter (3.5 vs. 5 mm), as evidenced by their ability to synthesize estradiol. The fact that there is no difference in androstenedione output in re¬ sponse to LH between the two strains points to differences in aromatase activity by the granulosa cells as the controlling factor. Two interesting correlations were evident: Nonatretic follicles from F + ewes had about one third as many gran¬ ulosa cells as the -I- + ewes, and corpora lutea of the F + ewes weighed 0.39 times as much as those of + + ewes. From this observation, the logical conclusion is reached that a granulosa mass from 3F + follicles equals the estrogen production rate of one follicle from the -I- + group. In comparing highly fecund F + and Booroola ewes to control flocks with ovulation rates of —1.4, the Booroolas ovulate significantly earlier (7.5 hr) but usually with no differences

416

/ Chapter 11

in peak LH concentrations or the interval between the onset of estrus and LH discharge (430). This again points to dif¬ ferences in follicular responsiveness established in the late luteal phase as the principal factor leading to increased ovu¬ lation rate in the Booroola. On day 13 of the cycle, the corpus luteum is still func¬ tional in Welsh mountain sheep, and the concentration of progesterone in ovarian venous plasma is about a 1,000fold higher than peripheral levels (431). Even at this stage, however, the largest nonatretic follicle secretes appreciable quantities of estrogen: from 145 to 390 pg/ml in ovarian venous plasma. This trend is accentuated on day 15, when, in most instances, the corpus luteum is regressing. From days 2 to 5, the largest follicle secreting estrogen has the enzyme 3(3-HSD restricted to the theca interna; after a hiatus on days 8 to 11, beginning on day 13 to 0, a presumably new set of follicles show enzymatic activity limited to the theca (100). Within 24 to 48 hr after injection of PMS on day 13, the number of 5-mm follicles increases, and concomitantly strong 3(3-HSD activity is demonstrable in the theca—but not in the granulosa—of all these follicles. A subsequent investigation revealed that in vivo exposure to PMS for only 5 min enables about 25% of follicles >2 mm in diameter to secrete estrogen in vitro (432). Length¬ ening the time of exposure to PMS to 12 hr fails to increase the proportion of estrogen-secreting follicles, consistent with the finding that for any follicular class, about a third of the follicles are nonatretic (e.g., ref. 420). Thus, for the ewe the concept of estrogen “activated” versus “nonactivated” follicles is extremely important, as only one or two of the largest follicles fall into the former category. The notable exception is between days 2 and 4 of the cycle, for example, many explanted follicles are steroidogenically active (432) at a time approximating the second surge of FSH. Secretion rates of androstenedione and estradiol are increased (but not significantly) on day 3 of the cycle (433). The concentration of FSH in follicular fluid is about threefold higher in follicles in which the estradiol:androstenedione ratio is >1, and the FSH values are comparable to peripheral levels (433). In experiments involving ewes with ovary and adnexa autotransplanted to the neck, the secretion rate between days 12 and 15 of estradiol, but not progesterone, increased within 5 min of each LH pulse; everything points to a causal re¬ lationship between the two events (419). Addition of 0.25 to 10.0 (xg LH to explanted follicles of PMS-treated ewes results in drastic curtailment of estrogen production 48 hr later, whereas progesterone production increases dramati¬ cally by 24 hr (434). However, within 4 hr of the addition of LH, a transitory increase in estrogen accumulation oc¬ curs, and this is followed by inhibition. It must be kept in mind that the amounts of LH added in vitro were comparable to estrous surge levels and not to the pulsatile pattern en¬ countered on days 12 to 14. An interesting approach combined topical autoradiogra¬ phy of [125I]hCG binding sites with determinations of fol¬ licular fluid concentrations of steroids (435). During the

luteal phase of the ewe, all follicles >2 mm were dissected, antral fluid aspirated, and the follicle prepared for autora¬ diography. Follicles fell into three categories of hCG bind¬ ing: to theca and granulosa (type I); LH receptors restricted to theca (type II); and no LH receptors in either cell type (type III). During the luteal phase, the type I follicles con¬ tained the highest values of progesterone in follicular fluid (ca. 120 ng/ml), which were 10-fold higher than the con¬ centrations of testosterone and estradiol. During the follic¬ ular phase, these follicles had the highest concentration of estradiol (120 ng/ml), and the type II follicles contained high levels of testosterone (55 ng/ml) and very low levels of estradiol (2.5 ng/ml). LH was detectable (>2.5 ng/ml) in 86% of follicles examined, there being no significant differences between small (5 mm), except during behavioral estrus (433). As is already evident, the follicles destined to ovulate in the ewe do not differentiate until about the time of luteolysis (436). This has been demonstrated during the normal estrous cycle (e.g., see ref. 431) and following induced luteolysis and other experimental manipulations. Utilizing ovarian or utero-ovarian autotransplants, with luteolysis induced on day 10, there is an abrupt two- to threefold increase in the episodic rate of release of LH, with no change in pulse amplitude; concomitantly, the pulse rate of plasma FSH was unchanged, with the pulse amplitude reduced by 60% (249). The episodic release of LH was mirrored within a few min¬ utes by a rise in estradiol and androgens. As luteolysis progressed, plasma FSH dropped because of the reduced amplitude of the pulses. A surge of FSH accompanied the LH peak, followed by a second FSH peak about 22 hr later. Very similar results are obtained in ewes with ovaries in situ and with prostaglandin-induced luteolysis (with or with¬ out PMS) on days 9 and 10 (437). The ewes were ovariectomized from 0 to 48 hr after the onset of luteolysis and all follicles >1.0 mm in diameter dissected; the follicles were classified as healthy or atretic based on whether they con¬ tained >50% or =£50% of the maximal number of granulosa cells for a given follicular diameter. It was not until 10 hr after the induction of luteolysis that a large estrogen-acti¬ vated follicle (S:5 mm in diameter) was consistently present. Analysis of the percentage of healthy follicles by diameters suggests that the dominant follicle is capable of doubling the number of granulosa cells in 6 to 10 hr. As the dominant follicles rise to prominence, there is a steady fall in the number of healthy follicles (>1 mm) from 50% at time 0 to less than 20% by 24 hr; this decline is possibly related to the decrease in FSH levels. Injection of PMS temporarily delays at 10 hr the fall in follicular numbers, apparently by recruiting smaller stages; by 24 hr, however, the percentage of healthy follicles reverts to control levels. Another ap¬ proach to identifying the ovulatory follicle is to synchronize estrus by using progesterone implants; removal of the im¬ plants sets in motion the same set of changes already de¬ scribed following spontaneous or induced regression of the corpus luteum (438). As the preovulatory period progressed,

Follicular Selection / the dominant follicles were larger and had greater con¬ centrations of estradiol in follicular fluid and increased numbers of LH/hCG receptors in theca and granulosa cells than did nonovulatory follicles. Suffolk sheep have an ovulation rate of 1.3, whereas Finnish Landrace ovulate 2.7 ova. Correlated with these rates, Suffolk ewes have 1.2 follicles with hCG binding to both theca and granulosa cells compared with 2.9 follicles in Finnish Landrace (438). It is apparent from these studies that the differentiation of the dominant follicles is associated with increased LH levels but with FSH unaffected or, if anything, reduced. In support of this contention, intravenous administration of LHRH antiserum in the preovulatory period promptly elim¬ inates the pulsatile release of LH without affecting basal levels; the pulsatile release of estradiol is abolished with no change in secretion rate of testosterone or plasma levels of FSH (439). Anestrous ewes have 35 follicles >1 mm di¬ ameter, whereas there are 24 follicles on days 9 to 10 of the estrous cycle (440). The anestrous pattern is associated with elevated levels of FSH and progesterone and, con¬ versely, a reduction in the number of high-amplitude LH peaks (2.4/9 hr vs. 5.3/9 hr on day 10 of the cycle). Con¬ tinuous infusion of low doses of GNRH in anestrous ewes resulted in 23 of 24 ovulating, with estrus detected 37 hr, on average, after the start of infusion (441). LH concentra¬ tion in sera significantly increased; FSH values were not reported. The corpora lutea induced by GNRH were func¬ tional for at least 9 days. Intravenous pulse administration of LH (10 (xg every 1 or 2 hr for 29-91 days) to anestrous ewes simulates pulse rates comparable to those in the luteal and follicular phases (440). This regimen is sufficient to induce normal cyclic progestational activity for at most two cycles; the period of subsequent acyclicity is associated with reduced FSH concentrations. Injection of anestrous ewes with bovine follicular fluid lowered plasma FSH concen¬ trations by 70% and reduced the number of healthy follicles S:2 mm in diameter. Hourly intravenous injections of 50 (xg FSH plus follicular fluid reversed the effects and sig¬ nificantly increased the number of large nonatretic follicles (^3 mm) (442). Collectively, these results suggest that following the onset of luteolysis, pulsatile release of LH is the controlling factor in the emergence of the dominant follicles, with FSH playing a strictly subordinate role. This does not rule out a possible primary role of FSH at earlier critical stages of follicular maturation. Indeed, injection of 500 IU PMS along with prostaglandins increases the ovulation rate in Romney ewes (437). Moreover, infusion of FSH from 24 hr before until 60 hr after induced luteolysis in the ewe results in a mean ovulation rate of 8, as compared with a rate of 3 for animals in which infusion was started concurrently with the induc¬ tion of luteolysis (Baird and Webb, personal communica¬ tion, cited in ref. 436). Similarly, after prostaglandin-induced luteal regression, intravenous injection of bovine follicular fluid significantly decreases FSH levels without affecting

417

the number of ovulations (443). Infusion of ovine FSH for 48 hr plus injection of follicular fluid resulted in an ovulation rate of 14.6 ova.

FOLLICULAR DEVELOPMENT IN THE COW Following the estrous surge there is a very definite second peak of circulating FSH, lasting for 2 to 3 days (212,444). FSH levels have been reported to peak on days 4, 8, 12 or 13, 17, and 18, but these results are based on relative per¬ centage changes (444). Schams et al. believe these peaks are temporally related to periods of enhanced follicular growth, but this is pure conjecture. During the early luteal phase (day 4), 8.5 pulses of FSH are recorded per 12 hr compared with 6.3 on day 11; pulse amplitude is also unchanged: 18.5 vs. 16.6 ng/ml (445). Separate FSH pulses (with minimal LH increases) are encountered about 41% of the time in the midluteal phase; these are concomitant with or soon fol¬ lowed by pulses of progesterone (445). With the onset of luteolysis, no change in plasma FSH is observed for as long as 12 hr (446,447), but the levels fall significantly by 24 to 36 hr (446). The second postestrous peak in FSH on day 4 is not associated with an increase in LH. On day 4, there are 8 LH peaks per 12 hr, the number falling to 3.6 on day 11 when progesterone levels are maximal (445). Ninety-six percent of all LH/FSH pulses are followed within 60 min by a peak in estradiol (445). With the onset of induced luteolysis, a significant increase of mean plasma LH occurs within 1 to 3 hr; this is related to a 4-fold increase in pulse frequency and amplitude (445,447). During the luteal phase, plasma estradiol rises to a peak of 6 pg/ml on day 6 (448), and there are 7 pulses of estradiol per 12 hr; by the midluteal phase, this value has fallen by half, in conjunction with the change in LH pulse frequency (445). Following induced luteolysis, the pulse frequency for estradiol increases within 4 to 6 hr but without changes in pulse amplitude (446). In healthy preantral follicles, the principal thecal androgen pro¬ duced in vitro is androstenedione, with about a fifth as much testosterone present (149). During the early follicular phase, the average width of the theca interna is considerably thinner than in the preovulatory follicle (72.8 vs. 237 |xm) (449). Before the LH surge, 3(3-hydroxysteroid dehydrogenase is presumably deficient in cow granulosa cells, as is the case in sheep and numerous other species. A slow increase in peripheral estradiol levels begins before progesterone falls (448). Progesterone levels in the cyclic cow begin to increase from about day 4, reach a plateau around day 10, and decline to basal levels by day 17 or 18 (444). Luteal phase levels of progesterone average about 4 ng/ml (445,447), with pulse amplitude and mean concentrations peaking at about day 11. Progesterone drops very rapidly after induced luteolysis, falling 67% by 20 min (446). An inverse relationship exists between progesterone and LH levels in the cow (450), as in the ewe. Insertion of vaginal coils loaded with proges-

418

/ Chapter 11

terone into cattle during the luteal or follicular phases usually lowers serum LH, but FSH is unaffected. On removal of the coils and injection of PGF2c(, serum LH increases but not FSH. Hence, a case can be made for FSH being con¬ trolled by a separate releasing hormone or, more likely, by the secretion of inhibin (450). How do these cyclic changes in hormone levels relate to follicular development in the cow? When the largest follicle was marked between days 9 and 17, none ovulated; the likelihood of the largest follicle ovulating increased abruptly at day 18 and in 5 of 6 animals, the largest or next-to-largest follicle was selected by day 18 (451). When the largest follicle (F0 and second largest follicle (F2) are marked on different days of the cycle and examined 5 days later, the follicles are still present on days 8 and 13 but have regressed considerably by day 18 (452). Thus, between days 13 and 18, there is an increase in the rate of replacement and turn¬ over of large follicles. This is further accentuated by day 18 in that the F! follicle only ovulated when ovulation oc¬ curred within 3 days. The greatest number of small follicles (1-3 mm) is observed on day 3, presumably in response to the second FSH surge (452). Medium-sized follicles (3-6 mm) are most numerous on day 13. In contrast, the number of large follicles (>6 mm) does not vary throughout the cycle. When all follicles >5 mm in diameter are destroyed on day 13, it takes 5 days for F, and F2 follicles to emerge from the medium-sized group. Similarly, if the ovary bear¬ ing the F, follicle is removed between days 4 and 12, the remaining ovary compensates within 4 days, but if the ovary contralateral to the F) follicle is removed, follicular devel¬ opment is unchanged (453). The dominant follicle may therefore inhibit smaller follicles by an intraovarian mech¬ anism. When all grossly visible follicles are eliminated from both ovaries on day 10 by electrocautery and x-irradiation of the ovaries, luteal life span is significantly extended (454). This indicates that the normal presence of follicles during mid- to late diestrus is essential for luteolysis in the cow. The above results are consistent with earlier histological studies, which showed that 23.7% normal and 76.3% atretic follicles were present per pair of ovaries during the cycle (455). Follicles undergoing early atresia (defined in ref. 455) constituted 20.3% of the atretic population. Normal follicles >5 mm were not found during the luteal phase, whereas follicles 8 to 18 mm developed during the follicular phase. Using the mitotic time of sheep granulosa cells, the time required for bovine follicles to pass from one class to another has been analyzed (456). The time required for Graafian follicles to grow from 0.4 to 10 mm is about 22 days, the approximate length of the estrous cycle. It is therefore tempting to speculate that the postestrous FSH surge stimulates the follicles that will ultimately ovulate 20 to 22 days later. Histological signs of atresia were first observed in follicles at least 1.7 mm in diameter (456). The maximum growth rate of granulosa cells in cow follicles is reached in larger follicles (1.5 mm diameter) than in the sheep (0.10 mm), suggesting that bovine Graafian follicles need to persist longer

and grow to a greater size before they can respond to an ovulatory amount of LH. There is an excellent correlation between macroscopic and histological criteria for follicular normality (93.3%) and atresia (95.5%) in bovine follicles ranging from 2 to 22 mm in diameter (457). As to be ex¬ pected, for follicles from 2 to 5 mm in diameter, there is a progressive decline in follicular fluid estradiol as atresia proceeds from early to advanced stages (457). The number of large follicles (>8 mm) increases from day 6 (1.3) to day 12 (1.8) and day 18 (2.1), and healthy follicles with FSH binding to granulosa cells and hCG bind¬ ing to granulosa and thecal cells are already present at day 6 (458). Thus, even at day 6, follicles that could be thought of as preovulatory have differentiated, since removal of the corpus luteum or treatment with prostaglandins results in ovulation 2 or 3 days later. However, the ultimate normal fate of these follicles is atresia. On day 18, two large follicles 'are invariably present, one with considerably more estradiol than the other, the latter evidently destined to degenerate (458) . Ovaries collected from cows (with no information avail¬ able about their reproductive history) had follicular fluid aspirated from all surface follicles, and the follicles were classified into three size ranges based on antral fluid volume (459) . As follicle size increased, FSH concentration in fol¬ licular fluid was unchanged, LH decreased significantly by 27%, and prolactin increased 2-fold. These changes were associated with a shift in the estradiol:androgen ratio, such that 83% of the large follicles were predominantly estro¬ genic, as compared with only 7% of the small follicles. From days 3 to 7 of the cycle, the number of estrogenactive follicles (based on steroid determinations from fol¬ licular fluid) decreased from 2 to 1 per heifer, with a gradual increase to day 7 in follicular diameter (14 mm) and number of granulosa cells (12 x 106) (212). Through day 7, an average of one large estrogen-inactive follicle was present per heifer, which was usually judged to be atretic on his¬ tological grounds. On day 9, all follicles were estrogen inactive; by day 13, however, small active follicles (10 mm) reappeared. The estrogen-active follicle is distinguishable by specific hCG binding to both theca and granulosa cells and FSH binding to granulosa cells. This was especially evident on day 17, when receptor levels were about twice as high in both compartments compared with the inactive follicle (212). The proestrous follicle (24 hr after progesterone declined to 1 cm) have estradiol concentrations 150-fold lower than normal pre¬ ovulatory follicles (460). Thus, in the cow, as in the ewe, regression of the corpus luteum leads to the final selection of the ovulatory follicle, principally as a result of increased LH activity, after FSH and LH priming actions. Ovaries were removed from cows 2 hr after behavioral estrus and the nine largest follicles from each pair were dissected. The dominant follicle was 15 mm in diameter; others were as small as 2 to 3 mm (461). Estradiol secretion by the largest follicle in vitro was at a rate of 2 to 3 orders of magnitude greater than the smaller follicles, but small follicles ipsilateral to the dominant follicle secreted 3 times as much estradiol as the population on the contralateral ovary. Injection of porcine FSH for 4 days beginning on day 9 of the bovine cycle results in superovulation of 11 eggs per animal after induced luteolysis (462). Twelve hours after the onset of estrus, estrogen concentrations in follicular fluid were similar to those observed in normal preovulatory follicles except for much lower amounts of estradiol.

FOLLICULAR DEVELOPMENT DURING THE PORCINE ESTROUS CYCLE Serum FSH shows a minor rise, concomitant with the proestrous LH surge, and a second rise in FSH about 27 hr later, paralleling an increase in progesterone (463). Mean values of FSH are maximal on day 3, when estradiol is minimal. Contrary to the ewe and cow, the gilt shows greater pulsatile activity for FSH and LH during the luteal than the follicular phase of the cycle. On day 12 of the cycle, pul¬ satile release of LH was usually followed within about an hour by pulses of estradiol. Two separate surges of prolactin are observed during the cycle, one during the proestrous increase of estradiol and one during estrus. Maximal values of serum progesterone are reached between days 9 and 14 of the cycle, before drastically falling on day 15 (463,464). The hormonal pattern in the pig has not been studied as closely as other domestic species, but the available evidence indicates that the demise of luteal function does not increase basal or pulsatile levels of LH (463). Plasma concentrations of estrone and estradiol do not begin to increase during proestrus until plasma progesterone falls between days 15 and 18, and other than the proestrous surge, levels range from 8 to 12 pg/ml throughout the rest of the cycle (464). The selection of follicles for ovulation occurs before day 17, as evidenced by the lack of compensatory ovulation when unilateral ovariectomy is performed on day 15 or 17 (465). On day 3 of the cycle no follicles >4 mm are present in the ovaries, but by day 9 follicles between 4 and 8 mm begin to repopulate the ovary (466). On day 13, the largest follicles range from 3 to 6 mm, with an average diameter of 4 mm (467). By day 16, average diameter is 4.8 mm, and only one of six gilts possessed large follicles (6-9 mm). Injection of 1,000 IU hCG on day 12 results in the presence

419

on day 16 of both medium and large follicles in all animals with follicular estrogen and progesterone concentrations 2and 40-fold greater than the levels in control follicles, re¬ spectively. In randomly selected gilts, the right ovary on day 13 contained 48 follicles 1 to 6 mm; by day 19, there were 29 follicles in this size range and there was an average of only 1.5 large follicles (7-10 mm) per ovary (465). Fol¬ licular selection is evidently completed by day 17 since 91% of 5- to 6-mm follicles marked between days 17 and 21 are represented 6 days later by corpora lutea (468). When all grossly visible follicles (>1 mm) of Poland China gilts are destroyed by electrosurgical cauterization on day 14 (late luteal phase), the largest clear follicles present 6 days later were 8 mm in diameter (469). Thus, the follicles grew 7 mm or 0.4515 log mm3/day. Based on this figure, it is estimated that the ultimate ovulatory follicles, which are 7 to 11 mm at estrus, begin to develop around day 5 or 6 and that they require 15.6 days to complete development. Again, the second postestrous FSH increase may be an important selective factor. As in the case of the other barnyard species, the largest follicles present at metestrus (>8 mm) have significantly greater follicular fluid levels of estrogens than androgens, and this relationship persists through diestrus and proestrus (470). Similarly, prepubertal gilts treated with PMS show a daily follicular increase of 1 to 2 mm in diameter; this is associated with increasing follicular fluid concentrations of estrone, estradiol, and progesterone, comparable to the lev¬ els in the cyclic sow before and at the onset of estrus (471).

FOLLICULAR DEVELOPMENT DURING THE MENSTRUAL CYCLE OF MACACA AND HUMAN Follicles from Macaca mulatta pass from preantral to antral stages between 200 and 250 |xm (43). A significant increase in the mean percentage of follicles 100 to 200 p,m is evident in the periovulatory period, evidently influenced by the midcycle increases in steroids (estrogens?) and go¬ nadotropins (FSH?). Atresia is very limited in preantral follicles from 40 to 159 p,m in diameter, varying from less than 1% to a maximum of about 3%. Primordial follicles constitute 80 to 95% of the preantral follicles and range from 26,000 to 242,000 per pair of ovaries. In general, the mean number of preantral follicles varies directly with the size of the primordial follicle pool; similar numbers of pri¬ mordial and preantral follicles are present in the right and left ovaries (43). Very few antral follicles >1 mm are present during the luteal phase until the corpus luteum shows histological signs of regression in the premenstrual period (472). This coin¬ cides with a significant increase in serum FSH without a change in LH levels (e.g., 473,474). The greatest number of atretic follicles throughout the cycle are 0.5 to 1.0 mm in diameter, medium-sized follicles (472). During the midluteal phase, preantral follicles (=£257 p.m in diameter) in-

420

/ Chapter 11

corporate [3H]thymidine with a labeling frequency from 10% in follicles with one layer of granulosa cells to 60% in follicles with 6 or more layers (475). Hence, the failure of advanced follicular development during the luteal phase is not attributable to a deficiency in growing preantral follicles. The limited follicular development during the luteal phase is most likely due to changes in pulse frequency and am¬ plitude of gonadotropins as GNRH pulses are modulated. Thus, in measuring bioactive LH, an average of about one pulse per 90 min occurs during the early luteal phase; the frequency changes to one pulse per 7 to 8 hr during the late luteal phase (476). During the early follicular phase (days 2-5), the largest healthy follicles are 2 mm or less and there are usually two follicles about the same size. However, by days 7 to 9, one follicle was considerably larger than the other in 3 of 4 sets of ovaries, and this was further accen¬ tuated by days 11-13 (preovulatory stage), when the largest follicle was usually about 6 mm. Based on in vivo obser¬ vations, the largest follicle grows from 1.7 to 9.4 mm during the 11 days before the LH surge, and the dominant follicle, destined to ovulate, can be confidently identified in 75% of repeated laparoscopies 7 days before the LH peak and in 100% of cases by 5 days before the peak (477). As the day of the LH peak approaches, the follicle grows about 1 mm/day, as compared with 0.5 mm daily 10 days before the LH peak. Peripheral levels of estradiol increase rapidly from day 3 to 0 (the day of the LH surge), presumably reflecting the ac¬ tivity of the dominant follicle.

literature, see ref. 480), and within 4 days after luteectomy, the contralateral ovary in monkeys with both ovaries present already had more medium-sized (0.5-1.0 mm) and large (1.1-1.5 mm) follicles (473). Another study involving lute¬ ectomy revealed that on days 17 to 19 of the cycle, the new dominant follicle always originated on the side opposite the ovarian vein with the highest concentration of progesterone (481). The fact that ovulation in about 90% of cases occurred in the contralateral ovary raised the possibility that trauma following follicular ablation or luteectomy might have tem¬ porarily affected the ipsilateral ovary, and the above ex¬ periments were therefore repeated in hemiovariectomized rhesus monkeys (480). After luteal extirpation in these an¬ imals a sustained and large increase in serum FSH was always observed, suggesting that when two ovaries are pres¬ ent, the contralateral ovary is responsible for the negative feedback on tonic FSH release, now thought to be due to its production of inhibin. Collectively, these experiments led to the concept of recruitment of follicles at the end of the luteal phase and selection of the dominant follicle by days 5 to 7 of the follicular phase (for references, see ref. 482).

As follicle diameter increases from 1 to 7 mm, serum FSH decreases; thereafter, however, increased diameter cor¬ relates with rising levels of FSH. Ovulation occurs with equal frequency in each ovary, regardless of the location of the previous corpus luteum (478). By day 8 of the cycle, the ovarian venous effluent on the side of the dominant follicle already contains significantly more estradiol than the contralateral drainage (473). Even on day 8 or 12 of the cycle, the dominant follicle may contain as many as 1.2% pyknotic granulosa cells and still be normal. Thus, the pres¬ ence of one or two pyknotic nuclei is hardly grounds for classifying any follicle as atretic. As already mentioned, a technique that has yielded a great deal of useful information about follicular development in large domestic species is extirpation of follicles and corpora lutea, and it has been equally successful in primates. Cautery of the largest follicle present on days 10 to 12 of the rhesus cycle blocks ovulation, and surges of LH and FSH occur 12.4 days later (479). Hence, the follicle destined to ovulate was already selected, and no others were able to immediately substitute for its loss. Following follicle cautery, basal levels of LH and FSH (measured daily in ketamine-injected mon¬ keys) were relatively stable, although a few animals showed slightly elevated levels the next day. Following luteectomy

Presumably the higher levels of FSH in the early follicular phase aid in the selection of the dominant follicle. When charcoal treated porcine follicular fluid was injected IP every 8 hr from days 1 to 4, serum FSH decreased 50% to 80% with no change in LH (483). Following this treatment, on days 10 to 14, the dominant follicle was much smaller than usual and contained very few granulosa cells; in vitro these cells secreted negligible amounts of progesterone and did not luteinize. Moreover, within 4 days the cells became necrotic. When porcine follicular fluid was administered daily from days 1 to 3, serum FSH was depressed from days 2 to 4, followed by a rebound to elevated levels on day 6 (474). Nevertheless, the animals ovulated with a fol¬ licular phase comparable in duration to control cycles. The resultant corpus luteum, secreted significantly less proges¬ terone during the first half of the luteal phase. Porcine follicular fluid (PFF) administered from days 1 to 5 or days 6 to 12 significantly reduced serum FSH con¬ centrations and estradiol without affecting LH (484). After cessation of PFF there was a rebound in FSH levels. Treat¬ ment during menses led to a delay in midcycle FSH and LH peaks until day 17, with formation of a normal, func¬ tional corpus luteum. On the other hand, when PFF was deferred until days 6 to 12, midcycle gonadotropin surges were delayed until day 26. It appears that the early treatment deferred the appearance of the dominant follicle, whereas later treatment resulted in atresia. The dominant follicle therefore requires the continued presence of FSH throughout its development.

4 to 6 days after the LH surge, preoperative levels of LH and FSH were maintained until 12.8 days later, when typical midcycle surges of the gonadotropins were observed (479). Ovulation invariably occurred in the contralateral ovary (for

The clear opposite of reducing serum FSH levels by PFF is to increase circulating levels by treatment with hMG (484). When hMG was administered from days 1 to 3 or 4 to 6, estradiol secretion increased immediately as more fol-

Follicular Selection / licles were recruited. However, when hMG was adminis¬ tered after day 7 (when the dominant follicle has emerged), the other follicles were now unable to respond with increased estradiol secretion; presumably most of them were now atretic. Thus, after the selection of the dominant follicle, the other follicles, members of the same cohort, are unresponsive. Evidence has been presented that a follicular regulatory protein (FRP) produced by the dominant follicle suppresses estrogen production by other developing antral follicles without affecting FSH levels; it is therefore distinct from inhibin (reviewed in ref. 482; also see section below on human folliculogenesis). The maturing follicle in cynomolgus monkeys is the source of increasing titers of estra¬ diol. When, after unilateral ovariectomy (ULO on day 0), estradiol antibodies are infused from days 5 to 10, serum FSH and LH increase, and 10 days after hemispaying the remaining ovary of two animals contained two large follicles and the third contained four (485). Accordingly, it appears that the dominant follicle’s production of estrogen acts as the principal modulator of development of other follicles by its ability to suppress gonadotropin levels. This is contrary to the notion that a follicular regulatory protein produced by the dominant follicle accounts for its outstripping the other members of the cohort. Another approach to analyzing follicular development in the rhesus is to induce follicular atresia by Silastic implants of estradiol on days 5 to 7 of the rhesus cycle (for literature, see ref. 486). The resultant serum concentrations of estra¬ diol, 100 to 400 pg/ml/24 hr, led to transient declines in FSH and LH, followed by a rebound and unusually high levels of both hormones on day 8. Dierschke et al. (486) believe that exogenous estrogen acts directly at the ovary or indirectly by the transitory depression of FSH to induce follicular atresia. It seems equally plausible that the rebound in LH is responsible for atresia: Injection of hCG (1,000 IU) on day 9 or 11, prior to the spontaneous midcycle surges of LH and FSH, leads to apparent atresia of the dominant follicle, as evidenced by the absence of midcycle increases in peripheral estrogen and gonadotropins (487). Progester¬ one administration by injection or Silastic implants on day 6 of the rhesus cycle similarly affects the dominant follicle, resulting in very low levels of serum estradiol (30-50 pg/ml) throughout the course of treatment (for literature, see ref. 488). During the follicular phase of the rhesus cycle, as follicles increase in size from 3 to 6 mm to 6 to 8 mm, follicular fluid estrogen increases from 100 to 2,200 ng/ml (489), in agreement with the estrogen-activated follicles already de¬ scribed for other species. Before the LH surge in vitro pro¬ gesterone secretion and morphological luteinization of gran¬ ulosa cells from follicles less than or greater than 6 mm are minimal in 2- to 8-day cultures (489). On the other hand, follicles removed during the early LH surge (when they are 6-8 mm) or mid-LH surge (10-11 mm) show in vitro mor¬ phological luteinization of granulosa cells and significant progesterone secretion by granulosa and thecal cells. More¬

421

over, the theca (without the addition of an androgen pre¬ cursor) produces considerable amounts of estrogen, whereas the granulosa cells are inactive. These results have been confirmed in short-term (3 hr) incubations of granulosa and thecal cells from dominant follicles removed on day 12 of the rhesus cycle—2 days before the normal LH surge (490). The follicles were 5.2 mm in diameter and contained 3.32 x 106 granulosa cells. A tritiated exchange assay with [l,23H]androstenedione (a measure of aromatase activity) showed that granulosa cells produced 341 fmol 3H20, compared with 89 fmol by thecal cells. In this connection, when peripheral estradiol was >150 pg/ml, aspiration of as many granulosa cells as possible from the dominant follicle results 15 min later in a fall of about 70% in the concentrations of progesterone, estradiol, and androstenedione from preaspiration levels (491).

FOLLICULAR DEVELOPMENT IN THE HUMAN Until recently, quantitative histological analyses of the human ovary were extremely limited—for obvious reasons. It is well established that the number of primordial and growing (>100 |xm) follicles diminishes with age: primor¬ dial ages 6 to 9, 484,000 vs. 8,236 at age 40 to 44; growing ages 6 to 9, 15,220 vs. 6,190 at age 40 to 44 (41). The number of Graafian follicles (>1 mm) averaged 63 per pair of ovaries over the same time span, with only three of seven women having follicles in this size range at ages 40 to 44. Another frequently cited study dealt with follicular devel¬ opment during the menstrual cycle, with histological eval¬ uation of the endometrium used to date the ovaries (492). The heart of that paper is Table 21. Presenting the data on 17 ovaries from women aged 18 to 33 years, the table shows the gradual emergence during the late follicular phase (days 11-14) of a large viable follicle, 10 to 13 mm in diameter, whereas on day 1 of the cycle, the largest follicles are 3 to 4 mm. There are more small antral follicles 1 to 2 mm in diameter present from day 12 to 14 (15.2/ovary) than in the earlier follicular phase; the number of healthy small follicles declines in the earlier luteal phase (days 16-18) to an av¬ erage of 8.5/ovary, with the maximal number present on days 20 to 27 (28/ovary). Although a heroic amount of effort went into Block’s studies, the knowledge gained was rather limited. The situation was dramatically changed by a series of interesting and provocative findings by Gougeon (42,493). The studies involved histological analyses of ovaries of 33 women with regular menstrual cycles (28 ± 2 days) who had been ovariectomized for extraovarian pathology. Days of the cycle were judged by plasma levels of estradiol, progesterone, LH, FSH, and, in some instances, endome¬ trial biopsy. Serial sections (10 |xm) of each ovary were cut, both ovaries being available for sectioning from 22 patients. Some salient findings are outlined in Table 6 (p. 402). The follicles are classified in eight stages, based primarily

422

/ Chapter

11

on diameter and number of granulosa cells. Estimates of the doubling time of granulosa cells are calculated from in vitro determinations of the mitotic time [1.36 hr, the same time as in the sheep (424)] and mitotic index (494). Based on these estimates, the length of time spent in each class of follicular development is listed in Table 6. The mean time for a preantral follicle (stage 1) to ovulation is about 85 to 90 days, about 3| cycles. Note that the most active stages in granulosa proliferation occur in classes 4 through 7. The overall rates of atresia are also shown in Table 6. Atresia of follicles 1 to 2 mm—classes 1 through 4—is relatively constant throughout the menstrual cycle and is unaffected by cyclic hormonal changes (495). In contrast, the atretic rate for classes 5 and 6 is higher and inversely proportional to circulating gonadotropin levels; it is highest during the early and midluteal phase, when almost all class 8 follicles are atretic. Table 6 lists the overall mitotic indices throughout the cycle (494). These rates, however, are not constant, and there is an especially brisk increase for class 5 and 6 follicles at the end of the cycle (day 28) (495). Analysis of the largest healthy and atretic follicles through¬ out the cycle revealed that the dominant follicle could be identified by size during the early follicular phase, days 1 through 5, although it did not differ in vascularization or thecal or granulosa development from other healthy folli¬ cles. By days 6 and 10, the dominant follicle (now 13.3 mm) did differ in these features from other healthy follicles (315). Excluding the dominant follicle, the next healthiest follicle never exceeded 6 mm in the follicular phase or 4 mm in the luteal phase. Based on the mitotic index, follicles 2 to 5 mm in diameter (classes 4 and 5 in Table 6) are very active in the late luteal phase and represent the population from which the dominant follicle may already be selected. It is interesting that the largest healthy follicle was invariably located on the ovary contralateral to the previous ovulation (315). A subsequent study (496) showed that this phenom¬ enon occurred in 87.6% of cases. This conclusion was based on the ability to make a histological identification of the age of the corpus luteum over five cycles. If the histological criteria are valid, this would be quite contrary to previous assertions that ovulation in the human is random, uninflu¬ enced by the side of the last corpus luteum. The pattern of alternating ovulations is explained on the basis of differences in the intraovarian hormonal milieu, such that a 5-day asyn¬ chrony exists in the population of follicles developing in the “ovulating” ovary and the contralateral one (497). It is proposed that this local effect comes into play when the follicles begin to develop an antral cavity (class 2). The follicles on the side of the corpus luteum, i.e., the ovulating ovary, have a significantly higher mitotic index but are smaller than in the contralateral side; in the next cycle, class 3 and 4 follicles now develop earlier in the ipsilateral, non¬ ovulating ovary. Alternation of ovulation is an interesting and unsettling phenomenon if true: Ultrasonography should be able to show whether, in fact, it is occurring. Table 6 also shows when follicles pass from one stage to another

during the menstrual cycle, illustrating how a class 1 follicle that will ovulate approximately 3 months later is selected. It is postulated that follicles move continuously out of the resting pool into class 1 and that the growth of follicles 6 mm) have a high pyknotic index (10.9 ± 2.1%) 2 days before the LH surge. Healthy dominant human follicles show a strong reaction for 3(3-HSD in the theca and limited activity in the granulosa; it is not until after the LH surge that an intense reaction develops in the granulosa cells. Follicles in advanced atresia show low or moderate 3(3-HSD activity in the theca and almost none in the granulosa. Criteria proposed for determining a follicle’s potential for further development are whether it contains (a) more than 50% of the maximal complement of granulosa cells for its size and (b) a normal oocyte (206). Thus, a 4-mm follicle has about 1 x 106 granulosa cells and a 12-mm normal follicle has 10 x 106 granulosa cells. Based on these cri¬ teria and the concentration of steroids in follicular fluid, 90% of follicles >1 mm in diameter were undergoing de¬ generative changes. After reaching a diameter of ~4 mm, only one or at most two follicles per follicular phase of the cycle are capable of continued mitotic activity to reach the 50 to 100 million granulosa cells characteristic of the pre¬ ovulatory follicle. According to the aforementioned histo¬ logic studies of Gougeon and hormonal correlates from an¬ tral fluid (499), it is evident that the follicle destined to ovulate has normally emerged by days 1 to 5 as a healthy 4-mm follicle. On day 1 of the cycle, although the dominant follicle cannot be grossly identified, ovarian venous plasma collected from both ovaries already shows unilaterally high concentrations of progesterone, estradiol, and estrone, and this is even more apparent by days 4 to 9 and thereafter,

Follicular Selection / when a large follicle can be identified (e.g., refs. 500-502). Moreover, by the midfollicular phase, high unilateral con¬ centrations of the above steroids in ovarian effluent are also associated with asymmetry in levels of androstenedione and testosterone. Granulosa cells harvested from 6- to 9-mm follicles on day 9 produce small amounts of estradiol (2 ng/106 cells) for 3 hr with 10~7 M testosterone or androstenedione but show an eightfold increase when FSH is combined with either of the androgens (247). In contrast, granulosa cells collected from a large dominant follicle (20 mm) accumulate ~60 ng estradiol/106 cells/3 hr in the presence of testos¬ terone, and FSH has no synergistic action. Thus, the early developing follicle depends on FSH for regulation of aromatase enzymes in granulosa cells. It is interesting that 5areduced androgens at physiological concentrations can in¬ hibit aromatization of testosterone in FSH-stimulated gran¬ ulosa cells of small (5-9 mm) follicles (247). For follicles 100 ng/ml, 96% of the follicles lack detectable amounts of FSH and antral fluid concentration of estradiol is depressed (506). In ad¬

423

dition, accompanying hyperprolactinemia, follicles >4 mm were apparently undergoing atresia, based on the reduced number of granulosa cells for their diameter. In large preovulatory follicles, both diced theca and gran¬ ulosa cells appear capable of producing progesterone, an¬ drostenedione, and estradiol, with progesterone the domi¬ nant steroid secreted by granulosa cells and androstenedione the major steroid produced by theca (for literature, see ref. 508). When expressed as ng steroid/mg protein x 2 hr, thecal and granulosa cells released estradiol at approxi¬ mately the same rate in the absence or presence of hCG (508). In contrast, collagenase-dispersed preparations, in¬ cubated with a fixed amount of testosterone, show that gran¬ ulosa cells have at least 700 times the aromatase activity of the theca when expressed as steroid/105 cells/24 hr (247). Considerable differences in experimental design make it difficult to compare the two studies. Based on these findings and others, McNatty (499,506) proposed that a preantral follicle destined for further de¬ velopment is transformed to an antral stage concurrent with high levels of FSH in plasma and its antral fluid. The latter minimizes atresia by increasing mitotic activity of granulosa cells and increasing aromatase activity in the cells, thus enhancing estrogen secretion. The combined interactions of FSH and estrogen act as a positive feedback to further en¬ hance granulosa cell mitosis and estrogen accumulation in the follicle. The action of LH on the theca stimulates an¬ drostenedione, which in the favored follicle is converted to estrogens and thus further shifts the estrogen:androgen ratio. Preantral follicles not emerging when the concentration of follicular fluid FSH can be increased are therefore much more likely to be androgenic follicles and hence prone to atresia. The potential to recruit follicles for ovulation obviously encompasses larger follicles—presumably ~4 mm at the onset of the follicular phase—as demonstrated by the ability of human menopausal gonadotropin or FSH administered on days 3 and 4 to increase the number of oocytes induced by hCG administered on day 8 (for literature, see ref. 509). Injection of hMG at the late luteal phase instead of the early follicular phase is even more successful in increasing the mitotic index of granulosa cells and increasing the induced ovulation rate to 3.7 ova (510). The human—like the rhesus—responds to removal of the dominant follicle or corpus luteum by ovulating 12.7 and 14.6 days later, respectively (511). Following either pro¬ cedure, ovulation invariably occurs in the ipsilateral ovary, contrary to the situation in the rhesus. Unlike the rhesus, there is a clear-cut, transient increase in FSH and LH after follicular ablation in the human (319,512). Whether a true species difference exists or whether effects of anesthesia are responsible remains to be established. It would indeed be unusual if recruitment of the next follicle in the rhesus is not associated with transient increases in gonadotropins. The restraining influence of the human corpus luteum on gonad¬ otropin levels is evident.

424

/ Chapter

11

A protein aspirated and pooled from human follicular fluid of hyperstimulated ovaries inhibits ovarian weight increase and serum estradiol in suitably primed hypophysectomized immature rats (513). The putative FRP is heat and trypsin labile and has also been recovered from the ovarian venous drainage on the side of the dominant follicle. FRP is pro¬ duced by granulosa cells (recoverable from spent medium during the first 48 hr of incubation) and does not affect FSH binding to rat granulosa cells but does decrease their aromatase activity (514). It is speculated that FRP may be an important intraovarian regulator of folliculogenesis and may account for the selection of the dominant follicle in primates and other species (for review, see ref. 482). The substance appears to be distinctly different from inhibin based on its biophysical properties (515). Considerably more work is needed before a physiological role of FRP can be established. At present, activity is based on pooled antral fluid from hyperstimulated ovaries and sampling from individual follicles (preferably from nonstimulated ovaries) is needed to establish whether a hier¬ archical relationship exists between FRP in the dominant follicle and other members of the cohort. FOLLICULAR DEVELOPMENT DURING PREGNANCY This topic is of interest for several reasons: (a) Is there any “wavelike” development of follicles during pregnancy comparable to the length of the estrous cycle and therefore recurring every n days? (b) How does the negative-feedback effect of high steroid levels affect the hypothalamic-pituitary axis and hence follicular growth? (c) In species in which corpora lutea are essential throughout pregnancy and pro¬ duce progesterone as the main hormone, the follicles ob¬ viously are the principal source of ovarian estrogens, (d) Are follicles developing during pregnancy “mature” enough to ovulate in response to hCG? Follicular development in the pregnant mouse can be divided into three stages: (a) From day 1 to 8 (day 1 = morning after mating), the largest follicles present are up to 473 |xm in diameter and atresia of large follicles is min¬ imal; (b) from day 10 to 14, follicular atresia is maximal and the largest follicles are usually 373 (Jim or less; (c) from day 16 to 19, follicular atresia is again minimal and from day 18 and on, large follicles reappear in preparation for postpartum ovulation (516). This profile correlates with the ability to induce ovulation with 5 IU hCG. Thus, between days 12 and 14, only 37% of injected animals ovulate; the nadir is reached at day 12, when the mean number of ovu¬ lations is 5.2—only half the normal number. Subcutaneous injection of 5 IU PMS on day 10 followed by hCG on day 12 results in 91% of the mice ovulating an average of 11.2 ova (516). When pregnant mice were pulse-labeled by injecting [3H]thymidine, small to large follicles were labeled on every day, showing that there is continuous growth of follicles

(50). At any time during pregnancy about 14 small follicles begin to grow compared with 19 per day during the estrous cycle, but there is no difference in the transit time. It is interesting that the labeling index of type 6 and 7 follicles (incipient formation of antral cavity and coalesced cavity, respectively) tend to decrease during midpregnancy (cf. above results on induced ovulation with hCG). It was estimated that type 3b follicles (with 21-60 granulosa cells) moving out of the resting pool on day 1 of pregnancy are the ones that ultimately ovulate at postpartum estrus. These morphological findings show interesting correla¬ tions with the gonadotropin pattern during mouse preg¬ nancy. Using 50 fxl of plasma, there are LH peaks on days 4 and 11 of gestation with no significant changes in FSH throughout this period (517). However, from days 12 to 15, LH is frequently undetectable in the amount of plasma as¬ sayed, whereas plasma FSH does not differ from earlier values. On the day of parturition, there is a sharp increase in both FSH and LH (517) coincident with restoration of follicular kinetics to the pattern characteristic of the estrous cycle (50). Follicular growth and gonadotropin profiles are quite dif¬ ferent between the pregnant mouse and rat. Until day 14 of pregnancy in the rat, healthy follicles up to 600 jxm are present, but this is followed by a hiatus on days 16 and 18, when the largest antral follicles are only 400 to 500 (xm. Differentiation of larger follicles begins on day 20, cul¬ minating in follicles >600 (xm on day 22 and at delivery on day 23 (518). Follicles 300 to 400 |xm in diameter are present on every day of pregnancy and are especially abun¬ dant on the first 4 days of gestation. Ovulation with 20 IU hCG (sc) could not be induced before day 21, but priming with 25 IU PMS on day 15, 72 hr before hCG treatment, resulted in ovulation. It was concluded that even though medium and large antral follicles are present throughout most of gestation, the follicles were “physiologically im¬ mature.” Subsequently, several other investigators who in¬ jected pregnant rats with hCG reported different results, depending possibly on differences in strain, colony, and route of administration (519). However, the consensus was that days 15 and 16 represented the nadir in follicular re¬ sponsiveness to hCG. In this connection, it is interesting that serum levels of FSH and LH are at their lowest values on day 16, with serum LH abruptly increasing on days 20 and 22 and with FSH recovering on day 18 (520). Furthermore, on day 16 the corpora lutea of pregnancy have the highest concentra¬ tions of estradiol with the lowest levels in the nonluteal ovary, suggesting that between days 14 and 18 secretion of testosterone and estradiol represents proportionately more luteal than follicular activity. Ovarian venous plasma on day 15 of pregnancy also does not contain detectable amounts of inhibin, a further indication of the nonfunctional status of the follicles (521). Richards and colleagues have recently explored this prob¬ lem in depth, focusing on follicular development on day 16

Follicular Selection / of pregnancy. Between days 14 and 19, FSH and hCG binding reach their nadir in granulosa cells, and hCG re¬ ceptors in thecal shells are also minimal (522). This was associated with minimal accumulation of estradiol in vitro, although providing testosterone as a substrate increased es¬ trogen levels to values observed on days 4 to 12. Hence, on day 15 the aromatizing system is still intact in the gran¬ ulosa compartment. Follicular morphology on day 16 re¬ vealed that the follicles lacked the features typical of steroidogenically active tissue (523). Following daily injection on days 14 and 15 of 1.5 IU hCG, however, the theca hypertrophied and the orientation of granulosa cells was altered with extensive lipid deposition in both tissues. The most salient finding was that isolated thecal shells produced considerable testosterone in vitro after exposure to hCG: from 17 to 175 pg/thecae/5 hr (523). The action of hCG is specific in that twice-daily injections of FSH on days 14 and 15 fails to increase thecal LH receptor content or in¬ trinsic ability of follicles to accumulate estradiol (524). The small antral follicles normally present on day 16, when incubated with [3H]progesterone, fail to convert it appre¬ ciably to labeled 17a-OH progesterone or androgens, in contrast to normal preovulatory follicles on day 23 (525). Collectively, these experiments demonstrate that the relative paucity of serum LH on day 16 accounts for inadequate biochemical development of the theca with a consequent deficit in 17a-hydroxylase and C-17,20 lyase. This condi¬ tion is then reversed by rising titers of LH commencing on day 20 of gestation. Follicular development in the pregnant hamster is dia¬ metrically opposed to the pattern described for the rat. Large healthy antral follicles >415 (xm in diameter are always present in the hamster ovary, with about 10 per pair of ovaries from day 4 to 10; this number reaches a peak of 35 to 40 on day 12 (526). This is paralleled by the ability of hCG to induce ovulation at any time during gestation: The mean of 10 to 13 ovulations from days 4 to 8 culminated in a peak of 35 ovulations on day 12 (526). Hence, at midpregnancy, the hamster matures sufficient follicles to result in superovulation. Healthy follicles 277 to 322 p,m in diameter (the smallest ones measured) were continuously present throughout gestation, indicating constant recruit¬ ment of preantral stages. The concentration of FSH on day 1 of the 16-day pregnancy was about fivefold greater than the levels on subsequent days (527). When sampled at 0900 hr at 2-day intervals throughout pregnancy, serum FSH was approximately twice as great as 0900 hr proestrous values (—180 versus 93 ng/ml), and serum LH was 20 ng/ml, which was comparable to 0900 hr proestrous values (528). In con¬ trast, in the pregnant rat, serum FSH is comparable to proes¬ trous values, but LH ranges from 4 to 8 ng/ml from day 4 to 18, whereas serum LH is approximately 30 ng on the morning of proestrus (399). Thus, the relative deficiency of LH may account for the differences in follicular develop¬ ment between the pregnant rat and hamster. Hamster fol¬ licles in vitro can be stimulated by 10 ng LH at any time

425

during gestation to accumulate estradiol, although the re¬ sponse is drastically curtailed by day 16—the day of delivery (528) . Again, this is a striking difference from the rat. Follicular development in the pregnant rabbit resembles the pattern found in the hamster. The intravenous injection of 25 IU hCG up to 12 days of pregnancy results in the ovulation of 11 ova, but similar treatment on day 21 leads to the ovulation of an average of 21 eggs (529). The en¬ hanced ovulatory response to hCG is associated with in¬ creasing numbers of antral follicles 1 mm in diameter or greater: on days 8 to 11 a mean number of 14 follicles compared to day 17, when the ovaries average 23 follicles (529) . Serum levels of FSH and LH measured at 3-day intervals throughout rabbit pregnancy do not correlate with the midgestational change in follicular number (530), but the sampling intervals tested in the hamster and other species are also not frequent enough for any valid comparisons. In estrous does, approximately 50% of the follicles are 1.6 mm (531). Some of these large follicles contain elevated levels of tes¬ tosterone and progesterone but low estradiol; presumably these follicles are atretic. After inducing ovulation with hCG, only 15% of the remaining follicles are large; by day 6 of pseudopregnancy, however, there is a redistribution, with 75% of the follicles being classified as large. Correlated with the recruitment of large follicles, on day 6, the follicles now contain high levels of estradiol, testosterone, and pro¬ gesterone. Rabbit corpora lutea first become dependent on estrogen on day 6, and the large follicles recruited by the periovulatory surge of FSH on days 1 and 2, with tonic levels of LH (530), evidently fulfill this need. The pregnant guinea pig resembles the hamster and rabbit in that large preovulatory follicles (500-700 p,m) are present throughout gestation, with the peak number attained on day 66 in preparation for postpartum estrus (532,533). Ovulation can be induced at any time in pregnant guinea pigs by intravenous injection of 50 IU hCG (534); the follicles are less sensitive on days 8 through 20 than on days 41 through 62, and the average ovulation rate is about twice as great in the last trimester (2.0 versus 3.9, respectively). There is very little information available about follicular development during pregnancy in large domestic species. In cross-bred pigs, the total number of follicles up to 8 mm in diameter is greater up to 40 days of gestation, and by 110 days (shortly before delivery) there are no 7- to 8-mm follicles and few in the 4- to 6-mm class (535). Small fol¬ licles (1-2 mm) were relatively constant from days 23 to 63 in three breeds of sow (127); over the same time span, medium follicles (3-5 mm) were also fairly constant (61), with large follicles (6-10 mm) averaging 4.8 per pregnant sow (536). In the pregnant cow, 32.6% of the follicles are normal and 67.4% are atretic (455). Follicles larger than 5 mm in diameter are absent throughout pregnancy, and in late preg¬ nancy (243 days) the largest follicles are only 2 mm. Small follicles 400 p,m are absent. In contrast, a few follicles in this range are always present in dams nursing two pups (548). Follicular immaturity in rats with eight pups was reflected in very low levels of estradiol, both in vivo and in vitro. On removal of the litter on day 3 of lactation, a significant increase is evident by 30 hr in the number of follicles 201 to 600 p.m in diameter, and the animals ovulate by 96 hr (549). The hormone profiles found in rats nursing eight pups exemplify a rather universal pattern during lac¬ tation: normal cyclic serum levels of FSH, elevated PRL, and extremely reduced LH. Daily injection of 0.5 to 1.0 IU hCG or 50 |xg ovine LH from day 2 to 5, followed by an ovulatory injection of hCG, results in ovulation of the nor¬ mal complement of ova by the next morning (548). After the removal of the litter on day 3 of lactation at 1100 hr, significant increases occur 24 hr later in serum LH and FSH with a concurrent fall in PRL (549). The missing ingredient during intense suckling stimulus therefore appears to be LH. In vitro production rates of testosterone and estradiol by the nonluteal ovary are drastically reduced in rats nursing two or eight pups on day 2 of lactation, but a positive staircase

Follicular Selection / increase occurs in the former group over the next 8 days. It thus appears likely that the deficiency in LH leads to decreased androgen production by the theca and presumably reduced estrogen production by the granulosa compartment. Reduced estrogen secretion may also reflect lower aromatase activity. This does not negate the role of high levels of PRL during lactation, which may act directly or indirectly on the ovary. For example, high levels of PRL inhibit follicular estrogen production in the rat (413,550). The respective roles of LH and PRL in affecting follicular function during lactation is a theme recurring throughout this section. In addition to the high baseline levels of PRL during lactation, the suckling-induced response must also be considered. In rats suckling eight pups, the magnitude of PRL release in 20-day postpartum mothers is considerably reduced and the return to baseline levels accelerated (551). Substituting 10day-old pups with the 20-day postpartum mother does not enhance PRL release, indicating that the intensity of the suckling stimulus is not a factor. As will be seen for other species, the hypothalamic-pituitary mechanism mediating suckling release of prolactin becomes refractory with time. Rats with prolonged lactation for 72 to 105 days (produced by substituting litters at approximately 2-week intervals) and then mated usually deliver at 22 days (552). This suggests that FSH and especially LH are of greater significance in the rat than suckling release of PRL in regulating follicular development. Bromocriptine treatment of suckling rats mated at post¬ partum estrus suppresses serum PRL and progesterone lev¬ els, and the uterus contains unimplanted embryos even if progesterone is injected concurrently with bromocriptine (553) . Removal of PRL evidently fails to increase secretion of follicular estrogen, which is the key to triggering ovu¬ lation in the rat. This also suggests that the primary role of PRL is at the hypothalamic-pituitary axis, not at the follic¬ ular level. In postpartum rats nursing seven pups, diestrus lasts for 3 weeks and is associated with reduced LH secre¬ tion, hyperprolactinemia, and increased serum progesterone (554) . Daily administration of bromocriptine to lactating rats shortens the duration of diestrus to 11 days by depress¬ ing the secretion of both PRL and progesterone to baseline levels. It is well established that lactation prevents the postcas¬ tration rise in gonadotropins in the rat (reviewed in ref. 555), and with the increase in understanding of neuroen¬ docrine mechanisms, it was logical to infer that the hypo¬ thalamic-pituitary axis is a controlling factor. Indeed, there is a significant fall during lactation in pituitary LHRH re¬ ceptor concentration (556). There is no change in pituitary affinity for GNRH receptors, but there are about 50% fewer binding sites than at estrus (557). Moreover, removal of an eight-pup litter from ovariectomized mothers results in a sharp increase in GNRH pituitary receptors 24 hr later. Rats ovariectomized on day 10 show a significant increase in LH secretion by the next day; this can be blocked by exogenous PRL without, however, affecting the LH response to LHRH

427

(555). The action of PRL is therefore presumably exerted in part at the hypothalamic level. On the other hand, pituitaries of lactating rats nursing eight pups exposed in vitro to pulsatile GNRH release as much FSH but considerably less LH than pituitaries of animals nursing two pups (558). This points to an action of the suckling stimulus (PRL?) directly modifying pituitary response to GHRH. As will become apparent, changes in the frequency and magnitude of GNRH release most likely account for the low serum levels of LH during lactation. Swiss mice nursing six young show a pattern very similar to the rat. During the first 11 days postpartum the largest vesicular follicles are 350 p,m in diameter; thereafter, the follicles enlarge to 450 |xm, the corpora lutea of pregnancy regress, and the vagina becomes mucified, indications of estrogen-progesterone interaction (559). The most extreme effect of lactation on follicular devel¬ opment has been observed in the hamster. Unlike the rat and mouse, there is no postpartum ovulation in the hamster; instead, the large number of vesicular follicles developed during pregnancy quickly regress and consequently the the¬ cal cells are incorporated into the interstitium (560). The net result is an ovary characterized by interstitial hypertro¬ phy, with follicular development limited to preantral folli¬ cles with seven to eight layers of granulosa cells. This un¬ usual ovary is maintained by as few as one or two suckling young. Removal of all young on day 2 or 14 of lactation results in ovulation exactly 4 days later (560). The hormonal basis for the acyclic ovary is a daily massive release at 1600 hr of FSH and LH (561), evidently in a ratio incompatible with the differentiation of antral follicles. Concomitant with the daily gonadotropin surge, serum progesterone increases as lactation progresses. The progesterone is secreted by the interstitium, and several lines of evidence indicate its release is in response to the LH surge (561). Because of the species’ economic importance, consid¬ erable attention has been devoted to follicular development in the postpartum cow. The degree of follicular inhibition is influenced by two factors: Suckling and its intensity delay the return to estrus. For example, beef cows that were nonsuckled, suckled once daily for 30 min, or suckled ad libitum by two calves ovulated, on average, 31, 41, and 76 days after parturition, respectively (562). Another factor com¬ plicating follicular development in the cow is the well-doc¬ umented inhibitory effect of the corpus luteum of pregnancy or the ipsilateral uterine horn (563). In nonsuckling dairy cows, nonatretic follicles begin to form an antral cavity at 0.16 mm in diameter (564). On day 15 postpartum, 77% of the largest healthy antral follicles present are 0.16 to 1.6 mm, and by day 35, only 1.5% of the smallest class are represented. Thus, during the early postpartum period, there is no constant replenishment of small antral follicles by preantral stages. On day 15 the percentage of follicles from 0.29 to 1.6 mm increased, presumably at the expense of the smallest antral follicles. At all times the ovary containing the corpus luteum had more nonatretic follicles than the

428

/ Chapter

11

contralateral ovary. In another study, follicles >3 mm were removed from the ovaries of suckling and nonsuckling beef cows on day 5 postpartum, and the number of healthy and atretic follicles was estimated based on estrogen content in follicular fluid and histology (563). Follicles from the nonsuckled cows were already considerably larger: 50% were >6 mm compared with 33% for suckled cows. Moreover, individual follicles from suckled cows had half the concen¬ tration of estrogen compared to follicles from the nonsuckled group. The “carry-over” effect of the corpus luteum was also evident in that the percentage of follicles containing estrogen in the ovary with the corpus luteum was about a third the value of the contralateral ovary (563). Another study evaluated follicular parameters in beef cows that were suckled or weaned on day 21 postpartum with the endpoint on day 25 (565). For combined follicles from 1 to >6 mm in the weaned group, there was a 68% increase in LH re¬ ceptors but no difference in FSH binding sites. The only hormone that differed in follicular fluid concentration was PRL, which was 53% higher in follicles from the weaned cows. Subsequently, the same authors (566) measured changes at 24-hr intervals after weaning on day 21 postpartum. Uti¬ lizing the largest follicle in the ovary, an increase in FSH receptors was observed 48 hr after weaning, whereas the increase in hCG receptors did not occur until 96 hr. In the suckling cow, serum levels of FSH and LH are depressed compared to weaned animals (567), and this is associated with an altered pattern of gonadotropin release. In cows nursing two young ad libitum, the LH pulse rate is about one every 6 hr, contrasted to three per 6 hr in weaned animals (562). After weaning on day 21, the number of LH pulses is approximately threefold greater in the weaned animals (565). In milked cows, FSH is secreted in the early postpartum period in discrete pulses comparable in fre¬ quency and magnitude to the profiles during pregnancy; there is a gradual increase in pulsatile release of LH in the first 1 to 2 weeks postpartum (see ref. 568). Again, LH levels are more affected than FSH levels by lactation in the cow. When pituitary explants of weaned and suckled cows are exposed to GNRH, LH secretion is doubled in the former group (566). Following weaning, over the next 4 days, basal levels of serum LH increase in linear fashion, whereas FSH is unaffected (566). It has frequently been noted that the first cycle after parturition in the cow is significantly shorter than subsequent ones judged both by serum progesterone levels and behavioral estrus. This inadequate luteal phase is similar to a disorder of the human female, and in both cases inadequate gonadotropin priming of follicles may be responsible. On day 7 postpartum, suckled beef cows have lower serum concentrations and pulse frequencies of LH and FSH than nonsuckled animals (567). The FSH was measured by a homologous radioimmunoassay, but in an¬ other study in which a heterologous FSH assay was used, serum levels of the hormone were within the normal estrous range (569). In summary, the suckling stimulus in the bovine alters

the frequency of LH (and FSH?) pulses; consequently, fol¬ licular development is impaired. As one might anticipate, pulsatile injection of GNRH has been used to reverse this situation and to induce ovulation within 4 days of treatment of suckling cows (570). Lactational anestrus in the sow usually lasts 6 weeks or longer and is, as usual, associated with altered follicular development. During the first 4 weeks of lactation, only 47% of the follicles >1 mm in diameter are normal, and the largest healthy follicles are 4 mm. On the day after weaning, there are numerous follicles 10 mm and 15 follicles 5 to 10 mm in diameter (572). Follicular development in the sow is therefore held in abeyance until the suckling stimulus is ^removed. Before weaning, serum FSH values are similar to those observed during the estrous cycle, whereas LH values are depressed (573). Serum PRL is elevated compared to cyclic values. Pigs ovariectomized 2 to 4 days after farrow¬ ing show a prompt increase in FSH throughout a 30-day period of lactation, while serum LH does not differ between ovariectomized and intact animals over this period (574). The divergent profiles evidently result from two different control mechanisms, with FSH being normally restrained in the intact sow by inhibin produced by the numerous 1to 5-mm follicles present throughout the postpartum period. After weaning, significant increases in serum FSH and LH presumably are responsible for the rapid maturation of fol¬ licles (572). Weaning to estrous intervals range from 3 to 10 days, and PRL levels decline to basal concentration 1 to 2 hr after weaning (575). Both LH and FSH concentrations rose in the 12-hr postweaning period and an increase in pulse frequency per 12 hr was especially pronounced for LH. As a corollary to these results, pulsatile administration of GNRH to lactating sows 25 days postpartum is as effec¬ tive as weaning in leading to fertile estrus within 4 days (576). Although there is no morphological evidence on follicular growth during lactation in the primate, indirect monitoring of gonadotropins and steroid hormones indicates that suck¬ ling inhibits follicular development. For example, in lac¬ tating rhesus monkeys, weaning normally occurs 9 to 12 months postpartum. Under these circumstances, serum LH is low during the first 9 months and values typical of the follicular phase of the cycle are not attained until about 1 year (577). In the absence of a suckling stimulus, basal LH levels begin to rise after the first month. Serum FSH is also reduced during the first 6 months postpartum in lactating rhesus, and normal follicular levels are not attained until 10 months. LH surges cannot be consistently elicited by in¬ jection of estradiol benzoate until 10 months in lactating monkeys, whereas in nonsuckling females, they can be elic¬ ited within the first month postpartum. This included a group of cycling monkeys who served as foster mothers after being

Follicular Selection / primed twice daily with thyrotropin-releasing hormone to become hyperprolactinemic. They responded to estradiol benzoate exactly like the normal postpartum suckling mon¬ keys, ruling out pregnancy per se as the factor responsible for ovarian refractoriness in lactating animals. In suckling or nonsuckling rhesus monkeys ovariectomized on day 24 or 25 postpartum, serum LH and FSH are suppressed by lactation (578). (Compare with results in the sow.) It is also noteworthy that pituitaries of lactating fe¬ males contain only about 8% as much LH compared to a pool of pituitaries from cycling rhesus; FSH concentrations were similar between the two groups. Basal serum PRL gradually falls in lactating rhesus but still runs at about 150 ng/ml from 60 to 180 days, whereas it declines drastically within 48 hr in nonsuckled monkeys (577). The diurnal rhythm of PRL secretion, which is absent during pregnancy, is resumed within the first week post¬ partum (579). These altered patterns of gonadotropin se¬ cretion associated with lactation are also reflected in low serum levels of estradiol and progesterone (577,579). In the former study, serum progesterone averaged 0.4 ± 0.1 ng/ml for up to 80 days postpartum, confirmatory evidence that suckling can partially “rejuvenate” the corpus luteum of pregnancy and restore some modicum of secretory activity, albeit at a low level. This is consistent with recent obser¬ vations that bromocriptine administered to suckling mon¬ keys curtails serum PRL and progesterone (580). In women, lactation delays the onset of first ovulation but the duration and patterns of suckling stimulus are vari¬ ables that confound the results (reviewed in refs. 581,582). No morphological studies are available, so that the extent to which follicular maturation is impaired is unknown. By 3 weeks after delivery, FSH levels are within the range of normal follicular values (583,584) and LH is reported to be within the normal range by 4 to 5 weeks (583). However, the pulsatile secretion of LH appears to be too low to induce estradiol secretion (584). Urinary estrogen is low in breast¬ feeding women until 40 weeks after delivery (582). The first ovulation in nonlactating women occurred between 43 and 87 days after delivery and was associated with increases in urinary pregnanediol or plasma progesterone (584). In the lactating group, there was no evidence of ovarian cyclic¬ ity until after weaning or after at least 150 days of lactation. After injection of 1 mg estradiol benzoate, a positive-feed¬ back increase in LH, was still absent in 7 lactating women at 100 days postpartum. For the first 2 weeks after delivery, the response of non-nursing mothers, to GNRH is negative; this is followed by gradually increasing responsiveness, with FSH responding before LH (585). Similar results have been reported for lactating and nonlactating women, both show¬ ing similar normal profiles of FSH and LH in response to GNRH by 4 to 5 weeks postpartum (583). Prolactin levels decline rapidly after delivery but are maintained by suckling in lactating women for long periods (see ref. 586). The question then is, With fairly early re¬ sumption of basal levels of FSH and LH and response to

429

GNRH, why is follicular maturation delayed so long in lactating women? More data is needed before it can be resolved whether PRL, directly or indirectly, plays a role or whether changes in GNRH pulsatile release is the con¬ trolling factor in lactational amenorrhea (see ref. 587).

SUMMARY AND CONCLUSIONS 1. Knowledge of the nongrowing pool of follicles—the primordial stages—is still woefully lacking. The fact that primary follicles, the first step in follicular differentiation, are always present in rather constant numbers, and regard¬ less of reproductive status, suggests that primordial follicles are recruited continuously on a daily basis. It is possible that their mobilization is random. On the other hand, prox¬ imity to nerve endings, blood vessels, larger follicles, or corpora lutea might conceivably influence their exit from the resting pool. There are even a few hints in the literature (e.g., ref. 61) that primordial follicles in the rat can be affected by gonadotropins. Whether primordial follicles en¬ ter the growing pool by chance or design, their time of transformation into primary follicles may be the decisive factor in deciding their ultimate fate. Gougeon has referred to “privileged follicles,” which emerge at a time when crit¬ ical changes in gonadotropin levels, e.g., the periovulatory period, may endow them with selective advantages for fur¬ ther normal differentiation. Gougeon believes that in the human, privileged status is bestowed on preantral follicles with 3 to 5 x 103 granulosa cells (class 1 follicles), whose development is initiated in the periovulatory window; these follicles develop a theca under the influence of high mid¬ cycle levels of LH. In the hamster, and other species as well, privileged follicles may involve still earlier stages in folliculogenesis: Primary follicles present on the morning of proestrus show dramatic increases in in vitro incorpo¬ ration of [3H]thymidine in response to the periovulatory increases in gonadotropins. It is therefore possible that ham¬ ster follicles developing in synchrony with surges of FSH and LH recurring every 4 days constitute the group selected to ovulate after 4 or 5 cycles of development. 2. It is often stated that primary and secondary follicles, i.e., preantral stages, are largely independent of gonado¬ tropic support. Although species differences may well exist, a strong case can be made that laboratory rodents are de¬ pendent on gonadotropins—especially FSH—at early stages of folliculogenesis. The literature on this subject is difficult to interpret because so much is based on subjective evalu¬ ations. Some investigators have been more impressed by the qualitative ability of some preantral follicles to develop following hypophysectomy than the fact that the number is drastically reduced from intact animals. Careful quantitative studies are needed for a number of species focusing on follicular development (healthy and atretic) in the posthypophysectomy period and the recrudescence in response to FSH preparations. In vitro steroidogenesis by preantral fol-

430

/

Chapter

11

licles from hypophysectomized versus that of intact animals would be another useful endpoint. However, there may very well be species differences in how far follicular development proceeds in the absence of gonadotropins. 3. An interesting finding of the past decade is the obser¬ vation that regional differences exist in the granulosal com¬ partment of antral follicles. This is reflected in distinct lo¬ calization of peptide receptors, enzymes, steroidogenic potential, and presumably metabolic processes as well be¬ tween the membrana granulosa and the cumulus oophorus. These structural and functional distinctions originate from the intimate relationship between the oocyte and its sur¬ rounding investment of granulosa cells. One wonders how early in the history of the preantral follicle can the future differences between cumulus and membrana granulosa be discerned. The different roles of the membrana granulosa and the cumulus-oocyte complex in the postovulatory period are obviously related to their different topographic and func¬ tional organizations in the maturing follicle. 4. Several lines of evidence show that in the rat and hamster the second periovulatory surge of FSH is respon¬ sible for the final recruitment of the large growing follicles that will ovulate 4 to 5 days later. This does not rule out the necessity of low tonic levels of LH since its neutrali¬ zation in the hamster by anti-LH precipitates atresia in the face of normal serum levels of FSH. The role of the second FSH increase during the first few postovulatory days is less clear in species with long cycles. Is it also essential for the selection of the follicles that ultimately ovulate? Early in the cycle, follicles in the ewe and cow are 0.4 and 0.5 mm, respectively, and kinetic studies indicate that follicles in this size range grow in the course of the current cycle to become the ovulatory follicles. Is it possible that further differen¬ tiation of these follicles is influenced by the postestrous FSH peak? This is a difficult question to resolve experimentally; for example, in sheep, suppression of the second FSH peak by inhibin is usually followed by a rebound in FSH levels. 5. A wide variety of stimuli induce superovulation, in¬ cluding pituitary and placental gonadotropins, antisera to some steroids, antisera to inhibin, and clomiphene—an es¬ trogen antagonist. It appears that highly pure FSH-like com¬ pounds (with high FSH.LH ratios) stimulate development of supernumerary follicles better than gonadotropins con¬ taining low FSH.LH ratios. However, in the hamster and guinea pig, continuous LH stimulation can induce sponta¬ neous superovulation. Superovulation may result from a combination of recruitment of new follicles into the popu¬ lation and prevention of atresia of larger follicles; this may depend on the stage of the cycle that gonadotropic stimulus is provided. The mechanism of superovulation may also involve an increase in ovarian blood flow, and thus the delivery rate of essential nutrients for the sudden increase in follicular growth. The temporal changes in ovarian blood flow after a superovulatory stimulus are largely unknown. It is tempting to speculate that increases in ovarian blood flow parallel (or precede) increases in the number of folli¬

cles. Superovulation also involves stimulation of ovarian FSH and LH receptors. FSH’s ability to increase granulosa cell division, aromatase activity, and LH receptor formation and LH’s ability to stimulate thecal androgen production are instrumental in causing superovulation. Excessive LH stimulation (in the case of gonadotropins with low FSH:LH ratios) may in some species down-regulate LH receptors, reduce androgen secretion, and induce atresia of developing follicles. A better understanding of the mechanism(s) of superovulation ultimately will provide insight into the nature of superovulatory compounds, the appropriate timing of the stimuli, and the factors reducing the variability of the num¬ ber of ova shed and increasing the quality of the ova. 6. Whether species have short or long cycles, the demise of luteal function accelerates follicular development and estrogen secretion. Thus, the main variable in determining the duration of the estrous cycle in the rat is whether the corpora lutea secrete progesterone for 2 or 3 days. The precision of the 4-day cycle of the hamster correlates with the rigid life span of the corpora lutea. The final impetus for follicular selection in the human is geared to luteolysis, which triggers a small, short-lived spurt of FSH. In the sheep and cow, the follicular phase is especially clearly demarcated by the spontaneous or induced regression of the corpus luteum. The luteal-follicular shift, therefore, is of paramount importance in the final selection of the ovulatory follicle(s) and is associated with progesterone withdrawal. In most cases, after luteolysis, baseline levels of FSH are unaffected, but LH increases as a consequence of an in¬ creased pulsatile release pattern. Although, I (G. S. Greenwald) have previously speculated that progesterone may directly affect follicular responsiveness, the overwhelming mass of evidence points to a classical inhibitory action on the hypothalamic-pituitary axis and, with the withdrawal of progesterone, increased secretion of LH. Presumably, the presence of higher titers of LH leads to increased se¬ cretion of androgens by the theca and concomitantly in¬ creases levels of estrogen and the development of a positivefeedback circuit in the now dominant follicles. According to this concept, although FSH is essential for the recruitment of the developing follicles (the cohort leading to the selection of the dominant follicles), the final maturation becomes more and more dependent on LH. This is also clearly shown in the pregnant rat, in which the maturation of the follicles destined to ovulate at postpartum estrus is brought about by rising titers of LH, beginning on day 20. 7. The estrous cycles of hamster, rat, and mouse usually recur at 4- to 5-day intervals, as compared with 18 to 22 days for sheep, pigs, and cows. One would intuitively as¬ sume that folliculogenesis proceeds at different rates de¬ pending on cycle length and indeed this is the case. For the laboratory rodents, it is estimated that about 3 weeks are required between the time a follicle enters the growing pool and ovulates. In contrast, similar estimates for the ewe are 6 months, and for the human, 85 days. Consequently, per¬ turbations are quickly sensed and promptly responded to by

Follicular Selection / rodent follicular populations, as exemplified by their rapid mobilization after unilateral ovariectomy or administration of PMS—and their possible rapid onset of regression after hypophysectomy. 8. There is no evidence to support the concept of “waves” of follicular activity during cycles of long duration or preg¬ nancy. In both situations, the population of primary follicles is continually replenished from the resting pool, and all stages of follicular growth are represented—from primary to tertiary. For example, in sheep and cow, estrogen-secreting antral follicles are present during the luteal phase, but their full development is held in check by progesterone, and the follicles wax and wane as their physiological life span is exceeded. A critical unanswered question is whether the follicles ultimately selected to ovulate at the end of preg¬ nancy (rat) or the cycle (ewe) are ever exposed to elevated levels of FSH or whether they can mature after exposure to tonic levels of FSH but with increased levels of LH super¬ imposed as the controlling factor. As previously mentioned, the concept of follicles emerging from the resting pool when FSH levels are high (in the immediate postovulatory period) may be the signal. For example, it takes about 21 days in the cyclic rat for a primary follicle to grow into a large preovulatory follicle. Is it merely coincidental that post¬ partum ovulation routinely occurs on day 23. Obviously, this line of reasoning does not apply to species with long gestation lengths. 9. Recent years have witnessed renewed interest in fol¬ licular degeneration based on models to induce atresia or investigations of the spontaneous event. Species seem to be divided into two categories in terms of the hormonal with¬ drawal pattern by antral follicles: in rodents, the loss of thecal androgen seems to precede the loss of granulosal aromatase activity, whereas in the ewe and human the re¬ verse sequence exists. In both groups, the loss of estrogen secretion by the atretic follicle is the common denominator. The deficit in estrogen secretion in a young antral follicle presumably curtails mitoses in the granulosa cells, and the follicle therefore lags further and further behind. The ques¬ tion remains, however, whether the loss of estrogen (or androgen) production is a primary or secondary event in atresia of tertiary follicles. It is virtually impossible to an¬ swer this question by examining follicles undergoing spon¬ taneous atresia because of the difficulty in pinpointing the moment when degeneration begins and determining the cri¬ teria for this early step. The histologic presence of a few pyknotic nuclei may be normal and of no consequence to a granulosa population of several thousand to million cells (depending on the species). By the time the follicle can definitely be classified as atretic, it is too late to establish causal relationships. Hence, it is our prejudice that exper¬ imental models in which atresia occurs as a timed, pre¬ dictable event will be the only way to determine the prime mover. Although emphasis on atresia has focused on tertiary follicles, certainly the process is even more widespread in younger follicles, and entirely different mechanisms may

431

be involved in their regression. A few studies have looked at FSH and LH receptors at the onset of atresia and found no differences in total numbers from normal follicles. A more meaningful endpoint may be the number of occupied receptors that are coupled to adenylate cyclase. We believe that a change distal to the peptide receptor may be the earliest signal of atresia. The most dramatic histological signs of atresia occur in the granulosa cells, and consequently this compartment has received the most attention. The theca is the most resistant portion of the atretic follicle, and morphological alterations may not occur until several days after granulosa cells are affected. It is possible though—at least for rodents—that the theca may be the key to antral follicular development and atresia. Hisaw originally postulated that an “undevel¬ oped” theca might be responsible for atresia. Although the morphological development of the theca in rodents may be normal, we suggest that biochemical “immaturity” may lead to atresia by impaired production of androgens, which in turn deprives the granulosa cells of substrate for conversion to estrogens. We base this belief on the pattern of steroido¬ genesis in induced and spontaneous atresia and the ability of exogenous LH to salvage the developing follicles that normally become atretic between days 3 and 4 of the hamster estrous cycle. This does not exclude an additional role for LH in recruiting more than the normal number of 20 de¬ veloping follicles in the hamster. The critical factor in the final maturation of developing follicles may be the LH re¬ ceptor level in theca interna cells. This may determine the ability of the cells to produce androstenedione and the vas¬ cularity of the theca. These two properties of the theca are so inextricably bound that it may be impossible to dissociate them. At present, however, it appears that the inception of atresia is not associated with decreases in vascularity, which may be a secondary event comparable to the sequence in luteolysis. Although the theca may be the pivotal tissue in rodent atresia, in other species the receptor deficiency may reside in the granulosa compartment involving LH and/or FSH receptors. Reasons for this possibility have been cited. 10. In this account, the follicle has largely been consid¬ ered as an isolated component, uninfluenced by other ovar¬ ian compartments. The interrelationships between follicles and corpora lutea, follicles and interstitium have been barely touched. Moreover, do tertiary follicles affect preantral stages if they share a common vascular supply? Coculture of these various tissues might be a rewarding approach. 11. Follicular development during lactation is impaired because of altered pulsatile release of LH and elevated levels of PRL. The problem of hormonal control during lactation is also complicated by species differences. What remains to be established are the relative roles of GNRH and PRL and how much of their effects are exerted indirectly or directly at the ovarian level. 12. Fifteen years ago, one could confidently predict that in vitro studies of antral follicles, granulosa, and thecal cells and peptide receptor distribution and follicular kinetics would

432

/

Chapter 11

be profitable areas of investigation. It is also equally obvious that molecular biology—an area untouched in this review— will have significant impact on the problems of folliculogenesis, as will the in vitro effects of growth factors inter¬ acting with peptide and steroid hormones. It is worth re¬ iterating, however, that more classical techniques still have a great deal to offer in this field and should not be neglected or discarded.

ACKNOWLEDGMENTS The original work of the authors included in this chapter was supported by NIH Grants HD00596-25 (G.S.G.) and HDK00478 and HD20389 (P.F.T.). We thank Mrs. Darlene Limback for help in assembling the bibliography and Mrs. Linda Carr for typing the manuscript.

REFERENCES 1. Greenwald, G. S. (1972): Editorial. Of eggs and follicles. Am. J. Anat., 135:1-4. 2. Hisaw, F. L. (1947): Development of the graafian follicle and ovu¬ lation. Physiol. Rev., 27:95-119. 3. Brambell, F. W. R. (1956): Ovarian changes. In: Marshall’s Phys¬ iology of Reproduction edited by A. S. Parkes, pp. 397-542. Long¬ mans, Green, New York. 4. Falck, B. (1959): Site of production of oestrogen in rat ovary as studied in micro-transplants. Acta Physiol. Scand. Suppl. 163, 47:5101. 5. Greenwald, G. S. (1974): Role of follicle-stimulating hormone and luteinizing hormone in follicular development and ovulation. In: Handbook of Physiology!Endocrinology, Vol. 4: Part 2, edited by E. Knobil and W. H. Sawyer, pp. 293-323. American Physiological Society, Washington, D.C. 6. Channing, C. P., and Tsafriri, A. (1977): Mechanism of action of luteinizing hormone and follicle-stimulating hormone on the ovary in vitro. Metabolism, 26:413-468. 7. Lindner, H. R., Amsterdam, A., Salomon, Y., et al. (1977): Intraovarian factors in ovulation: determinants of follicular response to gonadotrophins. J. Reprod. Fertil., 51:215-235. 8. Armstrong, D. T., and Dorrington, J. H. (1977): Estrogen biosyn¬ thesis in the ovaries and testes. In: Regulatory Mechanisms Affecting Gonadal Hormone Action, edited byJ. A. Thomas andR. H. Singhal, pp. 215-258. University Park Press, Baltimore. 9. Richards, J. S. (1980): Maturation of ovarian follicles: actions and interactions of pituitary and ovarian hormones on follicular cell dif¬ ferentiation. Physiol. Rev., 60:51-89. 10. diZerega, G. S., and Hodgen, G. D. (1981): Folliculogenesis in the primate ovarian cycle. Endocrine Rev., 2:27-49. 11. Erickson, G. F. (1983): Primary cultures of ovarian cells in serumfree medium as models of hormone-dependent differentiation. Mol. Cell. Endocrinol., 29:21-49. 12. Hsueh, A. J. W., Adashi, E. Y., Jones, P. B. C., and Welsh, T. H., Jr. (1984): Hormonal regulation of the differentiation of cul¬ tured ovarian granulosa cells. Endocrine Rev., 5:76-127. 13. Tsafriri, A., and Braw, R. H. (1984): Experimental approaches to atresia in mammals. Oxford Rev. Reprod. Biol., 6:226-265. 14. Hillier, S. G. (1985): Sex steroid metabolism and follicular devel¬ opment in the ovary. Oxford Rev. Reprod. Biol., 7:168-222. 15. Zuckerman, S. S., and Weir, B. J., eds. (1977): The Ovary, 2nd ed. Academic, New York. 16. Jones, R. E., ed. (1978): The Vertebrate Ovary. Comparative Biology and Evolution. Plenum, New York. 17. Crighton, D. B., Foxcroft, G. R., Haynes, N. B., and Lamming, G. E. (1978): Control of Ovulation. Butterworths, Boston. 18. Midgley, A. R., Jr., and Sadler, W. A., eds. (1979): Ovarian Fol¬ licular Development and Function. Raven Press, New York.

19. Channing, C. P., Marsh, J. M., and Sadler, W. A., eds. (1979): Ovarian Follicular and Corpus Luteum Function. Plenum, New York. 20. Edwards, R. G. (1980): Conception in the Human Female. Aca¬ demic, Ne\y York. 21. Motta, P. M., and Hafez, E. S. E., eds. (1980): Biology of the Ovary. Martinus Nijhoff Publishers, The Hague. 22. Schwartz, N. B., and Hunzicker-Dunn, M., eds. (1981): Dynamics of Ovarian Function. Raven Press, New York. 23. Channing, C. P., and Segal, S. J., eds. (1982): Intraovarian Control Mechanisms. Plenum, New York. 24. Greenwald, G. S., and Terranova, P. F., eds. (1983): Factors Reg¬ ulating Ovarian Function. Raven Press, New York. 25. Austin, C. R., and Short, R. V., eds. (1982-84): Reproduction in Mammals, 2nd ed. Cambridge University Press, New York. 26. Lamming, G. E.,ed. (1984): Marshall’sPhysiology of Reproduction. Vol. 1, 4th Ed., Reproductive Cycles of Vertebrates. Churchill Liv¬ ingstone, New York. 27. Guraya, S. S. (1985): Biology of Ovarian Follicles in Mammals. Springer-Verlag, New York. 28. Toft, D. O., and Ryan, R. J., eds. (1985): Proceedings of the Fifth Ovarian Workshop. Ovarian Workshops, Champaign, IL. 29. Mauleon, P. (1967): Cinetique de l’ovogenese chez les mammiferes. Arch. Anat. Microsc. Morphol. Exp., 56:125-150. 30. Gerard, P., and Herlant, M. (1953): Sur la persistance de phenomenes d’oogenese chez les lemuriens adultes. Arch. Biol., 64:97-111. 31. Peters, H., Byskov, A. G., and Faber, M. (1973): Intraovarian reg¬ ulation of follicle growth in the immature mouse. In: The Devel¬ opment and Maturation of the Ovary and its Functions, edited by H. Peters, pp. 20-23. Excerpta Medica, Amsterdam. 32. Lunenfeld, B., Kraiem, Z., and Eshkol, A. (1975): The function of the growing follicle. J. Reprod. Fertil., 45:567-574. 33. Peters, H. (1969): The development of the mouse ovary from birth to maturity. Acta Endocrinol., 62:98-116. 34. de Wolff-Exalto, E. A. (1982): Influence of gonadotrophins on early follicle cell development and early oocyte growth in the immature rat. J. Reprod. Fertil., 66:537-542. 35. Hage, A. J., Groen-Klevant, A. C., and Welschen, R. (1978): Fol¬ licle growth in the immature rat ovary. Acta Endocrinol., 88 375— 382. 36. Uilenbroek, J. T. J., de Wolff-Exalto, E. A., and Blankenstein, M. A. (1976): Serum gonadotrophins and follicular development in immature rats after early androgen administration. J. Endocrinol 68' 461-468. 37. Vomachka, A. J., and Greenwald, G. S. (1979): The development of gonadotropin and steroid hormone patterns in male and female hamsters from birth to puberty. Endocrinology, 105:960-966. 38. Krarup, T., Pedersen, T., and Faber, M. (1969): Regulation of oocyte growth in the mouse ovary. Nature, 224:187-188. 39. Gosden, R. G., Laing, S. C., Felicio, L. S., Nelson, J. F., and Finch, C. E. (1983): Imminent oocyte exhaustion and reduced fol¬ licular recruitment mark the transition to acyclicity in aging C57BL/6J mice. Biol. Reprod., 28:255-260. 40. Mandl, A. M., and Shelton, M. (1959): A quantitative study of oocytes in young and old nulliparous laboratory rats. J. Endocrinol 18:444-450. 41. Block, E. (1952): Quantitative morphological investigations of the follicular system in women. Variations at different ages Acta Anat 14:108-123. 42. Gougeon, A. (1981): Cinetique de la croissance et de l’involution des follicules ovariens pendant le cycle menstruel chez la femme. Thesis, Universite Pierre and Marie Curie. 43. Koering, M. J. (1983): Preantral follicle development during the menstrual cycle in the Macaca mulatto ovary. Am. j Anat 166 429433. 44. Mauleon, P., and Rao, K. H. (1963): Variations genetiques des populations foliculaires dans les ovaires de rates impuberes. Ann. Biol. Anim. Bioch. Biophys., 3:21-31. 45. Mauleon, P., and Pelletier, J. (1964): Variations genetiques du fonctionnement hypophysaire de trois souches de rattes immatures, relafions avec la fertilite. Ann. Biol. Anim. Bioch. Biophys., 4:10546. Merchant, H. (1975): Rat gonadal and ovarian organogenesis with and without germ cells. An ultrastructural study. Dev. Biol., 44:1-

Follicular Selection / 47. Merchant-Larios, H., and Centeno, B. (1981): Morphogenesis of the ovary from the sterile W/Wv mouse. In: Eleventh International Con¬ gress of Anatomy: Advances in the Morphology of Cells and Tissues, pp. 383-392. Liss, New York. 48. Pedersen, T. (1970): Follicle kinetics in the ovary of the cyclic mouse. Acta Endocrinol., 64:304-323. 49. Butcher, R. L., and Kirkpatrick-Keller, D. (1984): Patterns of fol¬ licular growth during the four-day estrous cycle of the rat. Biol. Reprod., 31:280-286. 50. Pedersen, T., and Peters, H. (1971): Follicle growth and cell dy¬ namics in the mouse ovary during pregnancy. Fertil. Steril., 22:4252. 51. Groen-Klevant, A. C. (1981): An autoradiographic study of follicle growth in the ovaries of cyclic rats. Acta Endocrinol.. 96:377-381. 52. Greenwald, G. S. (1974): Quantitative aspects of follicular devel¬ opment in the untreated and PMS-treated cyclic hamster. Anat. Rec., 178:139-143. 53. Chiras, D. D., and Greenwald, G. S. (1980): Analysis of ovarian follicular development and thymidine incorporation in the cyclic golden hamster. Am. J. Anat., 157:309-317. 54. Roy, S. K.. and Greenwald, G. S. (1986): Quantitative analysis of in vitro incorporation of [JH] thymidine intotiamster follicles during the oestrous cycle. J. Reprod. Fertil., 77:143-152. 55. Zarrow, M. X., and Wilson, E. D. (1961): The influence of age on superovulation in the immature rat and mouse. Endocrinology, 69:851— 855. 56. Welschen, R., and Rutte, M. (1971): Ovulation in adult rats after treatment with pregnant mare serum gonadotrophin during oestrus. Acta Endocrinol., 68:41-49. 57. Greenwald, G. S. (1962): Analysis of superovulation in the adult hamster. Endocrinology, 71:378-389. 58. Chiras, D. D., and Greenwald, G. S. (1978): Ovarian follicular development in cyclic hamsters treated with a superovulatory dose of pregnant mare’s serum. Biol. Reprod., 19:895-901. 59. Lane, C. E., and Greep, R. O. (1935): The follicular apparatus of the ovary of the hypophysectomized immature rat and the ef¬ fects of hypophyseal gonadotropic hormones on it. Anat. Rec., 63:139-146. 60. Paesi, F. J. A. (1949): The influence of hypophysectomy and of subsequent treatment with chorionic gonadotrophin on follicles of different size in the ovary of the rat. Acta Endocrinol., 3:89-104. 61. De Reviers, M. M. (1974): Etude quantitative de Paction des hor¬ mones gonadotropes hypophysaires sur la population folliculaire de l’ovaire de ratte immature—signification biologique du dosage de Phormone folliculo-stimulante par le test de Steelman et Pohley. Thesis, L’Universite de Tours. 62. Edwards, R. G., Fowler, R. E., Gore-Langton, R. E., Gosden, R. G., Jones, E. C., Readhead, C., and Steptoe, P. C. (1977): Nor¬ mal and abnormal follicular growth in mouse, rat and human ovaries. J. Reprod. Fertil., 51:237-263. 63. Kim, I., Shaha, C., and Greenwald, G. S. (1984): A species dif¬ ference between hamster and rat in the effect of oestrogens on growth of large preantral follicles. J. Reprod. Fertil., 72:179-185. 64. Nakano, R., Mizuno, T., Katayama, K., and Tojo, S. (1975): Growth of ovarian follicles in rats in the absence of gonadotrophins. J. Re¬ prod. Fertil., 45:545-546. 65. Hirshfield, A. N. (1985): Comparison of granulosa cell proliferation in small follicles of hypophysectomized, prepubertal, and mature rats. Biol. Reprod., 32:979-987. 66. Faddy, M. J. E., Jones, E. C., and Edwards, R. G. (1976): An analytical model for ovarian follicle dynamics. J. Exp. Zool., 197:173185. 67. Oakberg, E. F. (1979): Follicular growth and atresia in the mouse. In Vitro, 15:41-49. 68. Moore, P. J., and Greenwald, G. S. (1974): Effect of hypophysec¬ tomy and gonadotropin treatment on follicular development and ovu¬ lation in the hamster. Am J. Anat., 139:37-48. 69. Garza, F., Shaban, M. A., and Terranova, P. F. (1984): Luteinizing hormone increases the number of ova shed in the cyclic hamster and guinea-pig. J. Endocrinol., 101:289-298. 70. Kim, I., and Greenwald, G. S. (1986): Occupied and unoccupied FSH receptors in follicles of cyclic, hypophysectomized or hypophysectomized/gonadotropin-treated hamsters. Mol. Cell. Endocrinol., 44:141-145.

433

71. Dempsey, E. W. (1937): Follicular growth rate and ovulation after various experimental procedures in the guinea pig. Am. J. Physiol., 120:126-132. 72. Perry, J. S., and Rowlands, I. W. (1963): Hypophysectomy of the immature guinea-pig and the ovarian response to gonadotrophins. J. Reprod. Fertil., 6:393-404. 73. Dufour, J., Cahill, L. P. and Mauleon, P. (1979): Short- and long-term effects of hypophysectomy and unilateral ovariectomy on ovarian follicular populations in sheep. /. Reprod. Fertil., 57:301-309. 74. Goldenberg, R. L., Powell, R. D., Rosen, S. W., Marshall, J. R., and Ross, G. T. (1976): Ovarian morphology in women with anosmia and hypogonadotropic hypogonadism. Am. J. Obstet. Gynecol., 126:91-94. 75. Lim, H. T., Meinders, A. E., deHaan, L. D., and Bronkhorst, F. B. (1984): Anovulation presumably due to the gonadotrophinresistant ovary syndrome. Eur. J. Obstet. Gynecol. Reprod. Biol., 16:327-337. 76. Talbert, L. M., Raj, M. H. G., Hammond, M. G., and Greer, T. (1984): Endocrine and immunologic studies in a patient with resistant ovary syndrome. Fert. Steril., 42:741-744. 77. Lintem-Moore, S., and Moore, G. P. M. (1977): Comparative as¬ pects of oocyte growth in mammals. In: Reproduction and Evolution. Proceedings of the Fourth Symposium on Comparative Biology of Reproduction, Canberra, December 1976, edited by J. H. Calaby and C. H. Tyndale-Biscoe, pp. 215-219. Australian Academy of Science, Canberra City. 78. Lintem-Moore, S., and Moore, G. P. M. (1979): The initiation of follicle and oocyte growth in the mouse ovary. Biol. Reprod., 20:773778. 79. Moore, G. P. M., Lintem-Moore, S., Peters, H., and Faber, M. (1974): RNA synthesis in the mouse oocyte. J. Cell Biol., 60:416422. 80. Canipari, R., Pietrolucci, A., and Mangia, F. (1979): Increase of total protein synthesis during mouse oocyte growth. J. Reprod. Fer¬ til., 57:405-413. 81. Muller, U., and Urban, E. (1981): An oocyte-specific antigen and its possible role in the organization of the ovarian follicle of the rat. Differentiation, 20:274-277. 82. Takaoka, H., Satoh, H., Makinoda, S., Moriya, S., and Ichinoe, K. (1985): Granulosa-cell growth factor in oocyte and its transport systems. Acta Obstet. Gynaecol. Jpn., 37:92-98. 83. Hirshfield, A. N. (1986): Patterns of [3H] thymidine incorporation differ in immature rats and mature, cycling rats. Biol. Reprod., 34:229-235. 84. Cran, D. G., Moor, R. M., and Hay, M. F. (1980): Fine structure of the sheep oocyte during antral follicle development. J. Reprod. Fertil., 59:125-132. 85. Tesoriero, J. V. (1984): Comparative cytochemistry of the devel¬ oping ovarian follicles of the dog, rabbit, and mouse: origin of the zona pellucida. Gamete Res., 10:301-318. 86. Shimizu, S., Tsuji, M., and Dean, J. (1983): In vitro biosynthesis of three sulfated glycoproteins of murine zonae pellucidae by oocytes grown in follicle culture. J. Biol. Chem., 258:5858-5863. 87. Colonna, R., and Mangia, R. (1983): Mechanisms of amino acid uptake in cumulus-enclosed mouse oocytes. Biol. Reprod., 28:797803. 88. Wassarman, P. M., Bleil, J. D., Cascio, S. M., LaMarca, M. J., Letoumeau, G. E., Mrozak, S. C., and Schultz, R. M. (1981): Programming of gene expression during mammalian oogenesis. In: Bioregulators of Reproduction, edited by G. Jagiello and H. J. Vogel, pp. 119-150. Academic, New York. 89. Dunbar, B. S. (1983): Morphological, biochemical, and immuno¬ chemical characterization of the mammalian zona pellucida. In: Mechanism and Control of Animal Fertilization, pp. 139-175. Ac¬ ademic, New York. 90. Wolgemuth, D. J., Celenza, J., Bundman, D. S., and Dun¬ bar, B. S. (1984): Formation of the rabbit zona pellucida and its relatioship to ovarian follicular development. Dev. Biol., 106: 1-14. 91. Skinner, S. M., Mills, T., Kirchick, H. J., and Dunbar, B. S. (1984): Immunization with zona pellucida proteins results in abnormal ovar¬ ian follicular differentiation and inhibition of gonadotropin-induced steroid secretion. Endocrinology, 115:2418-2432.

434

/ Chapter 11

92. Michael, S. D. (1983): Interactions of the thymus and the ovary. In: Factors Regulating Ovarian Function, edited by G. S. Greenwald and P. F. Terranova, pp. 445—464. Raven, New York. 93. Lane, C. E., and Davis, F. R. (1939): The ovary of the adult rat. I. Changes in growth of the follicle and in volume and mitotic activity of the granulosa and theca during the estrous cycle. Anat. Rec., 73:429-442. 94. Ax, R. L., and Ryan, R. J. (1979): The porcine ovarian follicle. IV. Mucopolysaccharides at different stages of development. Biol. Reprod., 20:1123-1132. 95. Yanagishita, M., Rodbard, D., and Hascall, V. C. (1979): Isolation and characterization of proteoglycans from porcine ovarian follicular fluid. J. Biol. Chem., 254:911-920. 96. Schweitzer, M., Jackson, J. C., and Ryan, R. J. (1981): The porcine ovarian follicle. VII. FSH stimulation of in vitro [3H]-glucosamine incorporation into mucopolysaccharides. Biol. Reprod., 24:332-340. 97. Burghardt, R. C., and Anderson, E. (1981): Hormonal modulation of gap junctions in rat ovarian follicles. Cell Tissue Res., 214:181— 193. 98. Larsen, W. J., Tung, H. N., and Polking, C. (1981): Response of granulosa cell gap junctions to human chorionic gonadotropin (hCG) at ovulation. Biol. Reprod., 25:1119-1134. 99. Fletcher, W. H., and Greenan, J. R. T. (1985): Receptor mediated action without receptor occupancy. Endocrinology, 116:1660-1662. 100. Hay, M. F., and Moor, R. M. (1975): Distribution of A5-3(3-hydroxysteroid dehydrogenase activity in the graafian follicle of the sheep. J. Reprod. Fertil., 43:313-322. 101. Cran, D. G., Hay, M. F., and Moor, R. M. (1979): The fine structure of the cumulus oophorus during follicular development in sheep. Cell Tissue Res., 202:439-451. 102. Mestwerdt, W., and Muller, O. (1978): Elektronenoptischmorphometrische Untersuchungen zum Luteinisierungsprozess der Follikelgranulosazelle menschlicher Ovarien. Arch. Gynaekol., 225: 51-65. 103. Bomsel-Helmreich, O., Gougeon, A., Thebault, A., Saltarelli, D., Milgrom, E., Frydman, R., and Papiemik, E. (1979): Healthy and atretic human follicles in the preovulatory phase: differences in ev¬ olution of follicular morphology and steroid content of follicular fluid. J. Clin. Endocrinol. Metabol., 48:686-694. 104. Amin, H., Richart, R. M., and Brinson, A. O. (1976): Preovulatory granulosa cells and steroidogenesis. An ultrastructural study in the rhesus monkey. Obstet. Gynecol., 47:562-568. 105. Zoller, L. C. (1984): A comparison of rat and hamster preovulatory follicles: an examination of differences in morphology and enzyme activity using qualitative and quantitative analyses. Anat. Rec., 210:279-291. 106. Bjersing, L. (1978): Maturation, morphology, and endocrine function of the follicular wall in mammals. In: The Vertebrate Ovary. Com¬ parative Biology and Evolution, edited by R. E. Jones, pp. 181— 214. Plenum, New York. 107. Fortune, J. E. (1986): Bovine theca and granulosa cells interact to promote androgen production. Biol. Reprod., 35:292-299. 108. Makris, A., Olsen, D., and Ryan, K. J. (1983): Significance of the A5 and A4 steroidogenic pathways in the hamster preovulatory fol¬ licle. Steroids, 42:641-651. 109. Weisz, J., and Zoller, L. C. (1979): Quantitative cytochemistry in the study of regional specialization in the membrana granulosa of the ovulable type of follicle. In: Quantitative Cytochemistry and Its Applications, edited by J. R. Pattison, L. Bitensky, and J. Chayen, pp. 269-283. Academic, New York. 110. Zoller, L. C., and Weisz, J. (1979): A quantitative cytochemical study of glucose-6-phosphate dehydrogenase and A5-3p-hydroxysteroid dehydrogenase activity in the membrana granulosa of the ovulable type of follicle of the rat. Histochemistry, 62:125-135. 111. Zoller, L. C., and Enelow, R. (1983): A quantitative histochemical study of lactate dehydrogenase and succinate dehydrogenase activ¬ ities in the membrana granulosa of the ovulatory follicle of the rat. Histochem. J., 15:1055-1064. 112. Hillensjo, T., Magnusson, C., Svensson, U., and Thelander, H. (1981): Effect of luteinizing hormone and follicle-stimulating hor¬ mone on progesterone synthesis by cultured rat cumulus cells. En¬ docrinology, 108:1920-1924. 113. Erickson, G. F., Hofeditz, C., Unger, M., Allen, W. R., and Dulbecco, R. (1985): A monoclonal antibody to a mammary cell line

recognizes two distinct subtypes of ovarian granulosa cells. Endo¬ crinology, 117:1490-1499. 114. Staigmiller, R. B., and Moor, R. M. (1984): Effect of follicle cells on the maturation and developmental competence of ovine oocytes matured outside the follicle. Gamete Res., 9:221-229. 115. Charlton, H. M., Parry, D., Halpin, D. M. G., and Webb, R. (1982): Distribution of 125I-labelled follicle-stimulating hormone and human chorionic gonadotrophin in the gonads of hypogonadal (hpg) mice. J. Endocrinol., 93:247-252. 116. Armstrong, D. T., Weiss, T. J., Selstam, G., and Seamark, R. F. (1981): Hormonal and cellular interactions in follicular steroid bio¬ synthesis by the sheep ovary. J. Reprod. Fertil. Suppl., 30:143-154. 117. Channing, C. P., Schaerf, F. W., Anderson, L. D., and Tsafriri, A. (1980): Ovarian follicular and luteal physiology. In: Reproductive Physiology. HI. International Review of Physiology, edited by R. O. Greep, pp. 117-201. University Park Press, Baltimore. 118. Nakano, R., Sasaki, K., Shima, K., and Kitayama, S. (1983): Fol¬ licle-stimulating hormone and luteinizing hormone receptors on por¬ cine granulosa cells during follicular maturation: an autoradiographic study. Exp. Clin. Endocrinol., 81:17-23. 119. Uilenbroek J. T. J., and Richards, J. S. (1979): Ovarian follicular development during the rat estrous cycle: gonadotropin receptors and » follicular responsiveness. Biol. Reprod., 20:1159-1165. 120. Amsterdam, A., Koch, Y., Lieberman, M. E., and Lindner, H. R. (1975): Distribution of binding sites for human chorionic gonado¬ tropin in the preovulatory follicle of the rat. J. Cell. Biol., 67:894900. 121. Lawrence, T. S., Dekel, N., and Beers, W. H. (1980): Binding of human chorionic gonadotropin by rat cumuli oophori and granulosa cells: a comparative study. Endocrinology, 106:1114-1118. 122. Bortolussi, M., Marini, G., and Reolon, M. L. (1979): A histo¬ chemical study of the binding of l25I-HCG to the rat ovary throughout the estrous cycle. Cell Tissue Res., 197:213-226. 123. Oxberry, B. A., and Greenwald, G. S. (1982): An autoradiographic study of the binding of l25I-labeled follicle-stimulating hormone, human chorionic gonadotropin and prolactin to the hamster ovary throughout the estrous cycle. Biol. Reprod., 27:505-516. 124. Roy, S. K., and Greenwald, G. S. (1985): Evidence for binding sites for FSH and hCG in mammalian oocytes. In: Proceedings of the Fifth Ovarian Workshop, edited by D. O. Toft and R. J. Ryan, pp. 143-147. Ovarian Workshops, Champaign, IL. 125. Niimura, S., and Ishida, K. (1983): Histochemical demonstration of hydroxysteroid dehydrogenases in the oocytes in antral follicles of pigs, cattle and horses. Jpn. J. Anim. Reprod., 29:150-153. 126. Hiura, M., and Fujita, H. (1977): Electron microscopy of the cytodifferentiation of the theca cell in the mouse ovary. Arch Histol Jpn., 40:95-105. 127. O’Shea, J. D. (1970): An ultrastructural study of smooth muscle¬ like cells in the theca externa of ovarian follicles in the rat. Anat Rec., 167:127-140. 128. Amsterdam, A., Lindner, H. R., and Stewart, U. G. (1977): Lo¬ calization of actin and myosin in the rat oocyte and follicular wall by immunofluorescence. Anat. Rec., 187:311-328. 129. Capps, M. L., Lawrence, I. E., Jr., and Burden, H. W. (1981): Cellular junctions in perifollicular contractile tissue of the rat ovary during the preovulatory period. Cell Tissue Res., 219:133-141. 130. Walles, B., Edvinsson, L., Owman, C., Sjoberg, N. O., and Sporrong, B. (1976): Cholinergic nerves and receptors mediating con¬ traction of the graafian follicle. Biol. Reprod., 15:565-572. 131. Kobayashi, Y., Sjoberg, N. O., Walles, B., Owman, C., Wright, K. H., Santulli, R., and Wallach, E. E. (1983): The effect of ad¬ renergic agents on the ovulatory process in the in vitro perfused rabbit ovary. Am. J. Obstet. Gynecol., 145:857-864. 132. Wylie, S. N., Roche, P. J., and Gibson, W. R. (1985): Ovulation after sympathetic denervation of the rat ovary produced by freezing its nerve supply. J. Reprod. Fertil., 75:369-373. 133. Stoklosowa, S., Gregoraszczuk, E., and Channing, C. P. (1982): Estrogen and progesterone secretion by isolated cultured porcine thecal and granulosa cells. Biol. Reprod., 26:943-952. 134. Merk, F. B., Albright, J. T., and Botticelli, C. R. (1973): The fine structure of granulosa cell nexuses in rat ovarian follicles. Anat Rec 175:107-126. 135. Kranzfelder, D., Korr, H., Mestwerdt, W., and Maurer-Schultze, B. (1984): Follicle growth in the ovary of the rabbit after ovulation-

Follicular Selection /

136.

137.

138.

139.

140.

141.

142.

143.

144.

145. 146.

147.

148.

149.

150.

151.

152.

153.

154.

155. 156.

157.

inducing application of human chorionic gonadotropin. Cell. Tissue Res., 238:611-620. Makris, A., Klagsbrun, M. A., Yasumizu, T., and Ryan, K. J. (1983): An endogenous ovarian growth factor which stimulates BALB/3T3 and granulosa cell proliferation. Biol. Reprod., 29:11351141. O’Shea, J. D., Cran, D. G., Hay, M. F., and Moor, R. M. (1978): Ultrastructure of the theca interna of ovarian follicles in sheep. Cell Tissue Res., 187:457^172. Priedkalns, J., Weber, A. F., and Zemjanis, R. (1968): Qualitative and quantitative morphological studies of the cells of the membrana granulosa, theca interna and corpus luteum of the bovine ovary. Z. Zellforsch., 85:501-520. Wordinger, R. J., Rudick, V. L., and Rudick, M. J. (1983): Immunohistochemical localization of laminin within the mouse ovary. J. Exp. Zoo!., 228:141-143. Roy, S. K., and Greenwald, G. S. (1985): An enzymatic method for dissociation of intact follicles from the hamster ovary: histological and quantitative aspects. Biol. Reprod., 32:203-215. Stoklosowa, S., Bahr, J., and Gregoraszczuk, E. (1978): Some mor¬ phological and functional characteristics of cells of the porcine theca interna in tissue culture. Biol. Reprod., 19:712-719. Tsang, B. K., Ainsworth, L., Downey, B. R., and Marcus, G. J. (1985): Differential production of steroids by dispersed granulosa and theca interna cells from developing preovulatory follicles of pigs. J. Reprod. Fertil., 74:459-471. Katayama, E. (1984): Monolayer culture of human ovarian thecal cells—A study on morphological and functional characteristics. Acta Obstet. Gynaecol. Jpn., 36:927-936. Bogovich, K., and Richards, J. S. (1984): Androgen synthesis during follicular development: evidence that rat granulosa cell 17-ketosteroid reductase is independent of hormonal regulation. Biol. Reprod., 31:122— 131. Koninckx, P. R. (1981): New aspects of ovarian function in man and in rat. Thesis, Katholieke Universiteit Leuven, Leuven. diZerega, G. S., and Hodgen, G. D. (1980): Fluorescence locali¬ zation of luteinizing hormone/human chorionic gonadotropin uptake in the primate ovary. II. Changing distribution during selection of the dominant follicle. J. Clin. Endocrinol. Metab., 51:903-907. Bassett, D. L. (1943): The changes in the vascular pattern of the ovary of the albino rat during the estrous cycle. Am. J. Anat., 73:251— 291. Zeleznik, A. J., Schuler, H. M., and Reichert, L. E., Jr. (1981): Gonadotropin-binding sites in the rhesus monkey ovary: role of the vasculature in the selective distribution of human chorionic gonad¬ otropin to the preovulatory follicle. Endocrinology, 109:356-362. McNatty, K. P., Heath, D. A., Lun, S., Fannin, J. M., McDiarmid, J. M., and Henderson, K. M. (1984): Steroidogenesis by bovine theca interna in an in vitro perifusion system. Biol. Reprod., 30:159170. Koos, R. D., and LeMaire, W. J. (1983): Factors that may regulate the growth and regression of blood vessels in the ovary. Semin. Reprod. Endocrinol., 1:295-307. Frederick, J. L., Shimanuki, T., and diZerega, G. S. (1984): Ini¬ tiation of angiogenesis by human follicular fluid. Science, 224:389390. Frederick, J. L., Nuguyen, H., Preston, D. S., Frederick, J. J., Campeau, J. D., Ono, T., and diZerega, G. S. (1985): Initiation of angiogenesis by porcine follicular fluid. Am. J. Obstet. Gynecol., 152:1073-1078. Gospodarowicz, D., Cheng, J., Lui, G. M., Baird, A., Esch, F., and Bohlen, P. (1985): Corpus luteum angiogenic factor is related to fibroblast growth factor. Endocrinology, 117:2283-2391. Makris, A., Ryan, K. J., Takehiko, Y., Hill, C. L., and Zetter, B. R. (1984): The nonluteal porcine ovary as a source of angio¬ genic activity. Endocrinology, 15:1672-1677. Saidapur, S. K. (1978): Follicular atresia in the ovaries of nonmam¬ malian vertebrates. Int. Rev. Cytol., 54:225-244. Byskov, A. G. (1978): Follicular atresia. In: The Vertebrate Ovary. Comparative Biology and Evolution, edited by R. E. Jones, pp. 533— 562. Plenum, New York. Ingram, D. L. (1962): Atresia. In: The Ovary, edited by S. S. Zuckerman, A. M. Mandl, and P. Eckstein, pp. 247-273. Academic, New York.

435

158. Spanel-Borowski, K. (1981): Morphological investigations on fol¬ licular atresia in canine ovaries. Cell Tisue Res., 214:155-168. 159. Byskov, A. G. S. (1974): Cell kinetic studies of follicular atresia in the mouse ovary. J. Reprod. Fertil., 37:277-285. 160. Brailly, S., Gougeon, A., Milgrom, E., Bomsel-Helmreich, O., and Papiemik, E. (1981): Androgens and progestins in the human ovarian follicle: differences in the evolution of preovulatory, healthy non¬ ovulatory, and atretic follicles. J. Clin. Endocrinol. Metab., 53: 128-133. 161. Westergaard, L., McNatty, K. P., Christensen, I., Larsen, J. K., and Byskov, A. G. (1982): Flow cytometric deoxyribonucleic acid analysis of granulosa cells aspirated from human ovarian follicles. A new method to distinguish healthy and atretic ovarian follicles. J. Clin. Endocrinol. Metab., 55:693-698. 162. Koering, M. J., Goodman, A. L., Williams, R. F., and Hodgen, G. D. (1982): Granulosa cell pyknosis in the dominant follicle of monkeys. Fert. Steril., 37:837-844. 163. Odeblad, E. (1952): Contributions to the theory and technique of quantitative autoradiography with 32P with special reference to the granulosa tissue of the graafian follicles in the rabbit. Acta Radiol. (Stockh.) Suppl., 93:1-123. 164. Deane, H. W. (1952): Histochemical observations on the ovary and oviduct of the albino rat during the estrous cycle. Am. J. Anat., 91:363-414. 165. Bukovsky, A., Presl, J., and Zidovsky, J. (1979): Migration of lymphoid cells into the granulosa of rat ovarian follicles. IRCS Med. Sci., 7:603-604. 166. Bukovsky, A., Presl, J., and Holub, M. (1984): The ovarian follicle as a model for the cell-mediated control of tissue growth. Cell Tissue Res., 236:717-724. 167. Peluso, J. J., and England-Charlesworth, C. (1982): Development of preovulatory follicles and oocytes during the oestrous cycle of mature and aged rats. Acta Endocrinol., 100:434-443. 168. Peluso, J. J., England-Charlesworth, C., Bolender, D. L., and Steger, R. W. (1980): Ultrastructural alterations associated with the initiation of follicular atresia. Cell Tissue Res., 211:105-115. 169. Gondos, B. (1982): Ultrastructure of follicular atresia in the rat. Gamete Res., 5:199-206. 170. Ryan, R. J., and Lee, C. Y. (1976): The role of membrane bound receptors. Biol. Reprod., 14:16-29. 171. Peluso, J. J., Steger, R. W., and Hafez, E. S. E. (1977): Surface ultrastructural changes in granulosa cells of atretic follicles. Biol. Reprod., 16:600-604. 172. Hay, M. F., Cran, D. G., and Moor, R. M. (1976): Structural changes occurring during atresia in sheep ovarian follicles. Cell Tis¬ sue Res., 169:515-529. 173. Hay, M. F., Moor, R. M., Cran, D. G., and Dott, H. M. (1979): Regeneration of atretic sheep ovarian follicles in vitro. J. Reprod. Fertil., 55:195-207. 174. Moor, R. M., Hay, M. F., Dot, H. M., and Cran, D. G. (1978): Macroscopic identification and steroidogenic function of atretic fol¬ licles in sheep. J. Endocrinol., 77:309-318. 175. Meinecke, B., Meinecke-Tillmann, S., and Gips, H. (1982): Experimentelle Untersuchungen zur Steroidsekretion intakter und atretischer Follikel in vitro. Berl. Munch. Tierarztl. Wochenschr., 95:107111. 176. Henderson, K. M., Kieboom, L. E., McNatty, K. P., Lun, S., and Heath, D. A. (1984): [l25I]hCG binding to bovine thecal tissue from healthy and atretic antral follicles. Mol. Cell. Endocrinol., 34:91— 98. 177. Zachariae, F. (1957): Studies in the mechanism of ovulation. Au¬ toradiographic investigations on the uptake of radioactive sulphate (35S) into the ovarian follicular mucopolysaccharides. Acta Endo¬ crinol., 26:215-224. 178. Beilin, M. E., and Ax, R. L. (1984): Chondroitin sulfate: an indicator of atresia in bovine follicles. Endocrinology, 114:428^434. 179. Guraya, S. S. (1973): Follicular atresia. Proc. Indian Natl. Sci. Acad., 39:311-332. 180. Lobel, B. L., Rosenbaum, R. M., and Deane, H. W. (1961): Enzymic correlates of physiological regression of follicles and corpora lutea in ovaries of normal rats. Endocrinology, 68:232247. 181. Ryan, R. J. (1981): Follicular atresia: some speculations on bio¬ chemical markers and mechanisms. In: Dynamics of Ovarian Func-

436

/ Chapter 11

tion, edited by N. B. Schwartz and M. Hunzicker-Dunn. Raven Press, New York. 182. Breitenecker, G., Friedrich, F., and Kemeter, P. (1978): Further investigations on the maturation and degeneration of human ovarian follicles and their oocytes. Fertil. Steril., 29:336-341. 183. McNatty, K. P., Makris, A., De Grazia, C., Osathanondh, R., and Ryan, K. J. (1979): Steroidogenesis in granulosa cells and corpus luteum. The production of progesterone, androgens and oestrogens by human granulosa cells in vitro and in vivo. J. Steroid Biochem , 11:775-779. 184. Austin, C. R. (1961): The Mammalian Egg. Blackwell, Oxford, UK. 185. Byskov, A. G. (1979): Atresia, In: Ovarian Follicular Development and Function, edited by A. R. Midgley and W. A. Sadler, pp. 4157. Raven Press, New York. 186. Dawson, A. B. (1952): Argyrophilic inclusions in the cytoplasm of the ova of the rat in normal and atretic follicles. Anat. Rec , 112 37— 59. 187. Peluso, J. J., Bolender, D. L., and Perri, A. (1979): Temporal changes associated with the degeneration of the rat oocyte Biol Reprod., 20:423-430. 188. Westergaard, L. (1985): Follicular atresia in relation to oocyte mor¬ phology in non-pregnant and pregant women. J. Reprod Fertil 74:113-118. 189. Vasques-Nin, G. H., and Sotelo, J. R. (1967): Electron microscope study of the atretic oocytes of the rat. Z. Zelforsch. Abt. Histochem 80:518-533. 190. Baker, T. G., and Franchi, L. L. (1967): The fine structure of oogonia and oocytes in human ovaries. J. Cell. Sci., 2:213-234. 191. Centola, G. M. (1982): Light microscopic observations of alterations in staining of the zona pellucida of porcine follicular oocytes: possible early indications of atresia. Gamete Res., 6:293-304. 192. Donahue, R. P., and Stem, S. (1968): Follicular cell support of oocyte maturation: production of pyruvate in vitro. J. Reprod Fertil 17:395-398. 193. Albertini, D. F., and Anderson, E. (1974): The appearance and structure of intercellular connections during the ontogeny of the rabbit ovarian follicle with particular reference to gap junctions J Cell Biol., 63:234-250. 194. Osman, P. (1985): Rate and course of atresia during follicular development in the adult cyclic rat. J. Reprod. Fertil., 73 261270. 195. Moor, R. M., and Trounson, A. O. (1977): Hormonal and follicular factors affecting maturation of sheep oocytes in vitro and their sub¬ sequent developmental capacity. J. Reprod. Fertil., 49:101-109. 196. Guraya, S. S., and Greenwald, G. S. (1964): A comparative histochemical study of interstitial tissue and follicular atresia in the mammalian ovary. Anat. Rec., 149:411-434. 197. Mossman, H. W., Koering, M. J., and Ferry, D., Jr. (1964): Cyclic changes of interstitial gland tissue of the human ovary. Am J Anat 115:235-256. 198. Schwall, R., and Erickson, G. F. (1981): Functional and morpho¬ logical changes in rat theca cells during atresia. In: Dynamics of Ovarian Function, edited by N. B. Schwartz and M. HunzickerDunn, pp. 29-34. Raven Press, New York. 199. Terranova, P. F., Martin, N. C., and Chien, S. (1982): Theca is the source of progesterone in experimentally induced atretic follicles of the hamster. Biol. Reprod., 26:721-727. 200. Silavin, S. L., and Greenwald, G. S. (1984): Steroid production by isolated theca and granulosa cells after initiation of atresia in the hamster. J. Reprod. Fertil., 71:387-392. 201. O Shea, J. D., Hay, M. F., and Cran, D. G. (1978): Ultrastructural changes in the theca interna during follicular atresia in sheep. J. Reprod. Fertil., 54:183-187. 202. Bruce, N. W., and Moor, R. M. (1976): Capillary blood flow to ovarian follicles, stroma and corpora lutea of anaesthetized sheep. J. Reprod. Fertil., 46:299-304. 203. Findlay, J. K., and Carson, R. S. (1980): Selective binding of go¬ nadotrophins and the control of follicular growth and atresia. In: Advances in Physiological Science, Vol. 15: Reproduction and De¬ velopment, edited by B. Flerko, G. Setalo, and L. Tima, pp. 7989. Pergamon, Budapest. 204. Motta, P. M., and Familiari, G. (1981): Occurrence of contractile tissue in the theca externa of atretic follicles in the mouse ovary. Acta Anat., 109:103-114.

205. Mori, T., Fujita, Y., Nihnobu, K., Ezaki, Y., Kubo, K., and Nishimura, T. (1982): Steroidogenesis in vitro by human ovarian follicles during the process of atresia. Clin. Endocrinol., 16:391-400. 206. McNatty, K. P., Smith, D. M., Makris, A., Osathanondh, R., and Ryan, K. J. (1979): The microenvironment of the human antral follicle: interrelationships among the steroid levels in antral fluid, the population of granulosa cells, and the status of the oocyte in vivo and in vitro. J. Clin. Endocrinol. Metab., 49:851-860. 207. McNatty, K. P., Makris, A., Osathanondh, R., and Ryan, K. J. (1980): Effects of luteinizing hormone on steroidogenesis by thecal tissue from human ovarian follicles in vitro. Steroids, 36:53-63. 208. Maxson, W. S., Haney, A. F., and Schomberg, D. W. (1985): Steroidogenesis in porcine atretic follicles: loss of aromatase activity in isolated granulosa and theca. Biol. Reprod., 33:495-501. 209. Carson, R. S., Findlay, J. K., Clarke, I. J., and Burger, H. G. (1981): Estradiol, testosterone, and androstenedione in ovine follic¬ ular fluid during growth and atresia of ovarian follicles. Biol. Re¬ prod., 24:105-113. 210. Tsonis, C. G., Carson, R. S., and Findlay, J. K. (1984): Relation¬ ships between aromatase activity, follicular fluid oestradiol-17[3 and testosterone concentrations, and diameter and atresia of individual ovine follicles. J. Reprod. Fertil., 72:153-163. 211. Carson, R. S., Findlay, J. K., Burger, H. G., and Trounson, A. O. (1979): Gonadotropin receptors of the ovine ovarian follicle during follicular growth and atresia. Biol. Reprod., 21:75-87. 212. Ireland, J. J., and Roche, J. F. (1983): Development of nonovulatory antral follicles in heifers: changes in steroids in follicular fluid and receptors for gonadotropins. Endocrinology, 112:150-156. 213. McNatty, K. P., Lun, S., Heath, D. A., Kieboom, L. E., and Henderson, K. M. (1985): Influence of follicular atresia on LHinduced cAMP and steroid synthesis by bovine thecae interna. Mol. Cell Endocrinol., 39:209-215. 214. Braw, R. H., Bar-Ami, S., and Tsafriri, A. (1981): Effect of hypophysectomy on atresia of rat preovulatory follicles. Biol Reprod 25:989-996. " 215. Bill, C. H., II, and Greenwald, G. S. (1981): Acute gonadotropin deprivation. I. A model for the study of follicular atresia Biol Reprod., 24:913-921. 216. Braw, R. H., and Tsafriri, A. (1980): Follicles explanted from pen¬ tobarbitone-treated rats provide a model for atresia. J. Reprod Fertil 59:259-265. 217. Udenbroek, J. T. J., Woutersen, P. J. A., and van der Schoot, P. (1980). Atresia of preovulatory follicles: gonadotropin binding and steroidogenic activity. Biol. Reprod., 23:219-229. 218. Terranova, P. F. (1980): Effects of phenobarbital-induced ovulatory delay on the follicular population and serum levels of steroids and gonadotropins in the hamster: a model for atresia. Biol Reprod 23:92-99. ” 219. Peters, H., Byskov, A. G., Himelstein-Braw, R., and Faber, M. (1975): Follicular growth: the basic event in the mouse and human ovary. J. Reprod. Fertil., 45:559-566. 220. Peluso, J. J., and Steger, R. W. (1978): Role of FSH in regulating granulosa cell division and follicular atresia in rats. J. Reprod Fertil 54:275-278. 221. Braw, R. H., and Tsafriri, A. (1980): Effect of PMSG on follicular atresia in the immature rat ovary. J. Reprod. Fertil., 59:267-272. 222. Matson, P. L., Gledhill, B., and Collins, W. P. (1984): Effect of LH on steroidogenesis by hamster follicles isolated at defined stages of development. J. Reprod. Fertil., 70:675-681. 223. Bagnell, C. A., Mills, T. M., Costoff, A., andMahesh, V. B. (1982): A model for the study of androgen effects on follicular atresia and ovulation. Biol. Reprod., 27:903-914. 224. Magnusson, C., Bar Ami, S., Braw, R., and Tsafriri, A. (1983): Oxygen consumption by rat oocytes and cumulus cells during induced atresia. J. Reprod. Fertil., 68:97-103. 225. Hubbard, C. J., and Greenwald, G. S. (1985): Morphological changes in atretic graafian follicles during induced atresia in the hamster Anat. Rec., 212:353-357. 226. Hubbard, C. J.. and Greenwald, G. S. (1981): Changes in DNA, cyclic nucleotides and steroids during induced follicular atresia in the hamster. J. Reprod. Fertil., 63:455^161. 227. Hubbard, C. J., and Greenwald, G. S. (1983): In vitro effects of luteinizing hormone on induced atretic graafian follicles in the ham¬ ster. Biol. Reprod., 28:849-859.

Follicular Selection /

228. Shaha, C., and Greenwald, G. S. (1982): Autoradiographic analy¬ sis of changes in ovarian binding of FSH and hCG during in¬ duced follicular atresia in the hamster. J. Reprod. Fertil., 66:197-

250.

201. 229. Terranova, P. F. (1981): Steroidogenesis in experimentally induced atretic follicles of the hamster: a shift from estradiol to progesterone synthesis. Endocrinology, 108:1885-1890. 230. Na, J. Y., Garza, F., and Terranova, P. F. (1985): Alterations in follicular fluid steroids and follicular hCG and FSH binding during atresia in hamster. Proc. Soc. Exp. Biol. Med., 179:123-127. 231. Mizuno, O., Otani, T., Shirota, M., and Sasamoto, S. (1983): Mat¬ uration of ovarian follicles after inhibition of ovulation in rats. J. Endocrinol., 97:113-119. 232. Uilenbroek, J. T. J., van der Linden, R., and Woutersen, P. J. A. (1984): Changes in oestrogen biosynthesis in preovulatory rat follicles after blockage of ovulation with pentobarbitone sodium. J. Reprod. Fertil., 70:549-555. 233. van der Schoot, P., den Besten, D., and Uilenbroek, J. T. J. (1982): Atresia of preovulatory follicles in rats treated with sodium pento¬ barbital: effects of bromocriptine. Biol. Reprod., 27:189-199. 234. Freeman, M. E., Butcher, R. L., and Fugo, N. W. (1970): Alteration of oocytes and follicles by delayed ovulation. Biol. Reprod., 2:209215. 235. Peluso, J. J., Steger, R. W., and Hafez, E. S. E. (1977): Sequential changes associated with the degeneration of preovulatory rat follicles. J. Reprod. Fertil., 49:215-218. 236. Louvel, J. P., Harman, S. M., Schreiber, J. R., and Ross, G. T. (1975): Evidence for a role of androgens in follicular maturation. Endocrinology, 97:366-372. 237. Hillier, S. G., and Ross, G. T. (1979): Effects of exogenous tes¬ tosterone on ovarian weight, follicular morphology and intraovarian progesterone concentration in estrogen-primed hypophysectomized immature female rats. Biol. Reprod., 20:261-268. 238. Harmon, S. M., Louvet, J. P.. and Ross, G. T. (1975): Interaction of estrogen and gonadotrophins on follicular atresia. Endocrinology, 96:1145-1152. 239. Peluso, J. J., Charlesworth, J., and England-Charlesworth, C. (1981): Role of estrogen and androgen in maintaining the preovulatory fol¬ licle. Cell Tisue Res., 216:615-624. 240. Schreiber, J. R., Reid, R., and Ross, G. T. (1976): A receptor-like testosterone-binding protein in ovaries from estrogen-stimulated hy¬ pophysectomized immature female rats. Endocrinology, 98:12061213. 241. Schreiber. J. R., and Ross, G. T. (1976): Further characterization of a rat ovarian testosterone receptor with evidence for nuclear trans¬ location. Endocrinology, 99:590-596. 242. Saiduddin, S., and Zassenhaus, H. P. (1978): Effect of testosterone and progesterone on the estradiol receptor in the immature rat ovary. Endocrinology, 102:1069-1076. 243. Kohut, J. K., Jarrell, J. F., and Younglai, E. V. (1985): Does dihydrotestosterone induce atresia in the hypophysectomized im¬ mature female rat treated with pregnant mare’s serum gonadotropin? Am. J. Obstet. Gynecol., 151:250-255. 244. Farookhi, R. (1980): Effects of androgen on induction of gonado¬ tropin receptors and gonadotropin-stimulated adenosine 3'-5'-mono¬ phosphate production in rat ovarian granulosa cells. Endocrinology, 106:1216-1223. 245. Farookhi, R. (1981): Atresia: A hypothesis. In: Dynamics of Ovarian Function, edited by N. B. Schwartz and M. Hunzicker-Dunn, pp. 13-23. Raven Press, New York. 246. Tonetta, S. A., Spicer, L. J., and Ireland, J. J. (1985): CI628 inhibits follicle-stimulating hormone (FSH)-induced increases in FSH recep¬ tors of the rat ovary: requirement of estradiol for FSH action. En¬ docrinology, 116:715-722. 247. Hillier, S. G., van den Boogaard, A. M. J., Reichert, L. E., Jr., and van Hall, E. V. (1980): Intraovarian sex steroid hormone inter¬ actions and the regulation of follicular maturation: aromatization of androgens by human granulosa cells in vitro. J. Clin. Endocrinol. Metab., 50:640-647. 248. Greenwald, G. S., and Limback, D. L. (1984): Effects of treatment with cycloheximide at proestrus on subsequent in vitro follicular steroidogenesis in the hamster. Biol. Reprod., 30:1105-1116. 249. Baird, D. T., Swanston, I. A., and McNeilly, A. S. (1981): Rela¬ tionship between LH, FSH, and prolactin concentration and the se¬

251.

252.

253.

254.

255.

256.

257.

258.

259.

260. 261.

262.

263.

264.

265.

266.

267.

268. 269.

270.

271.

272.

437

cretion of androgens and estrogens by the preovulatory follicle in the ewe. Biol. Reprod., 24:1013-1025. Hutz, R. J., Dierschke, D. J., and Wolf, R. C. (1986): Markers of atresia in ovarian follicular components from rhesus monkeys treated with estradiol-17(3. Biol. Reprod., 34:65-70. Cole, H. H. (1975): Studies on reproduction with emphasis on go¬ nadotropins, antigonadotropins and progonadotropins. Biol. Reprod., 12:194-211. Moore, W. T., Jr., Burleigh, B. D., and Ward, D. N. (1980): Chorionic gonadotropins: comparative studies and comments on relationships to other glycoprotein hormones. In: Chorionic Go¬ nadotropin edited by S. J. Segal, pp. 89-126. Plenum, New York. Stewart, F., and Allen, W. R. (1979): The binding of FSH, LH and PMSG to equine gonadal tissues. J. Reprod. Fertil. Suppl., 27:431440. Licht, P., Gallo, A. B., Aggarwal, B. B., Farmer, S. W., Castelino, J. B., and Papkoff, H. (1979): Biological and binding activities of equine pituitary gonadotrophins and pregnant mare serum gonado¬ trophin. J. Endocrinol., 83:311-322. Murphy, B. D., Mapletoft, R. J., Manns, J., and Humphrey, W. D. (1984): Variability in gonadotrophin preparations as a factor in the superovulatory response. Theriogenology, 21:117-125. Combamous, Y., Guillou, F., Martinat, N., and Cahoreau, C. (1984): Origine de la double activite FSH + LH de la choriogonadotropine equine (eCG/PMSG). Ann. Endocrinol. (Paris), 45:261-268. Leaver, H. A., and Boyd, G. S. (1981): Action of gonadotrophic hormones on cholesterol side-chain cleavage and cholesterol ester hydrolase in the ovary of the immature rat. J. Reprod. Fertil., 63:101108. Younglai, E. V. (1984): Effects of pregnant mare’s serum gonado¬ trophin administered in vivo on steroid accumulation by isolated rabbit ovarian follicles. Acta Endocrinol., 107:531-537. Matson, P. L., Tyler, J. P. P., and Collins, W. P. (1981): Follicular steroid content and oocyte meiotic status after PMSG stimulation of immature hamsters. J. Reprod. Fertil., 61:443-452. Cran, D. G. (1983): Follicular development in the sheep after priming with PMSG. J. Reprod. Fertil., 67:415-423. Cahill, L. P., Mariana, J. C., and Mauleon, P. (1979): Total follicular populations in ewes of high and low ovulation rates. J. Reprod. Fertil., 5:27-36. Greenwald, G. S. (1961): Quantitative study of follicular develop¬ ment in the ovary of the intact or unilaterally ovariectomized hamster. J. Reprod. Fertil., 2:351-361. Monniaux, D., Mariana, J. C., and Gibson, W. R. (1984): Action of PMSG on follicular populations in the heifer. J. Reprod. Fertil., 70:243-253. Dott, H. M., Hay, M. F., Cran, D. G., and Moor, R. M. (1979): Effect of exogenous gonadotrophin (PMSG) on the antral follicle population in the sheep. J. Reprod. Fertil., 56:683-689. Moor, R. M., Kruip, T. A. M., and Green, D. (1984): Intraovarian control of folliculogenesis: limits to superovulation? Theriogenology, 21:103-116. Greenwald, G. S., and Terranova, P. F. (1983): Development in the cyclic hamster of refractoriness to the superovulatory action of antiLH serum. J. Reprod. Fertil., 69:297-301. Dhondt, D., Bouters, R., Spincemaille, J., Coryn, M., and Vandeplassche, M. (1978): The control of superovulation in the bovine with a PMSG-antiserum. Theriogenology, 9:529-534. Greenwald, G. S. (1979): Analysis of superovulation in the hamster: 1962-1978. Ann. Biol. Anim. Bioch. Biophys., 19:1483-1487. Greenwald, G. S. (1973): Effect of an anti-PMS serum on ovulation and estrogen secretion in the PMS-treated hamster. Biol. Reprod., 9:437-446. Hirshfield, A. N. (1984): Stathmokinetic analysis of granulosa cell proliferation in antral follicles of cyclic rats. Biol. Reprod., 31:52— 58. Krishna, A., Terranova, P. F., Matteri, R. L.. and Papkoff, H. (1986): Histamine and increased ovarian blood flow mediate LHinduced superovulation in the cyclic hamster. J. Reprod. Fertil., 76:23-29. Krishna, A., and Terranova, P. F. (1985): Alterations in mast cell degranulation and ovarian histamine in the proestrous hamster. Biol. Reprod., 32:1211-1217.

438

/

Chapter

11

273. Szego, C. M., and Gitin, E. S. (1964): Ovarian histamine depletion during acute hyperaemic response to luteinizing hormone. Nature, 201:682-684. 274. Lipner, H. (1971): Ovulation from histamine depleted ovaries. Proc. Soc. Exp. Biol. Med., 136:111-114. 275. Wallach, E. E., Wright. K. H., and Hamada, Y. (1978): Investi¬ gation of mammalian ovulation with an in vitro perfused rabbit ovary preparation. Am. J. Obstet. Gynecol., 132:728-738. 276. Knox, E., Lowry, S., and Beck, L. (1979): Prevention of ovulation in rabbits by antihistamine. In: Ovarian Follicular Development and Function, edited by A. R. Midgley and W. A. Sadler, pp. 159-163. Raven Press, New York. 277. Kobayashi, Y„ Wright, K. H., Santulli, R., Kitai, H., and Wallach, E. E. (1983): Effect of histamine and histamine blockers on the ovulatory process in the in vitro perfused rabbit ovary. Biol. Reprod., 28:385-392. 278. Scaramuzzi, R. J., Martensz, N. D., and Van Look, P. F. A. (1980): Ovarian morphology and the concentration of steroids, and of go¬ nadotrophins during the breeding season in ewes actively immunized against oestradiol-17(3 or oestrone. J. Reprod. Fertil., 59:303-310. 279. Scaramuzzi, R. J., Baird, D. T., Clarke, I. J., Martensz, N. D., and Van Look, P. F. A. (1980): Ovarian morphology and the con¬ centration of steroids during the oestrous cycle of sheep actively immunized against androstenedione. J. Reprod. Fertil., 58:27-35. 280. Land, R. B., Morris, B. A., Baxter, G., Fordyce, M., and Forster, J. (1982): Improvement of sheep fecundity by treatment with antisera to gonadal steroids. J. Reprod. Fertil., 66:625-634. 281. Wallace, J. M., and McNeilly, A. S. (1985): Increase in ovulation rate after treatment of ewes with bovine follicular fluid in the luteal phase of the oestrous cycle. J. Reprod. Fertil., 73:505-515. 282. Lasley, B. L. (1984): Treating infertility: CRES employs innovative method. CRES Rep., 2:No. 2. 283. Jones, G. S. (1984): Update on in vitro fertilization. Endocrine Rev 5:62-75. 284. Terranova, P. F., and Greenwald, G. S. (1979): Antiluteinizing hormone: chronic influence on steroid and gonadotropin levels and superovulation in the pregnant hamster. Endocrinology, 104 1013— 1019. 285. Greenwald, G. S., and Terranova, P. F. (1981): Induction of su¬ perovulation in the cyclic hamster by a single injection of antilu¬ teinizing hormone serum. Endocrinology, 108:1903-1908. 286. Terranova, P. F., and Greenwald, G. S. (1981): Alteration of the serum follicle-stimulating hormone to luteinizing hormone ratio in the cyclic hamster treated with antiluteinizing hormone: relationship to serum estradiol, free antiluteinizing hormone, and superovulation. Endocrinology, 108:1909-1914. 287. Opavsky, M. A., and Armstrong, D. T. (1985): The effectiveness of FSH in inducing superovulation is influenced by LH. Biol. Reprod Suppl. 1, 32:71. 288. Kim, I., and Greenwald, G. S. (1984): Hormonal requirements for maintenance of follicular and luteal function in the hypophysectomized cyclic hamster. Biol. Reprod., 30:1063-1072. 289. Terranova, P. F., and Greenwald, G. S. (1981): Increased ovulation rate in the cyclic guinea pig after a single injection of antiserum to LH. J. Reprod. Fertil., 61:37-42. 290. Fitzgerald, J. A., Ruggles, A. J., and Hansel, W. (1985): Increased ovulation rate of adult ewes treated with anti-bovine LH antiserum during the normal breeding season. J. Anim. Sci., 60:749-754. 291. Peppier, R. D. (1968): Method and mechanism of ovulatory com¬ pensation following unilateral ovariectomy in the rat. Thesis, Uni¬ versity of Kansas. 292. Jones, R. E., and Summers, C. H. (1984): Compensatory follicular hypertrophy during the ovarian cycle of the house gecko, Hemidactylus frenatus. Anat. Rec., 209:59-65. 293. Bleier, W. J., and Ehteshami, M. (1981): Ovulation following uni¬ lateral ovariectomy in the California leaf-nosed bat (Macrotus californicus). J. Reprod. Fertil., 63:181-183. 294. Hunter, J. (1787): An experiment to determine the effect of extir¬ pating one ovarium upon the number of young produced. Phil. Trans 17:233-239. 295. Arai, H. (1920): On the cause of the hypertrophy of the surviving ovary after semispaying (albino rat) and on the number of ova in it. Am. J. Anat., 28:59-79.

296. Lipschutz, A. (1928): New developments in ovarian dynamics and the law of follicular constancy. Br. J. Exp. Biol., 5:283-291. 297. Chiras, D. D., and Greenwald, G. S. (1978): Acute effects of uni¬ lateral ovariectomy on follicular development in the cylic hamster. J. Reprod. Fertil., 52:221-225. 298. Peppier, R. D., and Greenwald, G. S. (1970): Effects of unilateral ovariectomy on ovulation and cycle length in 4- and 5-day cycling rats. Am. J. Anat., 127:1-8. 299. Peppier, R. D., and Greenwald, G. S. (1970): Influence of unilateral ovariectomy on follicular development in cycling rats. Am. J Anat 127:9-14. 300. Otani, T., and Sasamoto, S. (1982): Plasma and pituitary hormone changes and follicular development after unilateral ovariectomy in cyclic rats. J. Reprod. Fertil., 65:347-353. 301. McLaren, A. (1966): Regulation of ovulation rate after removal of one ovary in mice. Proc. R. Soc. London [Biol.], 166:316-340. 302. Fleming, M. W., Rhodes, R. C., Ill, and Dailey, R. A. (1984): Compensatory responses after unilateral ovariectomy in rabbits. Biol. Reprod., 30:82-86. 303. Hermreck, A. S., and Greenwald, G. S. (1964): The effects of unilateral ovariectomy on follicular maturation in the guinea pig Anat. Rec., 148:171-176. 304. Land, R. B. (1973): Ovulation rate of Finn-Dorset sheep following unilateral ovariectomy or chlorpromazine treatment at different stages of the oestrous cycle. J. Reprod. Fertil., 33:99-105. 305. Findlay, J. K., and Cumming, I. A. (1977): The effect of unilateral ovariectomy on plasma gonadotropin levels, estrus and ovulation rate in sheep. Biol. Reprod., 17:178-183. 306. Brinkley, H. J., and Young, E. P. (1969): Effects of unilateral ovariectomy or the unilateral destruction of ovarian components on the follicles and corpora lutea of the nonpregnant pig. Endocrinology 84:1250-1256. 307. Brinkley, H. J., Wickersham, E. W., First, N. L., and Casida, L. E. (1964): Effect of unilateral ovariectomy on the structure and function of the corpora lutea of the pig. Endocrinology, 74 462467. 308. Sundaram, S. K., and Stob, M. (1967): Effect of unilateral ovar¬ iectomy on reproduction and induced ovulation in ewes J Anim Sci., 26:374-376. 309. Roy, S. K., and Greenwald, G. S. (1986): Effects of FSH and LH on incorporation of [3H] thymidine into follicular DNA J Reprod Fertil. 78:201-209. 310. Saiduddin, S., Rowe, R. F., and Casida, L. E. (1970): Ovarian follicular changes following unilateral ovariectomy in the cow. Biol Reprod., 2:408-412. 311. Johnson, S. K., Smith, M. F., and Elmore, R. G. (1985): Effect of unilateral ovariectomy and injection of bovine follicular fluid on gonadotropin secretion and compensatory ovarian hypertrophy in prepuberal heifers. J. Anim. Sci., 60:1055-1060. 312. Cochrane, R. L., and Holmes, R. L. (1966): Unilateral ovariectomy and hypophysectomy in the rhesus monkey. J. Endocrinol., 35:427313. Sopelak, V. M., and Hodgen, G. D. (1984): Contralateral tubalovanan apposition and fertility in hemiovariectomized primates Fer¬ til. Steril., 42:633-637. 314. Speert, H., Na Sh, W., and Kaplan, A. L. (1956): Tubal pregnancy. Some observations on external migration of the ovum and compen¬ satory hypertrophy of the residual ovary. Obstet. Gynecol., 7:322315. Gougeon, A., and Lefevre, B. (1983): Evolution of the diameters of the largest healthy and atretic follicles during the human menstrual cycle. J. Reprod. Fertil., 69:497-502. 316. Bast, J. D„ and Greenwald, G. S. (1977): Acute and chronic ele¬ vations in serum levels of FSH after unilateral ovariectomy in the cyclic hamster. Endocrinology, 100:955-966. 317. Welschen, R., Dullaart, J., and deJong, F. H. (1978): Interrelationships between circulating levels of estradiol-173, progesterone, FSH and LH immediately after unilateral ovariectomy in the cyclic rat Biol. Reprod., 18:421-427. ’ *•» V..U.OIVHOUH, ix. ix., roru, j. j., ana Day, B. N. (1985): Effect of follicular fluid treatment on follicle-stimulating hormone, luteinizing hormone and compensatory ovarian hypertro¬ phy in prepuberal gilts. Biol. Reprod., 32:111-119.

Follicular Selection / 319. Baird, D. T., Backstrom, T., McNeilly, A. S., Smith, S. K., and Wathen, C. G. (1984): Effect of enucleation of the corpus luteum at different stages of the luteal phase of the human menstrual cycle on subsequent follicular development. J. Reprod. Fertil 70 615624. 320. Baranczuk, R., and Greenwald, G. S. (1973): Peripheral levels of estrogen in the cycling hamster. Endocrinology, 92*805-812. 321. Curry, T. E., Jr., Lawrence, I. E., Jr., and Burden, H. W. (1984): Effect of ovarian sympathectomy on follicular development during compensatory ovarian hypertrophy in the guinea-pig. J. Reprod Fertil., 71:39-44. 322. Chatterjee, A., and Greenwald, G. S. (1972): The long-term effects of unilateral ovariectomy of the cycling hamster and rat. Biol. Re¬ prod., 7:238-246. 323. Peppier, R. D. (1971): Effects of unilateral ovariectomy on follicular development and ovulation in cycling, aged rats. Am.J.Anat., 132:423428. 324. Hirshfield, A. N. (1982): Follicular recruitment in long-term hemicastrate rats. Biol. Reprod., 27:48-53. 325. Hirshfield, A. N. (1983): Compensatory ovarian hypertrophy in the long-term hemicastrate rat: size distribution of growing and atretic follicles. Biol. Reprod., 28:271-278. 326. Butcher, R. L. (1977): Changes in gonadotropins and steroids as¬ sociated with unilateral ovariectomy of the rat. Endocrinology, 101:830— 839. 327. Baker, T. G., Challoner, S., and Burgoyne, P. S. (1980): The number of oocytes and the rate of atresia in unilaterally ovariectomized mice up to 8 months after surgery. J. Reprod. Fertil., 60:449^156. 328. Schwartz, N. B., Cobbs, S B., and Ely, C. A. (1972): What is the function(s) of the proestrous FSH surge in the rat. In: Endocrinology, Proceedings of the 4th International Congress of Endocrinology, Washington (June 1972), Excerpta Med. Int. Cong. Ser. No. 273, pp. 897-902. Excerpta Medica, Amsterdam. 329. Chiras, D. D., and Greenwald, G. S. (1977): An autoradiographic study of long-term follicular development in the cylic hamster. Anat. Rec., 188:331-337. 330. Welschen, R. (1973): Amounts of gonadotropins required for normal follicular growth in hypophysectomized adult rats. Acta Endocrinol. (Copenh.), 72:137-155. 331. Hirshfield, A. N., and Midgley, A. R., Jr. (1978): Morphometric analysis of follicular development in the rat. Biol. Reprod., 19:597— 605. 332. Ireland, J. J., and Richards, J. S. (1978): A previously undescribed role for luteinizing hormone (LH:hCG) on follicular cell differentia¬ tion. Endocrinology, 102:1458-1465. 333. Terranova, P. F., and Garza, F. (1983): Relationship between the preovulatory luteinizing hormone (LH) surge and androstenedione systhesis of preantral follicles in the cyclic hamster: detection by in vitro responses to LH. Biol. Reprod., 29:630-636. 334. Richards, J. S., and Midgley, A. R., Jr. (1976): Protein hormone action: a key to understanding follicular and luteal cell development. Biol. Reprod., 14:82-94. 335. Richards, J. S., Ireland, J. J., Rao, M. C., Bemath, G. A., Midgley, A. R., Jr., and Reichert, L. E., Jr. (1976): Ovarian follicular de¬ velopment in the rat: hormone receptor regulation by estradiol, fol¬ licle stimulating hormone and luteinizing hormone. Endocrinology, 99:1562-1570. 336. Midgley, A. R., Jr. (1973): Autoradiographic analysis of gonado¬ tropin binding to rat ovarian tissue sections. Adv. Exp. Med. Biol., 36:365-378. 337. Bast, J. D., and Greenwald, G. S. (1974): Serum profiles of folliclestimulating hormone, luteinizing hormone and prolactin during the estrous cycle of the hamster. Endocrinology, 94:1295-1299. 338. Bex, F. J., and Goldman, B. D. (1975): Serum gonadotropins and follicular development in the Syrian hamster. Endocrinology, 96:928933. 339. Siegel, H. I., Bast, J. D., and Greenwald, G. S. (1976): The effects of phenobarbital and gonadal steroids on periovulatory serum levels of luteinizing hormone and follicle stimulating hormone in the ham¬ ster. Endocrinology, 98:48-55. 340. Gay, V. L., Midgley, A. R., Jr., and Niswender, G. D. (1970): Patterns of gonadotropin secretion associated with ovulation. Fed. Proc., 29:1880-1887.

439

341. Butcher, R. L., Collins, W. E., and Fugo, N. W. (1974): Plasma concentrations of LH, FSH, prolactin, progesterone and estradiol173 throughout the 4-day estrous cycle of the rat. Endocrinology, 94:1704-1708. 342. DePaolo, L. U., Shander, D., Wise, P. M., Barraclough, C. A., and Channing, C. P. (1979): Identification of inhibin-like activity in ovarian venous plasma of rats during the estrous cycle. Endocrinol¬ ogy, 105:647-654. 343. Fujii, T., Hoover, D. J., and Channing, C. P. (1983): Changes in inhibin activity, and progesterone, oestrogen and androstenedione concentrations, in rat follicular fluid throughout the oestrous cycle. J. Reprod. Fertil., 69:307-314. 344. Sander, H. J., van Leeuwen, E. C. M., and deJong, F. H. (1984): Inhibin-like activity in media from cultured rat granulosa cells col¬ lected throughout the oestrous cycle. J. Endocrinol., 103:77-84. 345. Welschen, R., Hermans, W. P., and deJong, F. H. (1980): Possible involvement of inhibin in the interrelationship between numbers of antral follicles and peripheral FSH concentrations in female rats. J. Reprod. Fertil., 60:485-493. 346. Chappel, S. C. (1979): Cyclic fluctuations in ovarian FSH-inhibiting material in golden hamsters. Biol. Reprod., 21:447-453. 347. Erickson, G. F., and Hsueh, A. J. W. (1978): Secretion of inhibin by rat granulosa cells in vitro. Endocrinology, 103:1960-1963. 348. Lee, V. W. K. (1983): PMSG treated immature female rat—a model system for studying control of inhibin secretion. In: Factors Regu¬ lating Ovarian Function, edited by G. S. Greenwald and P. F. Ter¬ ranova, pp. 157-161. Raven Press, New York. 349. Schwartz, N. B., and Channing, C. P. (1977): Evidence for ovarian “inhibin”: suppression of the secondary rise in serum follicle stim¬ ulating hormone levels in proestrous rats by injection of porcine follicular fluid. Proc. Natl. Acad. Sci. USA. 74:5721-5724. 350. Chiras, D. D., and Greenwald, G. S. (1978): Effects of steroids and gonadotropins on follicular development in the hypophysectomized hamster. Am. J. Anat., 152:307-320. 351. Zeleznik, A. J., Midgley, A. R., Jr., and Reichert, L. E., Jr. (1974): Granulosa cell maturation in the rat: increased binding of human chorionic gonadotropin following treatment with follicle stimulating hormone in vivo. Endocrinology, 95:818-825. 352. Pencharz, R. I. (1940): Effect of estrogens and androgens alone and in combination with chorionic gonadotropin on the ovary of the hypophysectomized rat. Science, 91:554-555. 353. Williams, P. C. (1940): Effect of stilbestrol on the ovaries of hy¬ pophysectomized rats. Nature, 145:388-389. 354. Smith, B. D., and Bradbury, J. T. (1963): Ovarian response to gonadotropins after pre-treatment with diethylstilbestrol. Am. J. Physiol., 204:1023-1027. 355. Kudolo, G. B., Elder, M. G., and Myatt, L. (1984): A novel oes¬ trogen-binding species in rat granulosa cells. J. Endocrinol., 102:8391. 356. Greenwald, G. S. (1975): Proestrous hormone surges dissociated from ovulation in the estrogen treated hamster. Endocrinology, 97:878884. 357. Chappel, S. C., and Selker, F. (1979): Relation between the secretion of FSH during the periovulatory period and ovulation during the next cycle. Biol. Reprod., 21:347-352. 358. Greenwald, G. S., and Siegel, H. I. (1982): Is the first or second periovulatory surge of FSH responsible for follicular recruitment in the hamster? Proc. Soc. Exp. Biol. Med., 170:225-230. 359. Sheela Rani, C. S., and Moudgal, N. R. (1977): Role of the pro¬ estrous surge of gonadotropins in the initiation of follicular matur¬ ation in the cyclic hamster: a study using antisera to follicle stim¬ ulating hormone and luteinizing hormone. Endocrinology, 101: 1484-1494. 360. Page, R. D., and Butcher, R. L. (1982): Follicular and plasma patterns of steroids in young and old rats during normal and prolonged estrous cycles. Biol. Reprod., 27:383-392. 361. Fevold, H. L. (1941): Synergism of follicle stimulating hormone and luteinizing hormone in producing estrogen secretion. Endocrinology, 28:33-36. 362. Greep, R. O., Van Dyke, H. B., and Chow, B. F. (1942): Gonad¬ otropins of the swine pituitary. I. Various biological effects of pu¬ rified thylakentrin (FSH) and pure metakentrin (ICSH). Endocrinol¬ ogy, 30:635-649.

440

/

Chapter

11

363. Lostroh, A., and Johnson, R. E. (1966): Amounts of interstitial cell stimulating hormone and follicle stimulating hormone required for development, uterine growth and ovulation in the hypophysectomized rat. Endocrinology, 79:991-996. 364. Erickson, G. F., Wang, C., and Hsueh, A. J. W. (1979): FSH induction of functional LH receptors in granulosa cells cultured in a chemically defined medium. Nature, 279:336-337. 365. Wang, C., Hsueh, A. J. W., and Erickson, G. F. (1981): LH stim¬ ulation of estrogen secretion in cultured granulosa cells. Mol. Cell. Endocrinol., 24:17-28. 366. Goodwin, J. A., and Terranova, P. F. (1985): Relationship between LH dependency of preantral follicles and the secondary FSH surge: effects of hypophysectomy and correlation with hCG and FSH bind¬ ing and follicular steroids. In: Proceedings of the 5th Ovarian Work¬ shop, edited by D. O. Toft and R. J. Ryan, pp. 243-248. Raven Press, New York. 367. Cameron, J. L., and Chappel, S. C. (1985): Follicle-stimulating hormone within and secreted from anterior pituitaries of female golden hamsters during the estrous cycle and after ovariectomy. Biol. Reprod., 33:132-139. 368. Sheela Rani, C. S., and Moudgal, N. R. (1977): Examination of the role of FSH in periovulatory events in the hamster. J. Reprod. Fertil 50:37-45. 369. Dorrington, J. H., Moon, Y. S., and Armstrong, D. T. (1975): Estradiol biosynthesis in cultured granulosa cells from hypophysec¬ tomized immature rats: stimulation by follicle stimulating hormone. Endocrinology, 97:1328-1331. 370. Hillier, S. G., Zeleznik, A. J., and Ross, G. T. (1978): Independence of steroidogenic capacity and luteinizing hormone receptor induction in developing granulosa cells. Endocrinology, 102:937-946. 371. Hillier, S. G., Zeleznik, A. J., Knazek, R. A., and Ross, G. T. (1980): Hormonal regulation of preovulatory follicle maturation in the rat. J. Reprod. Fertil., 60:219-229. 372. Wang, C., Hsueh, A. J. W., and Erickson, G. F. (1979): Induction of functional prolactin receptors by follicle stimulating hormone in rat granulosa cells in vivo and in vitro. J. Biol. Chem., 254 11330— 11336. 373. Jones, P. B. C., Welsh, T. H., Jr., and Hsueh, A. J. W. (1982): Regulation of ovarian progestin production by epidermal growth factor in cultured rat granulosa cells. J. Biol. Chem., 257:1126811273. 374. Ireland, J. J., and Richards, J. S. (1978): Acute effects of estradiol and FSH on specific binding of human [I125] iodo FSH to rat ovarian granulosa cells in vivo and in vitro. Endocrinology, 102:876-883. 375. Gruenberg, M. L., Steger, R. W., andPeluso, J. J. (1983): Follicular development, steroidogenesis and ovulation within ovaries exposed in vitro to hormone levels which mimic those of the rat estrous cycle Biol. Reprod., 29:1265-1275. 376. Saidapur, S., and Greenwald, G. S. (1978): Peripheral blood and ovarian levels of sex steroids in the cyclic hamster. Biol. Reprod 18:401-408. 377. Leavitt, N. N., Barcom, C. R., Bagwell, J. N., and Blaha, C. C. (1973): Structure and function of the hamster corpus luteum during the estrous cycle. Am. J. Anat., 136:235-250. 378. Terranova, P. F., and Greenwald, G. S. (1978): Steroid and gonad¬ otropin levels during the luteal-follicular shift of the cylic hamster. Biol. Reprod., 18:170-175. 379. Schwartz, N. B. (1969): A model for the regulation of ovulation in the rat. Recent Prog. Horm. Res., 25:1-55. 380. Smith, M. S., Freeman, M. E., and Neill, J. D. (1973): The control of progesterone secretion during the estrous cycle and early pseu¬ dopregnancy in the rat: prolactin, gonadotropin and steroid levels associated with rescue of the corpus luteum of pseudopregnancy. Endocrinology, 96:219-226. 381. Gallo, R. V. (1981): Pulsatile LH release during periods of low level LH secretion in the rat estrous cycle. Biol. Reprod., 24:771-777. 382. Fox, S. R., and Smith, M. S. (1985): Changes in the pulsatile pattern of luteinizing hormone secretion during the rat estrous cycle. En¬ docrinology, 116:1485-1492. 383. Szoltys, M. (1981): Oestrogens and progestagens in rat ovarian fol¬ licles during the oestrous cycle. J. Reprod. Fertil., 63:221-224. 384. Hamberger, L., Nordenstrom, K., Rosberg, S., and Sjogren, A. (1978): Acute influence of LH and FSH on cyclic AMP formation in isolated granulosa cells of the rat. Acta Endocrinol., 88:567-579.

385. Naftolin, F., Brown-Grant, K., and Corker, C. S. (1972): Plasma and pituitary luteinizing hormone and peripheral plasma oestradiol concentrations in the normal oestrous cycle of the rat and after ex¬ perimental manipulation of the cycle. J. Endocrinol., 53:17-30. 386. Kalra, S. P., and Kalra, P. S. (1974): Temporal interrelationships among circulating levels of estradiol, progesterone and LH during the rat estrous cycle. Endocrinology, 95:1711-1718. 387. Goodman, R. L. (1978): A quantitative analysis of the physiological role of estradiol and progesterone in the control of tonic and surge secretion of luteinizing hormone in the rat. Endocrinology, 102:142150. 388. Goodman, R. L., and Daniel, K. (1985): Modulation of pulsatile luteinizing hormone secretion by ovarian steroids in the rat. Biol. Reprod., 32:217-225. 389. Devorshak-Harvey, E., Peluso, J. J., Bona-Gallo, A., and Gallo, R. V. (1985): Effect of alterations in pulsatile luteinizing hormone release on ovarian follicular atresia and steroid secretion on diestrus 1 in the rat estrous cycle. Biot. Reprod., 33:103-111. 390. Okamoto, M. T., Nobunaga, T., and Suzuki, Y. (1972): Delay in ovulation with pentobarbital anesthesia applied at various stages of the 4-day cyclic rat. Endocrinol. Jpn., 19:11-17. 391. Dominquez,. R., and Smith. E. R. (1974): Barbiturate blockade of ovulation on days other than proestrus in the rat. Neuroendocrinol¬ ogy, 14:212-223. 392. Schwartz, N. B., and Gold, J. J. (1967): Effect of a single dose of anti-LH serum at proestrus on the rat estrous cycle. Anat. Rec., 157:137-150. 393. Laurence, K. A., and Ichikawa, S. (1969): Effects of antiserum to bovine LH on the estrous cycle and early pregnancy in the female rat. Int. J. Fertil., 14:8-15. 394. Greenwald, G. S. (1965): Effect of a single injection of diethylstilbestrol or progesterone on the hamster ovary. J. Endocrinol., 33:1323. 395. Greenwald, G. S. (1977): Exogenous progesterone: influence on ovulation and hormone levels in the cyclic hamster. J. Endocrinol., 73:151-155. 396. Greenwald, G. S. (1978): Modification by exogenous progesterone of estrogen and gonadotropin secretion in the cyclic hamster. En¬ docrinology, 103:2315-2322. 397. Beattie, C. W., and Corbin, C. W. (1975): The differential effects of diestrous progestogen administration on proestrous gonadotropin levels. Endocrinology, 97:885-890. 398. Garza, F., and Terranova, P. F. (1984): Inhibition of thecal androstenedione production by exogenous progesterone in the cyclic ham¬ ster. J. Reprod. Fertil., 70:493-498. 399. Taya, K , Terranova, P. F., and Greenwald, G. S. (1981): Acute effects of exogenous progesterone on follicular steroidogenesis in the cyclic rat. Endocrinology, 108:2324-2330. 400. Varga, B., and Greenwald. G. S. (1979): Cyclic changes in uteroovarian blood flow and ovarian hormone secretion in the hamster: effects of adrenocorticotropin, luteinizing hormone, and folliclestimulating hormone. Endocrinology, 104:1525-1531. 401. Varga, B., Horvath, E., Folly, G., and Stark, E. (1985): Study of the luteinizing hormone-induced increase of ovarian blood flow dur¬ ing the estrous cycle in the rat. Biol. Reprod., 32:480-488. 402. Peluso, J. J., Luttmer, S., and Gruenberg, M. L. (1984): Modulatory action of FSH on LH-induced follicular growth in rats. J Reprod Fertil., 72:173-177. 403. Greenwald, G. S. (1978): Follicular activity in the mammalian ovary. In: The Vertebrate Ovary, edited by R. E. Jones, pp. 639-689. Plenum, New York. 404. Nordenstrom, K., and Johanson, C. (1985): Steroidogenesis in iso¬ lated rat granulosa cells—changes during follicular maturation. Acta Endocrinol., 108:550-556. 405. Richards, J. S., Jonassen, J. A., and Kersey, K. A. (1980): Evidence that changes in tonic luteinizing hormone secretion determine the growth of preovulatory follicles in the rat. Endocrinology, 107:641— 647. 406. Roser, S., and Block (1971): Etude comparative des variations de la progesterone plasmatique ovarienne au cours de cycles de respectivement 4 et 5 jour, chez la ratte. C. R. Soc. Biol., 165:1995-1998 407. van der School, P., and de Greef, W. J. (1976): Dioestrous proges¬ terone and pro-estrous luteinizing hormone in 4- and 5-day cycles of female rats. J. Endocrinol., 70:61-68.

Follicular Selection / 408. Nequin, L. G., Alvarez, J., and Shwartz, N. B. (1979): Measurement of serum steroid and gonadotropin levels and uterine and ovarian variables throughout 4 day and 5 day estrous cycles in the rat. Biol. Reprod., 20:659-670. 409. van der Schoot, P., and Uilenbroek, J. T. J. (1983): Reduction of 5-day cycle length of female rats by treatment with bromocriptine. J. Endocrinol., 97:83-89. 410. Boehm, N., Plas-Roser, S., and Aron, C. (1984): Prolactin and the control of cycle length in the female rat. Acta Endocrinol., 106:188— 192. 411. Richards, J. S., and Bogovich, K. (1982): Effects of human chorionic gonadotropin and progesterone on follicular development in the im¬ mature rat. Endocrinology, 111:1429-1438. 412. Fortune, J. E., and Vincent, S. E. (1983): Progesterone inhibits the induction of aromatase activity in rat granulosa cells in vitro. Biot. Reprod., 28:1078-1089. 413. Uilenbroek, J. T. J., van der Schoot, P., den Besten, D., and Lankhorst, R. R. (1982): A possible direct effect of prolactin on follicular activity. Biol. Reprod., 27:1119-1125. 414. Magoffin, D. A., and Erickson, G. F. (1981): LH induction of androgen biosynthesis in cultured ovarian cells: inhibitory effect on prolactin. In: Dynamics of Ovarian Function, edited by N. B. Schwartz and M. Hunzicker-Dunn, pp. 55-60. Raven Press, New York. 415. L’Hermite, M., Niswender, G. D., Reichert, L. E., Jr., andMidgley, A. R., Jr. (1972): Serum follicle-stimulating hormone in sheep as measured by radioimmunoassay. Biol. Reprod., 6:325-332. 416. Goodman, R. L., Pickover, S. M., and Karasch, F. J. (1981): Ovar¬ ian feedback control of follicle-stimulating hormone in the ewe: evi¬ dence for selective suppression. Endocrinology, 108:772-777. 417. McNatty, K. P., Dobson, C., Gibb, M., Kieboom, L., andThurley, D. C. (1981): Accumulation of luteinizing hormone, oestradiol and androstenedione by sheep ovarian follicles in vivo. J. Endocrinol., 91:99-109. 418. Hauger, R. L., Karsch, F. J., and Foster. D. L.( 1977): A new concept for control of the estrous cycle of the ewe based on the temporal relationships between luteinizing hormone, estradiol and progester¬ one in peripheral serum and evidence that progesterone inhibits tonic LH secretion. Endocrinology, 101:807-817. 419. Baird, D. T., Swanston, I., and Scaramuzzi, R. J. (1976): Pulsatile release of LH and secretion of ovarian steroids in sheep during the luteal phase of the estrous cycle. Endocrinology, 98:1490-1495. 420. Brand, A., anddeJong, W. H. R. (1973): Qualitative and quantitative micromorphological investigations of the tertiary follicle population during the oestrous cycle in sheep. J. Reprod. Fertil., 33:431 —439. 421. Bherer, J., Matton, P., and Dufour, J. J. (1977): Fate of the two largest follicles in the ewe after injection of gonadotrophins at two stages of the estrus cycle. Proc. Soc. Exp. Biol. Med., 154:412414. 422. Tsonis, C. G., Cahill, L. P., Carson, R. S., and Findlay, J. K. (1984): Identification at the onset of luteolysis of follicles capable of ovulation in the ewe. J. Reprod. Fertil., 70:609-614. 423. Dailey, R. A., Fogwell, R. L., and Thayne, W. V. (1982): Distri¬ bution of visible follicles on the ovarian surface in ewes. J. Anim. Sci., 54:1196-1204. 424. Turnbull, K. E., Braden, A. W. H., and Mattner, P. E. (1977): The pattern of follicular growth and atresia in the ovine ovary. Aust. J. Biol. Sci., 30:229-241. 425. Cahill, L. P., and Mauleon, P. (1980): Influences of season, cycle and breed on follicular growth rates in sheep. J. Reprod. Fertil., 58:321-328. 426. Lahlou-Kassi, A., and Mariana, J. C. (1984): Ovarian follicular growth during the oestrous cycle in two breeds of ewes of different ovulation rate, the D’man and the Timahdite. J. Reprod. Fertil., 72:301-310. 427. Cummins, L. J., O’Shea, T., Bindon, B. M., Lee, V. W. K., and Findlay, J. K. (1983): Ovarian inhibin content and sensitivity to inhibin in Booroola and control strain Merino ewes. J. Reprod. Fertil., 67:1-7. 428. Robertson, D. M., Ellis, S., Foulds, L. M., Findlay, J. K., and Bindon, B. M. (1984): Pituitary gonadotrophins in Booroola and control Merino sheep. J. Reprod. Fertil., 71:189-197. 429. McNatty, K. P., Henderson, K. M., Lun, S., et al. (1985): Ovarian activity in Booroola X Romney ewes which have a major gene in¬ fluencing their ovulation rate. J. Reprod. Fertil., 73;109-120.

441

430. Bindon, B. M., Piper, L. R., and Thimonier, J. (1984): Preovulatory LH characteristics and time of ovulation in the prolific Booroola Merino ewe. J. Reprod. Fertil., 71:519-523. 431. Bjersing, L., Hay, M. F., Kan, G., et al. (1972): Changes in go¬ nadotrophins, ovarian steroids and follicular morphology in sheep at oestrus. J. Endocrinol., 52:465-479. 432. Hay, M. F., and Moor, R. M. (1975): Functional and structural relationships in the graafian follicle population of the sheep ovary. J. Reprod. Fertil., 45:583-593. 433. McNatty, K. P., Gibb, M., Dobson, C., Thurley, D. C., and Findlay, J. K. (1981): Changes in the concentration of gonadotrophic and steroidal hormones in the antral fluid of ovarian follicles throughout the oestrous cycle of the sheep. Aust. J. Biol. Sci., 34:67-80. 434. Moor, R. M. (1974): The ovarian follicle of the sheep: inhibition of oestrogen secretion by luteinizing hormone. J. Endocrinol., 61:455463. 435. England, B. G., Webb, R., and Dahmer, M. K. (1981): Follicular steroidogenesis and gonadotropin binding to ovine follicles during the estrous cycle. Endocrinology, 109:881-887. 436. Driancourt, M. A., Gibson, W. R.. and Cahill, L. P. (1985): Fol¬ licular dynamics throughout the oestrous cycle in sheep. A review. Reprod. Nutr. Dev., 25:1-15. 437. McNatty, K. P., Gibb, M., Dobson, C., et al. (1982): Preovulatory follicular development in sheep treated with PMSG and/or prosta¬ glandin. J. Reprod. Fertil., 65:111-123. 438. Webb, R., and England, B. G. (1982): Identification of the ovulatory follicle in the ewe: associated changes in follicular size, thecal and granulosa cell luteinizing hormone receptors, antral fluid steroids, and circulating hormones during the preovulatory period. Endocri¬ nology, 110:873-881. 439. McNeilly, A. S., Fraser, H. M., and Baird, D. T. (1984): Effect of immunoneutralization of LH releasing hormone on LH, FSH and ovarian steroid secretion in the preovulatory phase of the oestrous cycle in the ewe. J. Endocrinol., 101:213-219. 440. McNatty, K. P., Hudson, N. L., Henderson, K. M., et al. (1984): Changes in gonadotropin secretion and ovarian antral follicular ac¬ tivity in seasonally breeding sheep throughout the year. J. Reprod. Fertil., 70:309-321. 441. McLeod, B. J., Haresign, W., and Lamming, G. E. (1983): Induction of ovulation in seasonally anoestrous ewes by continuous infusion of low doses of Gn-RH. J. Reprod. Fertil., 68:489—495. 442. McNatty, K. P., Hudson, N., Gibb. M., et al. (1985): FSH influences follicle viability, oestradiol biosynthesis and ovulation rate in Rom¬ ney ewes. J. Reprod. Fertil., 75:121-131. 443. McNeilly, A. S. (1985): Effect of changes in FSH induced by bovine follicular fluid and FSH infusion in the preovulatory phase on sub¬ sequent ovulation rate and corpus luteum function in the ewe. J. Reprod. Fertil., 74:661-668. 444. Schams, D., Schallenberger, E., Hoffman, B., and Karg, H. (1977): The oestrous cycle of the cow: hormonal parameters and time rela¬ tionships concerning oestrus, ovulation, and electrical resistance of the vaginal mucus. Acta Endocrinol., 86:180-182. 445. Walters, D. L., Schams, D., and Schallenberger, E. (1984): Pulsatile secretion of gonadotrophins, ovarian steroids and ovarian oxytocin during the luteal phase of the oestrous cycle in the cow. J. Reprod. Fertil., 71:479-491. 446. Schallenberger, E., Schams, D., Bullermann, B., and Walters, D. L. (1984): Pulsatile secretion of gonadotrophins, ovarian steroids and ovarian oxytocin during prostaglandin-induced regression of the corpus luteum in the cow. J. Reprod. Fertil., 71:493-501. 447. Ireland, J. J., and Roche, J. F. (1982): Development of antral follicles in cattle after prostaglandin-induced luteolysis: changes in serum hormones, steroids in follicular fluid, and gonadotropin receptors. Endocrinology, 111:2077-2086. 448. Glencross, R. G.. Munro, I. B., Senior, B. E., and Pope, G. S. (1973): Concentrations of oestradiol- 17(3, oestrone and progesterone in jugular venous plasma of cows during the oestrous cycle and in early pregnancy. Acta Endocrinol., 73:374-384. 449. Dieleman, S. J., Kruip, T. A. M., Fontijne, P., de Jong, W. H. R., and van der Weyden, G. C. (1983): Changes in oestradiol, proges¬ terone and testosterone concentrations in follicular fluid and in the micromorphology of preovulatory bovine follicles relative to the peak of luteinizing hormone. J. Endocrinol., 97:31—42.

442

/

Chapter 11

450. Roche, J. F., and Ireland, J. J. (1981): The differential effect of progesterone on concentrations of luteinizing hormone and folliclestimulating hormone in heifers. Endocrinology, 108:568-572. 451. Dufour, J., Whitmore, H. L., Ginther, O. J., and Casida, L. E. (1972): Identification of the ovulating follicle by its size on different days of the estrous cycle in heifers. J. Anim. Sci., 34:85-87. 452. Matton, P., Adelakoun, V., Couture, Y., and Dufour, J. J. (1981): Growth and replacement of the bovine ovarian follicles during the estrous cycle. J. Anim. Sci., 52:813-820. 453. Staigmiller, R. B., and England, B. G. (1982): Folliculogenesis in the bovine. Theriogenology, 17:43-52. 454. Fogwell, R. L., Cowley, J. L., Wortman, J. A., Ames, N. K., and Ireland, J. J. (1985): Luteal function in cows following destruction of ovarian follicles at midcycle. Theriogenology, 23:389-398. 455. Choudary, J. B., Gier, H. T., and Marion, G. B. (1968): Cyc¬ lic changes in bovine vesicular follicles. J. Anim. Sci., 27:468471. 456. Scaramuzzi, R. J., Turnbull, K. E., and Nancarrow, C. D. (1980): Growth of graafian follicles in cows following luteolysis induced by the prostaglandin F2a analogue, cloprostenol. Aust. J. Biol. Sci., 33:63-69. 457. Kruip, T. A. M., and Dieleman, S. J. (1982): Macroscopic classi¬ fication of bovine follicles and its validation by micromorphological and steroid biochemical procedures. Reprod. Nutr. Dev., 22:465473. 458. Merz, E. A., Hauser, E. R., and England, B. G. (1981): Ovarian function in the cycling cow: relationship between gonadotropin bind¬ ing to theca and granulosa and steroidogenesis in individual follicles. J. Anim. Sci., 52:1457-1468. 459. Henderson, K. M., McNeilly, A. S., and Swanston, I. A. (1982): Gonadotrophin and steroid concentrations in bovine follicular fluid and their relationship to follicle size. J. Reprod. Fertil., 65:467473. 460. Fortune, J. E., and Hansel, W. (1985): Concentrations of steroids and gonadotropins in follicular fluid from normal heifers and heifers primed for superovulation. Biol. Reprod., 32:1069-1079. 461. Staigmiller, R. B., England, B. G., Webb, R., Short, R. E., and Bellows, R. A. (1982): Estrogen secretion and gonadotropin binding by individual bovine follicles during estrus. J. Anim. Sci., 55:14731482. 462. Thayer, K. M., Forrest, D. W., and Welsh, T. H., Jr. (1985): Real¬ time ultrasound evaluation of follicular development in superovulated cows. Theriogenology, 23:233. 463. van de Wiel, D. F. M., Erkens, J., Koops, W., Vos, E., and van Landeghem, A. A. J. (1981): Periestrous and midluteal time courses of circulating LH, FSH, prolactin, estradiol-17(3 and progesterone in the domestic pig. Biol. Reprod., 24:223-233. 464. Magness, R. R., Christenson, R. K., and Ford, S. P. (1983): Ovarian blood flow throughout the estrous cycle and early pregnancy in sows. Biol. Reprod., 28:1090-1096. 465. Clark, J. R., Brazier, S. G., Wiginton, L. M., Stevenson, G. R., and Tribble, L. F. (1982): Time of ovarian follicle selection during the porcine estrous cycle. Theriogenology, 18:697-709. 466. Parlow, A. F., Anderson, L. L., and Melampy, R. M. (1964): Pituitary follicle-stimulating hormone and luteinizing hormone con¬ centrations in relation to reproductive stages of the pig. Endocri¬ nology, 75:365-376. 467. Guthrie, H. D., and Knudsen, J. F. (1984): Follicular growth and production of estrogen and progesterone after injection of gilts with human chorionic gonadotropin on day 12 of the estrous cycle. J. Anim. Sci., 59:1295-1302. 468. Hunter, R. H. F., and Baker, T. G. (1975): Development and fate of porcine graafian follicles identified at different stages of the oestrous cycle. J. Reprod. Fertil., 43:193-196. 469. Dailey, R. A., Clark, J. R., Staigmiller, R. B., First, N. L., Chap¬ man, A. B., and Casida, L. E. (1976): Growth of new follicles following electrocautery in four genetic groups of swine. J. Anim. Sci., 43:175-183. 470. Bamberg, E., Choi, H. S., Hassaan, N. K., Klaring, W. J., Mostl, E., and Stockl, W. (1980): Steroidhormongehalt in Blut und Ovarfollikeln des Rindes wahrend des Zyklus. Zbl. Vet. Med., 27:186194. 471. Ainsworth, L., Tsang, B. K., Downey, B. R., Marcus, G. J., and Armstrong, D. T. (1980): Interrelationships between follicular fluid

steroid levels, gonadotropic stimuli, and oocyte maturation during preovulatory development of porcine follicles. Biol. Reprod., 23:621—

621. 472. Koering, M» J. (1969): Cyclic changes in ovarian morphology during the menstrual cycle in Macaca mulatto. Am. J. Anat., 126:73-101. 473. Koering, M. J., Baehler, E. A., Goodman, A. L., and Hodgen, G. D. (1982): Developing morphological asymmetry of ovarian fol¬ licular maturation in monkeys. Biol. Reprod., 27:989-997. 474. Stouffer, R. L., Hodgen, G. D., Ottobre, A. C., and Christian, C. D. (1984): Follicular fluid treatment during the follicular versus luteal phase of the menstrual cycle: effects on corpus luteum func¬ tion. J. Clin. Endocrinol. Metab., 48:1027-1033. 475. Zeleznik, A. J., Wildt, L., and Schuler, H. M. (1980): Character¬ ization of ovarian folliculogenesis during the luteal phase of the menstrual cycle in rhesus monkeys using [3H]thymidine autoradiog¬ raphy. Endocrinology, 107:982-988. 476. Ellinwood, W. E., Norman, R. L., and Spies, H. G. (1984): Chang¬ ing frequency of pulsatile luteinizing hormone and progesterone se¬ cretion during the luteal phase of the menstrual cycle of rhesus mon¬ keys. Biol. Reprod., 31:714-722. 477. Clark, J. R., Dierschke, D. J., Meller, P. A., and Wolf, R. C. (1979): Hormonal regulation of ovarian folliculogenesis in rhesus „ monkeys. II. Serum concentrations of estradiol-17p and follicle stim¬ ulating hormone associated with growth and identification of the preovulatory follicle. Biol. Reprod., 21:497-503. 478. Clark, J. R., Dierschke, D. J., and Wolf, R. C. (1978): Hormonal regulation of ovarian folliculogenesis in rhesus monkeys: I. Con¬ centrations of serum luteinizing hormone and progesterone during laparoscopy and patterns of follicular development during successive menstrual cycles. Biol. Reprod., 17:779-783. 479. Goodman, A. L., Nixon, W. E., Johnson, D. K., and Hodgen, G. D. (1977): Regulation of folliculogenesis in the cycling rhesus monkey: selection of the dominant follicle. Endocrinology, 100: 155-161. 480. Goodman, A. L., Nixon, W. E., and Hodgen, G. D. (1979): Between-ovary interaction in the regulation of follicle growth, corpus luteum function, and gonadotropin secretion in the primate ovarian cycle. III. Temporal and spatial dissociation of folliculogenesis and negative feedback regulation of tonic gonadotropin release after luteectomy in rhesus monkeys. Endocrinology, 105:69-73. 481. diZerega, G. S., and Hodgen, G. D. (1982): The interovarian pro¬ gesterone gradient: a spatial and temporal regulator of folliculo¬ genesis in the primate ovarian cycle. J. Clin. Endocrinol. Metab., 54:495-499. 482. diZerega, G. S., Campeau, J. D., Ujita, E. L., et al. (1983): The possible role for a follicular protein in the intraovarian regulation of folliculogenesis. Sem. Reprod. Endocrinol., 1:309-320. 483. Channing, C. P., Anderson, L. D., Hoover, D. J., Gagliano, P., and Hodgen, G. (1981): Inhibitory effects of porcine follicular fluid on monkey serum FSH levels and follicular maturation. Biol. Re¬ prod., 25:885-903. 484. diZerega, G. S., Turner, C. K., Stouffer, R. L., Anderson, L. D., Channing, C. P., and Hodgen, G. D. (1981): Suppression of folliclestimulating hormone-dependent folliculogenesis during the primate ovarian cycle. J. Clin. Endocrinol. Metab., 52:451-456. 485. Zeleznik, A. J., Hutchison, J. S., and Schuler, H. M. (1985): In¬ terference with the gonadotropin-suppressing actions of estradiol in macaques overrides the selection of a single preovulatory follicle. Endocrinology, 117:991-999. 486. Dierschke, D. J.. Hutz, R. J., and Wolf, R. C. (1985): Induced follicular atresia in rhesus monkeys: strength-duration relationships of the estrogen stimulus. Endocrinology, 117:1397-1403. 487. Williams, R. F., and Hodgen, G. D. (1980): Disparate effects of human chorionic gonadotropin during the late follicular phase in monkeys: normal ovulation, follicular atresia, ovarian acyclicity, and hypersecretion of follicle-stimulating hormone. Fertil. Steril 33 64-

68. 488. Wilks, J. W., Spilman, C. H., and Campbell, J. A. (1983): Arrest of folliculogenesis and inhibition of ovulation in the monkey follow¬ ing weekly administration of progestins. Fertil. Steril., 40:688-692. 489. Channing, C. P. (1980): Progesterone and estrogen secretion by cultured monkey ovarian cell types: influences of follicular size, serum luteinizing hormone levels, and follicular fluid estrogen levels. Endocrinology, 107:342-352.

Follicular Selection /

490. Vernon, M. W., Dierschke, D. J., Sholl, S. A., and Wolf, R. C. (1983): Ovarian aromatase activity in granulosa and theca cells of rhesus monkeys. Biol. Reprod.. 28:342-349. 491. Marut, E. L., Huang, S. C., and Hodgen, G. D. (1983): Distin¬ guishing the steroidogenic roles of granulosa and theca cells of the dominant ovarian follicle and corpus luteum. J. Clin. Endocrinol. Metab., 57:925-930. 492. Block, E. (1951): Quantitative morphological investigations of the follicular system in women. Variations in the different phases of the sexual cycle. Acta Endocrinol., 8:33-54. 493. Gougeon, A. (1981): Rate of follicular growth in the human ovary. In: Follicular Maturation and Ovulation—Proceedings of the IVth Reinier de Graaf Symposium, International Congress Ser. No. 560, edited by R. Rolland, E. V. van Hall, S. G. Hillier, K. P. McNatty, and J. Schoemaker, pp. 155-163. Excerpta Medica, Princeton, NJ. 494. Gougeon, A. (1984): Vitesse de croissance des follicules dans l’ovaire humain. Contraception-Fertilite-Sexualite, 12:839-845. 495. Gougeon, A. (1984): Influence des variations hormonales cycliques (steroides et gonadotropines) sur la croissance folliculaire dans l’ovaire humain. Contraception-Fertilite-Sexualite, 12:615-620. 496. Gougeon, A., and Lefevre, B. (1984): Histological evidence of al¬ ternating ovulation in women. J. Reprod. Fertil., 70:7-13. 497. Gougeon, A. (1984): Croissance folliculaire dans l’ovaire humain pendant le cycle menstruel: mise en evidence de regulations intraovariennes. Contraception-Fertilite-Sexualite, 12:733—738. 498. Gougeon, A. (1984): Le follicule ovulatoire humain. A quel moment du cycle est-il selectionne et par quels mecanismes? Une tentative de reponse. Contraception-Fertilite-Sexualite, 12:1397-1405. 499. McNatty, K. P. (1981): Hormonal correlates of follicular develop¬ ment in the human ovary. Aust. J. Biol. Sci., 34:249-268. 500. Lloyd, C. W., Lobotsky, J., Baird, D. T., et al. (1971): Concen¬ tration of unconjugated estrogens, androgens and gestagens in ovarian and peripheral venous plasma of women: The normal menstrual cycle. J. Clin. Endocrinol., 32:155-166. 501. deJong, F. H., Baird, D. T., and van der Molen, H. J. (1974): Ovarian secretion rates of oestrogens, androgens and progesterone in normal women and in women with persistent ovarian follicles. Acta Endocrinol., 77:575-587. 502. Baird, D. T., and Fraser, I. S. (1974): Blood production and ovarian secretion rates of estradiol-173 ar>d estrone in women throughout the menstrual cycle. J. Clin. Endocrinol. Metab., 38:1009-1017. 503. McNatty, K. P., Hunter, W. M., McNeilly, A. S., and Sawers, R. S. (1975): Changes in the concentration of pituitary and steroid hormones in the follicular fluid of human graafian follicles through¬ out the menstrual cycle. J. Endocrinol., 64:555-571. 504. McNatty, K. P., and Baird, D. T. (1978): Relationship between follicle-stimulating hormone, androstenedione and oestradiol in hu¬ man follicular fluid. J. Endocrinol., 76:527-531. 505. Queenan, J. T., O’Brien, K. G. D., Bains, L. M., Simpson, J., Collins, W. P., and Campbell, S. (1980): Ultrasound scanning of ovaries to detect ovulation in women. Fertil. Steril., 34:99-105. 506. McNatty, K. P. (1978): Cyclic changes in antral fluid hormone con¬ centrations in humans. Clin. Endocrinol. Metab., 7:577-600. 507. Bieszczad, R. R., McClintock, J. S., Pepe, G. J., and Dimino, M. J. (1982): Progesterone secretion by granulosa cells from dif¬ ferent sized follicles of human ovaries after short term incubation. J. Clin. Endocrinol. Metab., 55:181-184. 508. Dennefors, B. L., Nilsson, L., and Hamberger, L. (1982): Steroid and adenosine 3', 5'-monophosphate formation in granulosa and thecal cells from human preovulatory follicles in response to hu¬ man chorionic gonadotropin. J. Clin. Endocrinol. Metab., 54:436441. 509. Bemardus, R. E., Jones, G. S., Acosta, A. A., et al. (1985): The significance of the ratio in follicle-stimulating hormone and lutein¬ izing hormone in induction of multiple follicular growth. Fertil. Steril., 43:373-378. 510. Gougeon, A., Lefevre, B., and Testart, J. (1984): Recrutement et selection du follicule dominant pendant le cycle menstruel spontane ou stimule chez la femme. In: Periode Peri-ovulatoire. Colloque de la societe francaise pour l’etude de lafertilite, pp. 1—11. 511. Nilsson, L., Wikland, M., and Hamberger, L. (1982): Recruitment of an ovulatory follicle in the human following follicle-ectomy and luteectomy. Fertil. Steril., 37:30-34.

443

512. Araki, S., Chikazawa, K., Akabori, A., Ijima, K., and Tamada, T. (1983): Hormonal profile after removal of the dominant follicle and corpus luteum in women. Endocrinol. Jpn., 30:55-70. 513. diZerega, G. S., Marrs, R. P., Roche, P. C., Campeau, J. D., and Kling, O. R. (1983): Identification of proteins in pooled human follicular fluid which suppress follicular response to gonadotropins. J. Clin. Endocrinol. Metab., 56:35-41. 514. diZerega, G. S., Marrs, R. P., Campeau, J. D., and Kling, O. R. (1983): Human granulosa cell secretion of protein(s) which suppress follicular response to gonadotropins. J. Clin. Endocrinol. Metab., 56:147-155. 515. Chari, S., Daume, E., Sturm, G., Vaupel, H., and Schuler, I. (1985): Regulators of steroid secretion and inhibin activity in human ovarian follicular fluid. Mol. Cell. Endocrinol., 41:137-145. 516. Greenwald, G. S., and Choudary, J. B. (1969): Follicular devel¬ opment and induction of ovulation in the pregnant mouse. Endocri¬ nology, 84:1512-1516. 517. Murr, S. M., Bradford, G. E., and Geschwind, I. I. (1974): Plasma luteinizing hormone, follicle-stimulating hormone and prolactin dur¬ ing pregnancy in the mouse. Endocrinology, 94:112-116. 518. Greenwald, G. S. (1966): Ovarian follicular development and pi¬ tuitary FSH and LH content in the pregnant rat. Endocrinology, 79:572-578. 519. Taya, K., and Sasamoto, S. (1977): Induction of ovulation by ex¬ ogenous gonadotrophin during pseudopregnancy, pregnancy or lac¬ tation in rats. J. Reprod. Fertil., 51:467^168. 520. Taya, K., and Greenwald, G. S. (1981): In vivo and in vitro ovarian steroidogenesis in the pregnant rat. Biol. Reprod., 25:683-691. 521. Taya, K., Kimura, J., and Sasamoto, S. (1984): Inhibin activity in ovarian venous plasma during pregnancy, pseudopregnancy and lac¬ tation in the rat. Endocrinol. Jpn., 31:427-433. 522. Richards, J. S., and Kersey, K. A. (1979): Changes in theca and granulosa cell function in antral follicles developing during pregnancy in the rat: gonadotropin receptors, cyclic AMP and estradiol-17(3. Biol. Reprod., 21:1185-1201. 523. Carson, R. S., Richards, J. S., and Kahn, L. E. (1981): Functional and morphological differentiation of theca and granulosa cells during pregnancy in the rat: dependence on increased basal luteinizing hor¬ mone activity. Endocrinology, 109:1433-1441. 524. Bogovich, K., Richards, J. S., and Reichert, L. E., Jr. (1981): Obligatory role of luteinizing hormone (LH) in the initiation of pre¬ ovulatory follicular growth in the pregnant rat: specific effects of human chorionic gonadotropin and follicle-stimulating hormone on LH receptors and steroidogenesis in theca, granulosa and luteal cells. Endocrinology, 109:860-867. 525. Bogovich, K., and Richards, J. S. (1982): Androgen biosynthesis in developing ovarian follicles: evidence that luteinizing hormone reg¬ ulates thecal 17a-hydroxylase and C17-20-lyase activities. Endocri¬ nology, 111:1201-1208. 526. Greenwald, G. S. (1967): Induction of ovulation in the pregnant hamster. Am. J. Anat., 121:249-258. 527. Bast, J. D., and Greenwald, G. S. (1974): Daily concentrations of gonadotrophins and prolactin in the serum of pregnant or lactating hamsters. J. Endocrinol., 63:527-532. 528. Greenwald, G. S., Voogt, J. L., and Limback, D. (1984): In vitro follicular and luteal steroidogenesis in the pregnant hamster with preliminary studies in the rat. Biol. Reprod., 30:93-104. 529. Adams, C. E. (1968): Ovarian response to human chorionic gonado¬ trophin and egg transport in the pregnant and post-parturient rabbit. J. Endocrinol., 40:101-105. 530. Osteen, K. G., and Mills, T. M. (1979): Serum LH and FSH levels in the pregnant rabbit. Proc. Soc. Exp. Biol. Med., 162:454—457. 531. Osteen, K. G., and Mills, T. M. (1980): Changes in the size, dis¬ tribution and steroid content of rabbit ovarian follicles during early pseudopregnancy. Biol. Reprod., 22:1040-1046. 532. Labhsetwar, A. P., and Diamond, M. (1970): Ovarian changes in the guinea pig during various reproductive stages and steroid treat¬ ments. Biol. Reprod., 2:53-57. 533. Bujard, E. (1953): L’ovaire de cobaye (etudes statistiques des fol¬ licules ovariques) I. L’ovaire gravide. Rev. Suisse Zool., 60:615652. 534. Rowlands, I. W. (1956): The corpus luteum of the guinea pig. In: Cl BA Foundation Colloquia on Ageing, Vol. 2, edited by G. Wolstenholme and E. Millar, pp. 69-85. Little, Brown, Boston.

444

/ Chapter 11

535. Melampy, R. M., Henricks, D. M., Anderson, L. L., Chen, C. L., and Schultz, J. R. (1966): Pituitary follicle-stimulating hormone and luteinizing hormone concentrations in pregnant and lactating pigs. Endocrinology, 78:801-804. 536. Dufour, J. J., and Fahmy, M. H. (1974): Follicular and luteal changes during early pregnancy in three breeds of swine. Can. J. Anim. Sci., 54:29-33. 537. Schallenberger, E., Rampp, J., and Walters, D. L. (1985): Gonad¬ otrophins and ovarian steroids in cattle. II. Pulsatile changes of con¬ centrations in the jugular vein throughout pregnancy. Acta Endocri¬ nol., 108:322-330. 538. Schallenberger, E., Schondorfer, A. M., and Walters, D. L. (1985): Gonadotrophins and ovarian steroids in cattle. I. Pulsatile changes of concentrations in the jugular vein throughout the oestrous cycle. Acta Endocrinol., 108:312-321. 539. Govan, A. D. T. (1968): The human ovary in early pregnancy. J. Endocrinol., 40:421-428. 540. Govan, A. D. T. (1970): Ovarian follicular activity in late pregnancy. J. Endocrinol., 48:235-241. 541. Stamp, J., and Visfeldt, J. (1974): Ovarian morphology in early and late human pregnancy. Acta Obstet. Gynecol. Scand., 53:211-218. 542. Dekel, N., David, M. P., Yedwab, G. A., and Kraicer, P. F. (1977): Follicular development during late human pregnancy. Int. J. Fertil., 22:24-29. 543. Dennefors, B. L., and Nilsson, L. (1981): Steroid production and responsiveness to gonadotropin and prostaglandins of human ovarian follicles at term pregnancy. Fertil. Steril., 35:232-233. 544. Westergaard, L., McNatty, K. P., and Christensen, I. J. (1985): Steroid concentrations in fluid from human ovarian antral follicles during pregnancy. J. Endocrinol., 107:133-136. 545. Parlow, A. F., Daane, T. A., and Dignam. W. J. (1970): On the concentration of radioimmunoassayable FSH circulating in blood throughout human pregnancy. J. Clin. Endocrinol., 31:213-214. 546. Mishell, D. R , Jr., Thomeycroft, I. H., Nagata, Y., and Nakamura, R. M. (1973): Steroid and gonadotropin levels in normal pregnancies and pregnancies following HMG therapy. In: Gonadotropin in Fe¬ male Infertility, edited by E. Rosemberg, pp. 201-207. Excerpta Medica, Amsterdam. 547. Ranta, T., Chen, H. C., Shimohigashi, Y., Baukal, A. J., Knecht, M., and Catt, K. (1985): Enhanced follicle-stimulating hormone activity of deglycosylated human chorionic gonadotropin in ovarian granulosa cells. Endocrinology, 116:59-64. 548. Taya, K., and Greenwald, G. S. (1982): Mechanisms of suppression of ovarian follicular development during lactation in the rat. Biol. Reprod., 27:1090-1101. 549. Taya, K., and Sasamoto, S. (1980): Initiation of follicular maturation and ovulation after removal of the litter from the lactating rat. J. Endocrinol., 87:393-400. 550. Tsai-Morris, C. H., Ghosh, M., Hirshfield, A. N., Wise, P. M., and Brodie, A. M. H. (1983): Inhibition of ovarian aromatase by prolactin in vivo. Biol. Reprod., 29:342-346. 551. Selmanoff, M., and Selmanoff, C. (1983): Role of pup age, estradiol 17-0 and pituitary responsiveness in the differences in the sucklinginduced prolactin response during early and late lactation. Biol. Re¬ prod., 29:400-411. 552. Bruce, H. M. (1961): Observations on the suckling stimulus and lactation in the rat. J. Reprod. Fertil., 2:17-34. 553. Gosden, R. G., Russell, J. A., Clarke, J., and Piper, I. (1981): Effects of inhibiting prolactin secretion on the maintenance of em¬ bryonic diapause in the suckling rat. J. Endocrinol., 88:197-203. 554. Hansen, S., Sodersten, P., and Eneroth, P. (1983): Mechanisms regulating hormone release and the duration of dioestrus in the lac¬ tating rat. J. Endocrinol., 99:173-180. 555. Smith, M. S. (1981): Site of action of prolactin in the suppression of gonadotropin secretion during lactation in the rat: effect on pi¬ tuitary responsiveness to LHRH. Biol. Reprod., 24:967-976. 556. Reeves, J. J., Tamavsky, G. K., and Platt, T. (1982): Pituitary and ovarian luteinizing hormone releasing hormone receptors during the estrous cycle, pregnancy and lactation in the rat. Biol. Reprod., 27:316-319. 557. Smith, M. S. (1984): Effects of the intensity of the suckling stimulus and ovarian steroids on pituitary gonadotropin-releasing hormone receptors during lactation. Biol. Reprod., 31:548-555.

558. Smith, M. S. (1982): Effect of pulsatile gonadotropin-releasing hor¬ mone on the release of luteinizing hormone and follicle-stimulating hormone in vitro by anterior pituitaries from lactating and cycling rats. Endocrinology, 110:882-890. 559. Greenwald, G. S. (1958): A histological study of the reproductive tract of the lactating mouse. J. Endocrinol., 17:17-23. 560. Greenwald, G. S. (1965): Histologic transformation of the ovary of the lactating hamster. Endocrinology, 77:641-650. 561. Bridges, R. S., and Goldman, B. D. (1975): Diurnal rhythms in gonadotropins and progesterone in lactating and photoperiod induced acyclic hamsters. Biol. Reprod., 13:617-622. 562. Garcia-Winder, M., Imakawa, K., Day, M. L., Zalesky, D. D., Kittok, R. J., and Kinder, J. E. (1984): Effect of suckling and ovariectomy on the control of luteinizing hormone secretion during the postpartum period in beef cows. Biol. Reprod., 31:771-778. 563. Beilin, M. E., Hinshelwood, M. M., Hauser, E. R., and Ax, R. L. (1984): Influence of suckling and side of corpus luteum or pregnancy on folliculogenesis in postpartum cows. Biol. Reprod., 31:849-855. 564. Dufour, J. J., and Roy, G. L. (1985): Distribution of ovarian fol¬ licular populations in the dairy cow within 35 days after parturition J. Reprod. Fertil., 73:229-235. 565. Walters, D. L., Kaltenbach, C. C., Dunn, T. G., and Short, R. E. (1982): Pituitary and ovarian function in postpartum beef cows. I. Effect of suckling on serum and follicular fluid hormones and fol¬ licular gonadotropin receptors. Biol. Reprod., 26:640-646. 566. Walters, D. L., Short, R. E., Convey, E. M., Staigmiller, R. B., Dunn, T. G., and Kaltenbach, C. C. (1982): Pituitary and ovarian function in postpartum beef cows. II. Endocrine changes prior to ovulation in suckled and nonsuckled postpartum cows compared to cycling cows. Biol. Reprod., 26:647-654. 567. Williams, G. L., Talavera, F., Petersen, B. J., Kirsch, J. D., and Tilton, J. E. (1983): Coincident secretion of follicle-stimulating hor¬ mone and luteinizing hormone in early postpartum beef cows: effects of suckling and low-level increases of systemic progesterone Biol Reprod., 29:362-373. 568. Peters, A. R., Pimentel, M. G., and Lamming, G. E. (1985): Hor¬ mone responses to exogenous GnRH pulses in post-partum dairy cows. J. Reprod. Fertil., 75:557-565. 569. Webb, R., Lamming, G. E., Haynes, N. B., and Foxcroft, G. R. (1980): Plasma progesterone and gonadotrophin concentrations and ovarian activity in post-partum dairy cows. J. Reprod. Fertil 59-133143. 570. Walters, D. L., Short, R. E., Convey, E. M., Staigmiller, R. B., Dunn, T. G., and Kaltenbach, C. C. (1982): Pituitary and ovarian function in postpartum beef cows. III. Induction of estrus, ovulation and luteal function with intermittent small-dose injections of GnRH. Biol. Reprod., 26:655-662. 571. Kunavongkrit, A., Einarsson, S., and Settergren, I. (1982): Follicular development in primiparous lactating sows. Anim. Reprod. Sci 5:47-56. 572. Cox, N. M., and Britt, J. H. (1982): Relationships between endog¬ enous gonadotropin-releasing hormone, gonadotropins, and follicular development after weaning in sows. Biol. Reprod., 27:70-78. 573. Edwards, S., and Foxcroft, G. R. (1983): Endocrine changes in sows weaned at two stages of lactation. J. Reprod. Fertil., 67:161-172. 574. Stevenson, J. S., Cox, N. M., and Britt, J. H. (1981): Role of the ovary in controlling luteinizing hormone, follicle-stimulating hor¬ mone, and prolactin secretion during and after lactation in pigs Biol Reprod., 24:341-353. 575. Shaw, H. J., and Foxcroft, G. R. (1985): Relationships between LH, FSH and prolactin secretion and reproductive activity in the weaned sow. J. Reprod. Fertil., 75:17-28. 576. Cox, N. M., and Britt, J. H. (1982): Pulsatile administration of gonadotropin releasing hormone to lactating sows: endocrine changes associated with induction of fertile estrus. Biol. Reprod., 27:1126577. Plant, T. M., Schallenberger, E., Hess, D. L., McCormack J T Dufy-Barbe, L., and Knobil, E. (1980): Influence of suckling on gonadotropin secretion in the female rhesus monkey (Macaca mu¬ latto). Biol. Reprod., 23:760-766. 578. Weiss, G., Butler, W. R., Dierschke, D. J., and Knobil, E. (1976): Influence of suckling on gonadotropin secretion in the postpartum rhesus monkey. Proc. Soc. Exp. Biol. Med., 153:330-331.

Follicular Selection 579. Williams, R. F., and Hodgen, G. D. (1980): Reinitiation of the diurnal rhythm ot prolactin secretion in postpartum rhesus monkeys Biol. Reprod., 23:276-280. 580. Richardson, D. W„ Goldsmith, L. T„ Pohl, C. R„ Schallenberger, E., and Knobil, E. (1985): The role of prolactin in the regulation of the primate corpus luteum. J. Clin. Endocrinol. Metab 60501504. 581. Thomson, A. M., Hytten, F. E., and Black, A. E. (1975): Lactation and reproduction. Bull. WHO, 52:337-349. 582. Howie, P. W , and McNeilly, A. S. (1982): Effect of breast-feeding patterns on human birth intervals. J. Reprod. Fertil., 65:545-557. 583. Jeppsson, S., Rannevik, G., Thorell. J. I., and Wide, L. (1977): Influence of LH/FSH releasing hormone (LRH) on the basal secretion of gonadotrophins in relation to plasma levels of oestradiol, proges¬

584.

585.

586. 587.

/

445

terone and prolactin during the post-partum period in lactating and in non-lactating women. Acta Endocrinol., 84:713-728. Baird, D. T., McNeilly, A. S., Sawers, R. S., and Sharpe, R. M. (1979): Failure of estrogen-induced discharge of luteinizing hormone in lactating women. J. Clin. Endocrinol. Metab., 49:500-506. Keye, W. R., Jr., and Jaffe, R. B. (1976): Changing patterns of FSH and LH response to gonadotropin-releasing hormone in the puerperium. J. Clin. Endocrinol. Metab., 42:1133-1138. McNeilly, A. S. (1980): Prolactin and the control of gonadotrophin secretion in the female. J. Reprod. Fertil., 58:537-549. McNeilly, A. S., Glasier, A., Jonassen, J., and Howie, P. W. (1982): Evidence for direct inhibition of ovarian function by prolactin. J. Reprod. Fertil., 65:559-569.

The Physiology of Reproduction, edited by E. Knobil and J. Neill et al. Raven Press, Ltd., New York © 1988.

CHAPTER 12

Mechanism of Mammalian Ovulation Harry Lipner Historical Background, 447 Smooth Muscle, 447 • The Vascular System, 449 • Depolymerization of Mucopolysaccharide, 449 • Neu¬ ral Control of Ovulation, 449 The Structure of the Ovarian Follicle, 450 General Features of Follicular Morphology, 450 • Detailed Features of Follicular Morphology, 450 The Neural Innervation of the Ovary, 454 Neural Innervation of Ovary and Follicles, 454 • Physio¬ logical Studies, 455 The Ovarian Blood Supply, 459 Morphology, 459 • Quantitation of the Preovulatory Blood Flow, 460 • Physiological Changes in Blood Flow, 460 • Vasodilatory Substances and Ovarian Blood Flow, 460

The Role of Smooth Muscle in Ovulation, 462 Follicular Fluid, 462 Physical Characteristics, 462 • Proteins, 463 • Hormones, 463 • Enzymes, 463 • Proteoglycans, 464 Preovulatory Morphological Changes Associated with Ovulation, 465 Preovulatory Chemical Changes Associated with Ovu¬ lation, 466 Protein Synthesis, 466 • Steroidogenesis, 466 • Prosta¬ glandins, 468 • Plasminogen, 472 • Collagenolysis, 475 Summary, 475 Acknowledgments, 477 References, 477

Ovulation marks the culmination of a series of events initiated by the surge of luteinizing hormone (LH) and char¬ acterized by resumption of meiosis and germinal vesicle breakdown, initiation of luteinization of the granulosa cells, and restructuring of the follicle wall, with resultant follicular rupture and release of a mature fertilizable ovum. Study of the control of ovulation has spawned a large literature, in¬ cluding that on hormonal controls and detailed morpholog¬ ical and biochemical analyses of the preovulatory follicle. These contributions have extended our understanding of the cellular mechanisms associated with steroidal and nonsteroi¬ dal biosynthesis, collagenolysis, and vascular changes as¬ sociated with ovulation but have not yet fully explained the intimate details of follicular rupture. There are a number of reviews that offer interesting insights into the phenomenon of ovulation (34,48,103,144,151-153,155,201,292,310 317,350,386,422).

occupied workers for the next 170 years in an attempt to settle a raging argument over whether there is smooth muscle in the follicle wall. When the question was resolved in the affirmative, the role of smooth muscle in ovulation was disputed and finally characterized as probably conducive but not essential. Beginning 70 years ago, the suggestion of a role for proteolytic enzymes initiated a new and con¬ tinuing study of their presence, substrate(s), and controls. A role for the nerve supply to the ovary was bitterly dis¬ puted, the answer depending on whether one accepted that nerves penetrated only to the follicle, supplied only the outer two layers, or reached all the layers of the follicle. Not to be excluded in the study of ovulation is the control of the vascular supply to both ovary and follicle, as changes in blood flow manifested by the follicle “blush” have been a subject of intense examination almost from the suggestion 130 years ago that the ovary is an erectile organ (425).

HISTORICAL BACKGROUND

Smooth Muscle

The study of ovulation began with the discovery of the follicle (227) and that it contained an ovum (118). The cellular composition of the ovary and the follicle wall then

Regnier de Graaf (227) described the ovary as differing from the testis because it lacked tubules and instead had vesicles on its surface, which he mistakenly called eggs or

447

448

/ Chapter 12

ova. One hundred twenty-five years later, Cruikshank (118) demonstrated ova in rabbit fallopian tubes. This observation initiated the search for an explanation of follicle rupture. Von Kolliker (258) was the first to note smooth muscle in the ovary, and Aeby (4,5) described smooth muscle in the frog ovary, suggested it was involved in the discharge of ova, and explained the observations by Pfluger (399) that frog ovaries showed spontaneous movements. Rouget (425) assumed that smooth muscle around the blood vessels in the ovary and mesovarium allowed the ovary to behave as an erectile organ, discharging ova as the erect penis dis¬ charged semen. This concept was accepted as late as 1905 by Heape (209). Grohe (200) described smooth muscle in pig ovaries in the hilus, extending to and encircling the follicle and in close association with blood vessels. Heape (209) noted that rabbits ovulate approximately 10 hr after mating and that only estrus does permit coition. Mature follicles in unmated rabbits fail to release their ova, and the ova and follicles degenerate and regress. He further dem¬ onstrated that an intact blood supply to the ovary was es¬ sential for follicle rupture but that the presence of sperm was unnecessary. He described growth of the follicles, the appearance of liquor folliculi, and the thinning of the wall to the point at which the contained ovum became visible, the blood vessels around the maturing follicle increased in number, enlarged, and became congested, and the follicle projected above the ovarian surface. After mating, the swol¬ len Graafian follicle underwent changes involving the re¬ lationship of the cells of the corona radiatum to the ovum, and approximately 10 hr after mating, follicle rupture oc¬ curred. Heape noted that pressure in the follicle was not an adequate explanation because unmated rabbits also had en¬ larged, projecting follicles. He thought that the innervation of the ovary and the possible behavior of the ovary as an erectile tissue made for a plausible explanation. Despite the earlier histological evidence of smooth muscle in the ovary and follicle, von Winiwarter and Sainmont (506) failed to demonstrate that either electrical or chemical stimuli caused follicle rupture, but they proceeded to demonstrate what they interpreted as smooth muscle in the cat ovarian follicle wall and subsequently in the human follicle (505). Thomson (466) introduced what should be called the “multifactor hypothesis of ovulation.” He suggested that the distribution of smooth muscle in the follicle wall, for whose presence he largely depended on the reports of earlier work¬ ers and his own observations based on tinctorial character¬ istics of tissues, conformed closely to the distribution of blood vessels and the capillary wreath previously described by Clark (108). Thomson (466) found smooth muscle in the human ovary to be less abundant than in other species but present in sufficient quantity to act as he thought it did in those species. He suggested that the increase in follicle size was caused by the increase in volume of the liquor folliculi and a consequent increase in intrafollicular pressure and that enzymes in the liquor folliculi, whose presence was dem¬

onstrated by Schochet (428), caused digestion of the theca, leading to ovulation. Thomson (466), as had Heape (209) and von Winiwarter (505) before him, accepted that there was an intimate association between ovulation and men¬ struation and that the vascular changes associated with the latter induced the vascular changes involved in the former. He therefore assumed that the increase in follicular volume must be caused by an increase in transudation resulting from vascular engorgement and that the combination of increased fluid, proteolysis of the theca, and smooth muscle contrac¬ tion would lead to the increase in intrafollicular pressure, resulting in ovulation. Comer (114,115) described smooth muscle in sow ovar¬ ian follicles, and Guttmacher and Guttmacher (202) con¬ firmed these observations and suggested that the smooth muscle was responsible for ovulation under the stimulation of the nerve fibers that they traced into the follicle wall. The significance of their study is that they caused contraction of sow follicle wall strips in vitro with low pH, barium chloride, and physostigmine sulfate, but caused relaxation with high pH, epinephrine, and atropine. They concluded that ovulation was contributed to by contraction of smooth muscle under autonomic control and that increased intra¬ follicular pressure resulting from increased arterial pressure was not sufficient to cause ovulation. Kraus (263) evaluated the roles of increased intrafollicular pressure, proteolytic enzymes and the presence of smooth muscle as factors in the mechanism of ovulation in the frog, hen, and rabbit. In the frog, pressure applied at the base induced rupture in the stigma area, but introduction of saline into the follicle was without effect. In the hen, application of external pressure induced rupture at unpredictable sites; however, late in follicular development, such pressure pro¬ duced rupture at the stigma even though the hens were dead. Saline injections into the follicles were ineffectual. In the rabbit, pressure induced rupture, whether it was applied externally or by intrafollicular injection of saline. Kraus (263) noted that, in the frog and hen, increased intrafollic¬ ular pressure may be an adequate explanation if combined with proteolytic changes in the follicle wall. In the rabbit, however, the pattern of discharge of follicular fluid argued against a sharp increase in intrafollicular pressure, but for an orderly progression of morphological change. Although smooth muscle had been reported in ovarian follicles by a number of workers, subsequent workers failed to confirm its presence in humans (429), swine (20), or rabbits (206). Claesson (107) attempted to resolve the di¬ vergent results obtained by histologists by utilizing polari¬ zation microscopy. He found perifollicular spindle-shaped cells resembling smooth muscle in the ovaries of cow, swine, rabbit, guinea pig, and rat. However, by using polarization microscopy he failed to find the high intrinsic birefringence characteristic of muscle, but did find the high form bire¬ fringence characteristic of connective tissue. The absolute objectivity of polarization microscopy completely negated

Mammalian Ovulation / all previous histological evidence, including Claesson’s own, of smooth muscle in the follicle wall and threw doubt on the reliability of any physiological data.

The Vascular System Direct microscopic observation of ovulation in rabbits was achieved by Walton and Hammond (495), who timed the process from coitus to rupture and confirmed the time reported earlier by Heape (209). They described the follicle as a broad-based convex structure, covered with a fine cap¬ illary network, protruding above the surface of the ovary. The first sign of impending ovulation was the appearance of the macula pellucida, which implied an obliteration of the blood flow to the apex, followed by further protrusion of the follicle. The macula pellucida became a pimple and ruptured, and the capillaries at the base of the pimple rup¬ tured before the pimple did. The rupture of the capillaries was interpreted to mean that thecal rupture preceded rupture of the surface. “The whole process of ovulation, the conges¬ tion and swelling of the follicle, the formation of a pimple at the surface and the gradual rupture and extrusion of con¬ tents strongly resembles the formation and rupture of a boil and, not unlikely, somewhat similar mechanisms are in¬ volved” (495). The investigators further noted that the liquor folliculi did not squirt from the follicle but flowed as the walls gradually collapsed. Kelly (248) confirmed with little variation the observations made by Walton and Hammond (495). Markee and Hinsey (310) confirmed the earlier ob¬ servations by Walton and Hammond (495) and Kelly (248) but noted that the mature follicle developed an avascular area surrounded by dilated vessels before there was any sign of a papilla. They also examined the rupture points and noted that they were uniformally oval shaped, with no ap¬ pearance of the tearing they assumed would be present if there had been an increase in intrafollicular pressure re¬ sulting either from smooth muscle contraction or from a rise in ovarian blood pressure. They thus concurred with Guttmacher and Guttmacher (202), who had failed to induce follicle rupture in Graafian follicles by injecting saline into ovarian arteries and raising the intraarterial pressure to 300 mm Hg. Despite suggestions that ovulation occurred with minimal intrafollicular pressure changes, contrary observations con¬ tinued to appear. Smith (444,445) noted that follicular fluid rose to 1 cm or less into a fine-tipped tube inserted into a Graafian follicle of an uninjected mature rabbit, but it rose to a height of 5 or 6 cm from the Graafian follicle of a pregnancy-urine-injected rabbit, suggesting that the latter fluid was under greater pressure. Hill et al. (214) applied the newly introduced cinemicrographic technique to the study of ovulation in the rabbit. These workers interpreted their observations as indicating that “follicular rupture is truly explosive in nature.” These observations supported the idea

449

that a tension was exerted on the intrafollicular contents and that rupture was a result of the pressure generated by the follicle wall. Although Schochet (428) had suggested that ovulation might be a result of enzymatic degradation of the follicle wall, and Thomson (466) had included this idea in what I call the multifactor hypothesis, little work was done on the enzymatic degradation of the follicle until Espey and Lipner (157) noted the effect of enzymes injected into the follicle.

Depolymerization of Mucopolysaccharide Because after 1947 it was generally accepted that smooth muscle was absent from the follicle wall or present in in¬ sufficient amounts to affect intrafollicular pressure (107), and Guttmacher and Guttmacher (202) had discounted the increase in intravascular pressure as a source of intrafollic¬ ular pressure, an alternative mechanism was needed to ex¬ plain the assumed increase in intrafollicular pressure. Zachariae (106,226,518-520) suggested that the mucopoly¬ saccharides synthesized by the granulosa cells and secreted into the follicle antrum were subjected to depolymerization to smaller molecular species, inducing an increase in the number of osmotically active molecules, and thus increased the volume of fluid in the follicle and the intrafollicular pressure. Nine hours after mating, rabbits were laparotomized, the ovaries were located, and Evans blue was admin¬ istered. Graafian follicles of unmated rabbits showed little accumulation of Evans blue at 20 min and slightly more at 30 min. The Graafian follicles of mated rabbits began to accumulate the dye within 5 min and were intensely stained by 20 min. The permeability of the follicle wall had in¬ creased, and, because Evans blue is bound to plasma pro¬ teins, it was unlikely that the mucopolysaccharide depoly¬ merization hypothesis could explain the increase in follicle volume or rupture as being caused by increased pressure. That ovulation might be a result of smooth muscle con¬ traction was again suggested by Lipner and Maxwell (295), who found that rabbit Graafian follicles autotransplanted to the anterior chamber of the eye underwent changes in shape within minutes after administration of pregnancy urine.

Neural Control of Ovulation A role for neural control of ovulation has periodically been suggested, beginning with the recognition that sym¬ pathetic nerves entered the ovary at the hilus along with the vacular supply (177,218,487). The nerves entering the ovary either accompanied blood vessels or went directly to the follicles (417). Opinions on the subsequent distribution of the nerves divided anatomists into three schools of thought: (a) those believing the nerves penetrated all layers of the follicle wall (63,182,210,417,507); (b) those believing only the two outermost layers of the follicle wall received neural

450

/

Chapter 12

innervation (1,202); and (c) those believing only the vessels in the follicle wall were innervated (129,308,415). Physiological studies initiated by von Winiwarter and Sainmont (506), who stimulated follicles in situ either chem¬ ically or electrically, were extended by Thomson (466), who noted that epinephrine was stimulatory in vitro, continued by Guttmacher and Guttmacher (202), who found that epi¬ nephrine was inhibitory and that acetylcholine was excita¬ tory and supported by the demonstration of norepinephrine in ovarian tissue (164). The whole significance of the neural innervation was challenged by the demonstration by Hinsey and Markee (217) that rabbits in which the vagal, thora¬ columbar, and sacral innervations to the ovary were de¬ stroyed could still ovulate in response to stimulation with pregnancy urine. Modem study of ovulation has made use of all the tools of anatomy, physiology, pharmacology, and biochemistry to analyze the mechanism. Although there are still some questions to be resolved, a more coherent de¬ scription of ovulation has now become possible.

THE STRUCTURE OF THE OVARIAN FOLLICLE

General Features of Follicular Morphology The structure of the ovarian follicle has been extensively reviewed by Nagel (344), Klebs (250), and most recently Mossman and Duke (334). Both structure and function have been reviewed by Bjersing (48), Erickson et al. (147), and Guraya (201). The adult ovary of mammals is invested with an epithelial membrane, the germinal epithelium, overlying a tough con¬ nective membrane, the tunica albuginea. Immediately be¬ neath the tunica albuginea is the ovarian cortex, in which are located the follicles in various stages of maturation or degeneration. The primordial follicles are surrounded by a single layer of granulosa cells; with maturation, the granulosa cells in¬ crease in number and begin to form layers, which appear stratified or pseudostratified (49,293). As the granulosa cells increase in number, an encapsulating sheath, the theca, com¬ posed of the surrounding stroma, delineates the developing follicle, separating it from adjacent primordial follicles. The theca becomes differentiated, as antrum formation begins, into a well-vascularized theca interna consisting of several layers of rounded steroid-secreting cells and an equally wellvascularized theca externa composed of spindle-shaped cells (371,372). The outer wall of the preovulatory follicle is generally described as being composed of six layers. The most su¬ perficial is the surface germinal epithelium, which rests on a basal lamina, which in turn merges with the tunica al¬ buginea; these three elements constitute the serosa of the ovary, which is comparable to the serosae of other organs suspended in the abdominal cavity. The wall of the follicle

abutting the tunica albuginea is composed of the theca ex¬ terna, then the theca interna, which rests on the lamina propria, which excludes all direct vascular supply to the innermost layer, the membrana granulosa.

Detailed Features of Follicular Morphology Germinal Epithelium The comparative aspects of the structure of the ovary have been examined by Mossman and Duke (334), and the following discussion is largely based on their evaluation of the literature. The ovary is attached to the body wall by the mesovarium and, except for the line of junction of the peritoneum, is completely enveloped. It is at this junction, the hilus, that blood vessels and nerves enter the ovary. The serosal layer enveloping the ovary consists, as it does for most mam¬ malian organs, of a simple squamous epithelium overlying a fibrous connective tissue layer. In the course of embryogenesis, the epithelial cells overlying the ovary undergo a transformation from squamous to low columnar epithelium lying upon a basement membrane, which separates the ep¬ ithelium from the connective tissue layer. The columnar nature of the epithelium persists especially in the region of the hilus, but ranges from squamous to low columnar in other areas of the ovary. In some species of small mammals, the basement membrane is absent, and there may not be a clear distinction between the underlying cells and the surface epithelium. Mossman and Duke (334) conclude that the serosa of the mammalian ovary is relatively embryonic, but that “it actually bears a much more intimate morphological and physiological relation to the organ as a whole than does that of other organs. . . . Even the adult ovary retains much of its embryonic character, its epithelium having a more or less continuous and formative relationship with it. For this reason, the epithelial covering of the ovary is regarded as different from the rest of the coelom, and in view of its function in gonadal development should be called surface germinal epithelium (epithelium germinale superficiale).” The relationship of the superficial germinal epithelium to cortical development of the ovary was first described by Pfluger (400) and in ovaries of fetal and neonatal dogs and humans (337). The superficial epithelium was termed ger¬ minal epithelium by Waldeyer (487). Ultrastructural ex¬ amination of germinal epithelium of the mouse ovarian sur¬ face by Wischnitzer (508) revealed an epithelium similar to that described by light microscopy, with the addition of microvillus projections and the usual cell organelles. Espey (148,150) described the germinal epithelium of the rabbit ovary but was more concerned with the underlying struc¬ tures. A later study noted the presence of electron-dense particles in the germinal epithelium (410). Byskov (86) also studied the surface germinal epithelium in the mouse ovary but only confirmed the description by Wischnitzer (508).

Mammalian Ovulation

In 1971, Motta et al. (335) examined the ovarian surface epithelium of guinea pigs, mice, rats, rabbits, and humans and correlated the changes in the epithelium with the fol¬ licular events preceding ovulation. The observations were similar in all the species examined. The cells were hemi¬ spheric, with the convexity directed outward and the surface covered with microvilli. The cells tended to flatten and become squamous only on the apices of enlarging pre¬ ovulatory follicles, with short evaginations appearing on their surfaces, suggesting increased cellular activity. The surface epithelium disappears shortly before follicular rup¬ ture. The proliferative activity of the surface epithelium is minimal prior to ovulation and shows a resurgence adjacent to the stigma immediately after ovulation (375). The role of the surface germinal epithelium in rupture of the follicle was again evaluated in a series of studies by Bjersing and Cajander (49-51), Cajander (87), and Cajander and Bjersing (88,89). They hypothesized that, with time, the surface germinal epithelium undergoes a series of trans¬ formations that provide the enzyme(s) that digest the surface layers of the follicle and induce rupture. The release of LH was the assumed initiator; however, this hypothesis fails to explain the changes in vascularity and neural activity or the series of biochemical changes (steroidal and nonsteroidal) that occur during the periovulatory period.

Tunica Albuginea The serosal cover of the ovary consists of the surface germinal epithelium, a thin, almost cell-free layer of amor¬ phous translucent material in which are embedded collagen bundles, and a relatively thin layer of fibroblasts surrounded by interwoven compact collagen fibers. It is this last area that is usually referred to as the tunica albuginea. Mossman and Duke (334) note that in small mammals (shrews and mice) the tunica albuginea is absent. In rats it appears as an indistinct subepithelial fibrous layer, but in the porcupine it is a distinct subepithelial fibrous capsule and should be considered the analog of the testicular tunica albuginea. The surface epithelium of the ovary in young women is under¬ layered by a thin subepithelial fibrous capsule with little transition into the ovarian cortex.

Theca Externa The outermost coat of the preovulatory follicle subjacent to the tunica albuginea is composed of several layers (two to five or seven) of fusiform cells and an extensive capillary network. The fusiform cells are of two types. One of these is designated as stromal or fibroblast cells and is charac¬ terized by an endoplasmic reticulum with cistemae rami¬ fying through the cytoplasm, which contains occasional mi¬ tochondria and free ribosomal particles. The second type of fusiform cell is ultrastructurally similar to smooth muscle. In these cells are bundles of thin filaments, numerous free

/

451

ribosomes, lipid droplets, glycogen granules, mitochondria with long cristae, and a Golgi vesicular complex. The plasma membrane contains caveolae intracellulares (invaginations) and occasionally myoneural junctions. The muscle cells are present in fascicles, in small groups, or as isolated cells. They are interspersed among the fibroblasts and form in¬ complete layers in the thecae extemae of developing, ma¬ ture, and atretic follicles and are present around corpora lutea and interstitial cells. Smooth muscle. Smooth muscle has been described in the theca externa in most mammals: rat (18,181,369,377), sheep (370), monkey (360,377), mouse (18,181,336), cat (18,68,181,360), guinea pig (68), rabbit (18,53,68,360), human (360), hamster (318,389,391), gerbil (318), and cow (490). Although the smooth muscle cells may be under neural control, cellular junctions among the perifollicular smooth-muscle cells may allow direct communication and sequential response (93). Collagen. The theca externa invariably contains collagen fibers occurring in bands and interspersed among the fibro¬ blasts and smooth muscle cells. The mechanisms determin¬ ing the rupture of the Graafian follicle primarily involve the degradation of the follicle wall (152,292). The primary com¬ ponent responsible for the wall’s tensile strength is its col¬ lagen content (149). It is therefore essential that some dis¬ cussion of collagen precede any review of the studies regulating its degradation. The following discussion is based on the reviews by Miller (320), Eyre (167), Jackson (222), and Fessler and Fessler (170). The word collagen is now used as a generic term de¬ scribing a population of distinct, but chemically related, macromolecular species (320). Table 1 lists the known ver¬ tebrate collagens and divides them into types based on their physicochemical properties. The properties shared by the 11 currently recognized collagens are that they (a) contain sizable domains in which the collagen helical fold is present and (b) contribute structurally to the formation of extracel¬ lular supramolecular aggregates. Miller (320) has divided the collagens into three molecular groups. Group 1 mole¬ cules are those collagens that give rise to fibers; Group 2 molecules are those composed of very large aggregates that give rise to the open mesh-like arrangements of basement membranes. The organization of the Group 3 molecules is currently undescribed and not of interest in the context of collagens found in the ovarian follicle wall. Collagen is a ubiquitous protein component of connective tissues. A generalized view of such tissues reveals them to be composed of a fibrous meshwork of collagen and elastin embedded in an amorphous polyionic matrix of proteogly¬ cans and glycoproteins. Interspersed among the fibers are the cells giving origin to these fibrous proteins. The array of connective tissues in vertebrates is large, and the amounts of collagen in these tissues are variable. Final collagen as it appears in tissues consists of three alpha polypeptide chains. Each chain consists of a central collagen fold with a short telopeptide chain at each end.

452

/

Chapter 12

TABLE 1. Properties of selected group I and group II collagen moleculesa

Collagen

Chain molecular weight

Type 1

95,000

Type II Type III Type IV

95,000 95,000-100,000 170,000-185,000

Type V

130,000-200,000

Molecular species MOkMI)] Ml)]3 Mll)]3 Mlll)]3 [ai(IV)]2[a2(IV)] [«i(IV)]3 [a2(IV)]3 MV)]2[a2(V)] MV)]3 MV)][a2(V)][a3(V)]

Form of aggregates Fibers Fibers-fibrils Reticular networks Open mesh-like aggregates of basement membranes Aggregates of unknown structure in pericellular and perifibrillar genes

aModified from Miller (320).

Roman numerals indicate the genetic types of collagen chain within a species. Types II, III, and probably IV are com¬ posed of three identical chains. Type I is a heterotrimer composed of two alpha-1 and one alpha-2 [(a!(I))2(a2(I))] chains. Type IV is found in basement membranes and is the best-characterized collagen type. It is the largest and has many 3-a hydroxyproline residues. The different collagen types tend to be distributed in particular tissues. Thus chon¬ drocytes are a primary source of Type II, which is largely present in hyaline cartilage. Type I is formed by osteocytes and corneal and tendon fibroblasts. Smooth-muscle cells are capable of synthesizing all of the types of collagen. Blood¬ vessel smooth-muscle cells in culture express both Type I and Type III collagens in variable amounts. Structures that resist deformation, such as bone, generally contain Type I collagen. Elastic connective tissues contain Types I and III, whereas Type IV is the collagen found between the under¬ lying stroma and epithelium. Type II is restricted to struc¬ tures that resist deformation, such as the cartilagenous tis¬ sues and vitreous humor of the eye. Type I and III collagens have been identified by immunohistochemical methods in the theca of the Graafian follicle (382) and appear as bundles of fibers distributed among the smooth-muscle cells and fibroblasts, whereas the lamina propria between the theca interna and the granulosa cells contains Type IV collagen (31,382). Martin and Miller-Walker (312) used a solution of sodium dodecyl sulfate incubated with hamster ovaries to strip away the overlying germinal epithelium and examine the distri¬ bution of collagen fibers in the follicle wall. They observed bands of parallel fibers arranged in layers beginning at the base of the follicle and extending into its apex, where they fan out into a mesh work covering the apex. Although similar studies have not been performed on the follicles of other species, ultrastructural study has indicated the presence of collagen fibers. Continued presence of collagen at ovulation is described in hamsters (312) and in rats (477), but dis¬ solution of collagen has been reported in rat (120,328, 362,385,412), human (179,359), and rabbit (148).

“Collagen degradation can follow two pathways. Its triple helical configuration is resistant to proteolysis. Collagenase, a specific metalloprotease, completely splits the molecule, producing fragments that are three-fourths and one-fourth of the molecule. At 37°C, these fragments unwind, giving rise to three separate gelatin chains, which are rapidly at¬ tacked by proteases. Extracellular serine proteases may at¬ tack the residual collagen molecule at the telopeptide por¬ tion, splitting the cross-linking and at 37°C denaturing the soluble telopeptide-free collagen to gelatin chains. A second pathway involves granulocytic or fibroblastic phagocytosis and intracellular degradation (510). The proteoglycan ground substance surrounding the collagen fibrils is degraded by serine proteases and elastase and these probably contribute to the complete proteolysis of collagenase-degraded colla¬ gen (343). The sources of collagenase. Granulocytic proteases. The human polymorphonuclear leukocyte (granulocyte) contains in its granules (a) neutral proteases that exhibit collagenase activity associated with a metalloprotease, (b) serine pro¬ teases having elastase and low-level collagenase activity, and (c) cathepsin G (chymotrypsin-like cationic protein). The collagenase is strongly bound to plasma a2-macroglobulin and less well bound to plasma P,-anticollagenase. The elastase is strongly bound to plasma a,-antitrypsin and mucus inhibitor I and weakly bound to a2-macroglobulin. The secretion of neutral proteases is low except during phagocytosis, and endogenous protease inhibitors normally keep the low levels in check, but local high concentrations of the enzymes may be achieved when the excess saturates the local inhibitory concentrations (354) during inflamma¬ tory reactions. Tissue collagenase. In addition to granulocytes, a wide variety of cells are capable of producing collagenase; how¬ ever, with the exception of the granulocyte, there is little evidence of intracellular storage of active collagenase. Col¬ lagenase production is dependent on induction of DNAmediated synthesis rather than on cellular release of pre¬ formed enzyme. Collagenase may be synthesized and stored

Mammalian Ovulation

in an inactive form, or both the enzyme and an inhibitor may be present simultaneously; their concurrent release would prevent the appearance of an active enzyme (511). Murphy and Sellers (343) hypothesize that all connective tissues synthesize collagenase inhibitors that complex with collagenase and that the resulting latent enzyme complex is, in turn, bound to collagen. In its latent form, collagenase is not subject to a2-macroglobulin inactivation. Latent col¬ lagenase may be a proenzyme requiring enzymatic activa¬ tion. A number of endogenous proteinases have been dem¬ onstrated to activate the latent collagenase by cleavage of a small peptide. The endogenous activator may also be present in a latent form requiring plasmin, kallikrein, or lysosomal proteinases such as cathepsin B and trypsin for activation (343). Plasmin is a very effective activator of latent collagenase; however, it is present in body fluids in the inactive plasminogen form and requires activation by plasminogen activator or streptokinase. Rheumatoid syno¬ vial cells secrete both latent collagenase and plasminogen activator and, in the presence of the local inflammation and large amounts of plasminogen, a situation adequate for ac¬ tivation of collagenase is created (501). Latent collagenase can also be activated by thiol-binding reagents such as aaminophenylmercuric acetate, A-ethylmaleimide, and pchloromercuribenzoate. The collagenase activity is as great with these compounds as with trypsin, and no synergism between trypsin and thiol-binding reagents is found (343).

Theca Interna Erickson et al. (147) elegantly summarized the literature on the structure and function of the interstitial cells of the ovary, and detailed references can be found in their review. They note that there are four varieties of interstitial cells. Primary interstitial cells constitute a transient population of endocrine cells contained in the medullary compartment of the developing ovary. These cells lack the ability to syn¬ thesize steroids de novo, but they convert pregnenolone sulfate to dehydroepiandrosterone and androstenedione (in humans). Theca interstitial cells are perhaps the most im¬ portant group of interstitial cells. These cells are found in the theca interna of the developing follicle. They are derived from stromal cells; cytodifferentiation occurs concurrent with their invasion by arterioles and the formation of a capillary wreath. The cells acquire 3(3-hydroxysteroid dehydrogenase and undergo the subsequent cytodifferentiation that char¬ acterizes the true theca interstitial cell. On the ultramicroscopic level, there appear to be at least two types of thecaintema cells, light and dark cells; both have the ultrastructure of steroidogenic cells. The third variety of interstitial cells are those found around atretic follicles. These cells are in¬ nervated by adrenergic nerves and, in culture, respond to catecholamine stimulation by increasing their production of androgen. Finally, the fourth variety of interstitial cells are those found in the hilus. These cells resemble the Leydig

/

453

cells of the testis and respond to LH stimulation by increas¬ ing their testosterone synthesis and secretion (265). Lamina Propria The theca interna and its vascular supply are separated from the membrana granulosa by the lamina propria. This amorphous boundary is composed of laminin, Type IV col¬ lagen, fibronectin, and heparan sulfate proteoglycan. In¬ duction of ovulation with human chorionic gonadotropin (hCG) causes fragmentation of the lamina and invasion by thecal cells and capillaries (31,382), probably under the influence of induced angiogenic factor (260,306). Membrana Granulosa The parietal membrana granulosa of the preovulatory fol¬ licle in the rabbit is described as a stratified epithelium or a pseudostratified epithelium four to five cells thick (49,293). The cells close to the lamina propria appear columnar, whereas those lining the antrum appear polyhedral. Many of the luminal cells have long slender projections extending to the lamina propria (49,293). The membrana granulosa in the human preovulatory fol¬ licle is six to 12 cells thick but appears similar to that in the rabbit. The cells of the basal layer are columnar, and those of the other layers are polyhedral. After the LH surge, the antral cells become disorganized, and the cells and their nuclei increase in size (60). The ultrastructural appearance of the cells of the membrana granulosa is consistent with a steroid secretory function. They contain a granular endo¬ plasmic reticulum, and at estrus 3(3 and 17[3-hydroxysteroid dehydrogenase activity is high, at the time that estrogen secretion by the follicle is maximal (48,326). Steroid biosynthesis is generally associated with secretory function of the cells of the membrana granulosa; however, other nonsteroidal factors are synthesized and secreted by these cells. Among these are the prostaglandins, inhibin, a 32K molecular weight protein that inhibits basal secretion of FSH (102,146) plasminogen activator (90,351) and pep¬ tide growth regulators (207,208). The granulosa cells also secrete the components of the extracellular matrix. Fibronectin is secreted by the granulosa cells before FSH induces the cytodifferentiation that inhibits its further production (443). The guinea pig ovarian follicle incorporates 35S04 into glycosaminoglycans, which also contribute to the extracellular matrix composing the early follicular fluid (110). Three varieties of granulosa cells occupy the area beneath the lamina propria in rat preovulatory follicles. Lying on the lamina propria is a parietal layer of columnar cells joined at their lateral margins by gap junctions (Type I). Adjacent to this basal layer and extending to the antrum are polygonal cells joined to each other as well as to the apices of the parietal cells by gap junctions (Type II). Antral cells floating in follicular fluid free of Contact with subjacent cells or with

454

/

Chapter 12

each other appear shortly before follicular rupture (Type III) (175). The Type I and Type II cells on the morning of proestrus contain small mitochondria with lamellar cristae, moderately developed Golgi apparatus, inconspicuous lysosomes, and only small amounts of lipid inclusions. Late in proestrus, Type I cells have only short segments of rough endoplasmic reticulum (RER), but the smooth endoplasmic reticulum (SER) shows profuse cisternal dilations. Mito¬ chondria are increased in size and number and appear spher¬ ical; the cristae are tubular. Lipid inclusions are plentiful, and Golgi cistemae are enlarged. The ultrastructural ap¬ pearance of the membrana granulosa has been similarly described in several species, including the rat (55,175), mouse (86), rabbit (148), cow (309,406), and human (180). Existence of cell-cell contacts was reported by Bjorkman (55) and Byskov (86); however, their significance was un¬ appreciated until Espey and Stutts (160) and Merk et al. (319) identified them as gap junctions between the granulosa cells and annular gap junctions contained within the cyto¬ plasm. Espey and Stutts (160) suggested the latter might be a mechanism for exchange of cytoplasm between the cells; their internalization might be a mechanism for decreased intercellular adhesion. The structure of the gap junction in different organisms and tissues has been reviewed by Larsen (271); Peracchia (393,394) and Loewenstein (299) have re¬ viewed structure, mechanism of formation, control of per¬ meability, number of particles, and adhesion. Uncoupling and decreased numbers of gap junction particles may result from decreased intracellular Ca2+ concentration, intracel¬ lular acidification (394), or decreased intracellular concen¬ trations of cAMP (299). Both follicle stimulating hormones (FSHs) and LH increase intracellular concentrations of cAMP; therefore the effects of these hormones should be to increase the numbers of gap junctions among the granulosa cells. Albertini and Anderson (14) reported that gap junctions appear among granulosa cells concurrently with the ap¬ pearance of the antrum (23). However, Fletcher (172,173) noted them among granulosa cells in 3-day-old rats, and Burghardt and Matheson (78) found them present in preantral follicles and probably present from the earliest appear¬ ance of granulosa cells. Albertini and Anderson (14) found that gap junctions of the granulosa cells increase in size as the follicles grow. Using tracers, Albertini et al. (15) dem¬ onstrated that annular gap junctions arise by invagination of the cell surface. Larsen and Tung (272) and Larsen et al. (273,274) described the probable mechanism for inter¬ nalization of gap junctions through an endocytotic process. The cAMP formed in rat granulosa cells in response to FSH exposure is transferred across heterologous gap junc¬ tions to co-cultured mouse myocardial cells, which manifest the effect by an increased beat (277). Furthermore, addition of FSH to the medium of cultures of rat granulosa cells causes the cells to change from a flattened epithelioid shape to a semispherical form but with retention of cytoplasmic processes in contact with adjacent cells and with the sub¬ strate (278). In in vivo studies, Burghardt and Matheson (78) found that FSH amplifies the sequence of gap junction

growth and turnover in young follicles and increases inter¬ nalization in mature follicles. Prolonged exposure of cul¬ tured granulosa cells to FSH increases membrane micro¬ viscosity (454) and probably reduces the lateral mobility of granules in the cell surface, thus contributing to the inter¬ nalization of gap junctions. Although Bjersing and Cajander (52) and Coons and Es¬ pey (113) had noted a decrease in the number of gap junc¬ tions after hCG and as the time of ovulation approaches, Larsen et al. (274), in a quantitative analysis of the gap junction, found that hCG induces an increase in the pro¬ portion of small surface gap junctions in the granulosa cells and a marked decrease in the proportion of large surface gap junctions. They also found a decrease in the amount of gap junction membranes in the granulosa cells. Burghardt and Matheson (78) hypothesize that synthesis and turnover °f gap junctions are amplified by mechanisms that increase ihtracellular cAMP and that the ratio of surface to inter¬ nalized gap junctions is dependent on the state of maturation of the follicle. FSH promotes formation of surface gap junc¬ tions in early stages of follicle growth and turnover of gap junctions in later stages of follicle growth, and LH takes over this role in the later stages (76-78). The significance of the ultrastructural organization of the membrane granulosa resides in the observation that the cells composing it are heterogeneous; those located near the basal lamina possess larger numbers of LH receptors than those closer to the antrum (21,241) and, by cell-cell communi¬ cation, permit uniform responses by cells separated from the vascular system (186). Prolactin receptors are more prev¬ alent in the antral than in the mural (parietal) granulosa cells (140), and the enzyme populations are a function of location in the membrane granulosa (521,522).

THE NEURAL INNERVATION OF THE OVARY Neural Innervation of Ovary and Follicles The gross anatomy of the nerve supply to the ovary was described in 1867 by Frankenhauser (177). The nerves orig¬ inate at spinal segments T10 and T„ (268,502), give rise to an aortorenal plexus, and follow the ovarian artery as a nerve plexus into the ovary at the hilus (345). Sensory fibers originate at spinal segment Ti0 and accompany the sym¬ pathetic fibers. The parasympathetic innervation is derived from the vagus (212,213,286). Spinal segments S-2 to S-4 may also contribute to the parasympathetic innervation (323); however, denervation studies suggest that these, if they exist, are inconsequential. In the rat the parasympathetic innervation is primarily vagal in origin (19,71,73,74,171,212). Furthermore, much of the innervation is probably composed of visceral afferent fibers, since in the rabbit only 10% of the abdominal vagal fibers are motor neurons (165). Early studies of the microscopic distribution of the auto¬ nomic nerve distribution utilized silver impregnation tech¬ niques, which were subject to artifact because of the ten¬ dency of the method to stain reticular and elastic fibers as

Mammalian Ovulation

TABLE 2. Comparison of adrenergic innervation by fluorescence microscopy with concentration of norepinephrine in mammalian ovariesa Innervation Rich Cow Sheep Cat Guinea pig Left Right Intermediary Human Pig Dogb Rat Opossumb Sparse Rabbit Mouse Hedgehog Syrian hamster Squirrel monkey Rhesus monkey

Concentrations (|xg/g)

Reference

1.96 ± 0.19

444

4.93 2.29 2.32 2.16

1.30 0.35 0.30 0.20

424 449 267 267

0.86 ± 0.30 0.57 0.25 ± 0.04

378 224 449 449 449 449 449

± ± ± ±

0.50 ± 0.03

0.20 0.22 0.63 0.71 0.41 0.17 0.35

± ± ± ± ±

0.02 0.02 0.11 0.12 0.09

± 0.06

442 58 449 449 449 224 380

aModified from Stefenson et al. (449). ^Qualitative evaluation.

well as nerve fibers (32,71). Development of the histochemical fluorescence method for biogenic amines (168,169) marks the beginning of the new era in the study of the adrenergic innervation of the ovary. Concurrently, refine¬ ments in the histochemical methods for demonstrating acetylcholinesterase and distinguishing it from nonspecific cholinesterases have led to more precise descriptions of the parasympathetic innervation of the ovary (67,239). Autonomic innervation of the ovary has been studied extensively in a number of species for the development of innervation (234) and for the distribution of nerve fibers in the cortex and in and around the follicles (69,70,378,379,424). The fluorescence has been identified as being primarily nor¬ epinephrine (378). Innervation of the cortical stroma and the follicles, as well as the nerve distribution in the theca externa and theca interna, has been found in all species examined. A distinguishing characteristic is the density of innervation, which varies from slight to intense. The ad¬ renergic innervation based on qualitative histochemical evaluation is compared with concentrations of norepineph¬ rine in Table 2 (in the case of sheep, dog, and opossum, only the qualitative histochemical evaluation is indicated) (449). The presence of specific acetylcholinesterase has been demonstrated histochemically, with concentration arrived at on the basis of qualitative evaluation. In human, dog, rat, and hamster, the appearance of the cholinergic fibers is similar, with a dense network of these fibers associated with blood vessels and follicles. The follicles of the human ovary

/

455

have an especially dense innervation. The incidence of cho¬ linergic fibers is less than that of adrenergic fibers. The rabbit ovary has a sparse and random distribution of cho¬ linergic fibers (449). In strips prepared from bovine ovarian follicles, the smooth muscle cells lie in the theca externa in groups parallel to the surface, with distances of 30 to 100 nm between them. Bundles of nonmyelinated nerves in the stroma send branches to the smooth muscle cells as well as to the smooth muscle of the blood vessels. The naked nerve terminals lie in close proximity to the smooth-muscle cells, as close as 50 nm. In the absence of specialized junctions, this is a distance short enough for diffusion to provide innervation. These nerve terminals contain dense-cored synaptic vesicles and are presumed to be adrenergic nerves. Also present are nerve terminals containing empty synaptic vesicles, and because these tend to correspond to the nerve distribution of cho¬ linesterase-containing nerves, they are presumed to be cho¬ linergic nerves (492). Similarly, the presence of both ad¬ renergic and cholinergic nerves in the ovarian follicle wall has been noted in humans (378,379,433), rats, guinea pigs, and rabbits (60,74,232), and cats (424). The concentration of norepinephrine in the ovary is a function of its innervation by the ovarian plexus, because crushing of the nerves on the guinea pig ovarian artery results, after 8 days, in a decrease in the concentration of norepinephrine in the treated ovary, whereas sectioning the hypogastric nerve has no ef¬ fect on the norepinephrine concentration (267).

Physiological Studies Noncontractile Effects of Autonomic Innervation Adrenergic innervation of the ovary has been associated with follicular growth (64,119), cyclicity (286,459), pu¬ berty (6,7,333), innervation of the guinea-pig ovarian in¬ terstitial gland (457), progesterone synthesis (3,105,240, 348,409,525), compensatory ovarian hypertrophy (73, 193), and ovulation (96,124,176,217,223,228,235,302, 497-500) (Table 3). Prostaglandins, Oxytocin, and Ovarian Contractility In addition to the autonomic transmitters, a number of substances also induce contractions of the ovary in vivo and in vitro or of follicle strips in vitro. Prostaglandin F2c[ in¬ duces contractions and prostaglandin E2 (PGE2) relaxation (116,117,130,187-189,303,450,483). Oxytocin induces ovarian contractions throughout the estrous cycle (419); however, Sterin-Borda et al. (450) observed more marked contractions during the periovulatory period (late proestrus), with a greater activity in the left ovary than in the right. Serotonin (5-hydroxytryptamine) causes contractions of the preovulatory hamster ovary (464). The complexity of neurohumoral-induced effects is further exaggerated by dem¬ onstrations of several [enkephalin, substance P, vasoactive intestinal peptide, neurotensin, somatostatin, gastric-releas-

456

/

Chapter 12

TABLE 3. Neural regulation of noncontractile mechanisms in the ovary Species Rat Rat Mouse

Rat

Rat Mouse

Treatment Surgical sympathectomy Ovarian autografts Denervation of ovary

Reserpine to immature rats treated with PMSG/hCG Ovarian autografts Denervation of ovary

Mouse

Norepinephrine to hypophysectomized mice

Mouse

Unilateral denervation of PMSG/hCG

Mouse Mouse Guinea pig

6-Hydroxydopamine 6-Hydroxydopamine None

Rat

Ovarian autografts

Rat

None

Rat

Denervation

Rat

Vagotomy and 6-hydroxydopamine

Rat

Adrenalectomized hyphophysectomized electrical stimulation of areas of brain p-adrenergic stimulation Ovarian denervation Ovarian denervation in pubescent rats Catecholamines in ovarian bursa

Rat Rat Rat

Rat

Response

Reference

Normal estrous cyclicity

459

Ovulation

124

Delayed vaginal opening, failure of normal estrous cyclicity, failure of follicle maturation, failure of corpus luteal formation Inhibition of ovulation effect partially reversed with pargyline

197

176

Normal ovulation Delayed vaginal opening, - decreased ovarian weight, erratic estrous cycles, decreased follicle maturation, failure of corpus luteal formation Slight regression of interstitial elements, increased numbers of type 3b and 4 follicles, decreased numbers of atretic follicles Devervated ovary had reduced numbers of maturing follicles; after gonadotropins effects were exacerbated Decreased litter size No effect on fertility Interstitial gland of ovary innervated by adrenergic nerves No correlation between density of innervation and plasma levels of progesterone and estradiol Interstitial gland of ovary innervated by autonomic nerves Decreased 33-hydroxysteroid dehydrogenase activity during pregnancy only Disrupts estrous cycles, interrupts estrous cycles, decreases compensatory ovary hypertrophy Increased secretion of progesterone and estradiol

133 198

199

64

96 228 457

105

276 72

73

244 245

Secretion of progesterone

525

Acute decreased secretion of progesterone Depletion of ovarian norepinephrine, no effect on puberty or first ovulation

6,7,356 7

Increased numbers of ovulation

427

Mammalian Ovulation / ing peptide, bombesin, neuropeptide Y (451)] different pep¬ tides and indolamine (serotonin) associated with autonomic nerves (83). Neural Innervation and Ovulation A frequently examined issue has been the role of the neural innervation in ovulation. In acute experiments, in which hormonal treatment to the animal is followed by in vitro perfusion of the ovary, ovulation occurs. Examples of such experiments are those by Wallach et al. (488). In such studies the nerve terminals may still provide the control of smooth-muscle contractility necessary for ovulation. This criticism has led to various attempts to demonstrate the independence of the ovulatory mechanism from the neural innervation demonstrated anatomically. Autotransplantation of the rat ovary into subcutaneous sites has no effect on the incidence of ovulation (124); however, Jacobowitz and Laties (223) noted that ovarian autotransplants into the anterior chamber of the cat eye were reinnervated by adrenergic fibers with the revascularization. Because Deanesley did not examine the transplants for a neural reinnervation, it is possible that the functional ovarian tissue ovulated because it was reinnervated. Spinal cord transection in the absence of abdominal vagotomy is an inadequate experimental determination of the role of neural innervation in follicular maturation and ovulation (184). Weiner et al. (497) utilized the technique of stripping the neural plexus around the rabbit ovarian artery to denervate the ovary. The ovaries lost all evidence of histofluorescence, but ovarian size and follicle number were unaffected. Rab¬ bits with one denervated ovary ovulate equally frequently on the denervated and control sides in response to hCG and with coital stimulation (498,499). The denervated ovary shows spontaneous contractile activity and responds to nor¬ epinephrine with a sharp increase in tone and rapid small contractions; exposure to the a-adrenergic blocking drug phenoxybenzamine totally inhibited the contractile activity, as did exposure to isoproterenol, a (3-adrenergic agonist, but propranolol, a (3-adrenergic blocking drug, enhanced the activity (500). Retention of, and even increased, sen¬ sitivity to agonists is a common characteristic of denervated smooth muscle (203), and because most receptor structures are retained in the membrane, inhibition of contractility by a-receptor antagonists is demonstrable (500). Demonstration that 6-hydroxydopamine causes selective degeneration of adrenergic terminals, i.e., a chemical sym¬ pathectomy (471), resulted in its application to ovarian func¬ tion. Administered to mice, 6-hydroxydopamine had no ef¬ fect on the estrous cycle or ovulation (96,228), nor did it affect these parameters of ovarian function when adminis¬ tered in rats (302). Autonomic Innervation and Ovarian Contractility Another approach to studying the role of autonomic in¬ nervation to the ovary has been the utilization of cholino¬

457

mimetic and sympathomimetic agonists and antagonists ap¬ plied in in vitro organ bath systems. This technique, with many modifications and refinements (427), has become stan¬ dard for studying the effects of various reagents on con¬ tractility of the follicle wall. Contractions of ovarian follicles were first reported in autotransplanted follicles in the an¬ terior chamber of the rabbit eye (295) in response to hCG stimulation. Guttmacher and Guttmacher (202) had dem¬ onstrated that ovarian strips could be induced to contract, but spontaneous contractions of ovaries of cats suspended in an organ bath were observed by Rocerto et al. (420). They also noted the enhanced contractile activity caused by norepinephrine and epinephrine, as well as the inhibition of norepinephrine-induced increased frequency of contractions caused by phenoxybenzamine. These observations have been confirmed in vivo in rabbits (482,483) and in monkeys (254,484). The in vitro method has been used to demonstrate the presence of spontaneous contractions in whole human ova¬ ries (116,383) and in human and canine ovarian strips (188.357.358.489) . The human ovarian strips showed min¬ imal spontaneous contractility, which, however, could be induced with cholinergic and adrenergic compounds (357.489) . In general, some cholinergic drugs (acetylcho¬ line, bethanechol, and neostigmine) enhanced contractility, some (methacholine and pilocarpine) exerted variable ef¬ fects, and atropine depressed response (126). Effects of cholinomimetic and sympathomimetic drugs on ovarian and follicle strip contractility have been studied most elegantly and systematically by the Swedish group (235,379,427,446,489^494). Both the cat and the guineapig ovary show an increase in tone and contractility when exposed in vitro to norepinephrine; however, the guineapig response appears only in the presence of a (3-receptor antagonist. The contractility induced with norepinephrine is abolished with a-receptor blockade. These effects were also examined in strips prepared from the protruding part of bovine or human follicles. The bovine follicle strip con¬ tracted in response to norepinephrine; contraction was blocked irreversibly by phenoxybenzamine and reversibly by piperoxan, both a-adrenergic receptor antagonists; however, the piperoxan is an a2-receptor antagonist and therefore con¬ traction is caused by competitive inhibition of norepineph¬ rine-induced contractions (490). The human follicle strip also contracted in response to norepinephrine; however, the addition of a (3-receptor antagonist intensified the contrac¬ tion (489). The order of potency for the catecholamines in the bovine follicle strip is norepinephrine > epinephrine > phenylephrine > isoproterenol (490). The contractile potency relationship of catecholamines, when tested on human follicle strips, is norepinephrine > epinephrine > phenylephrine > isoproterenol, indicating that an a-adrenergic receptor is involved in generating the response. Relaxation induced by the catecholamines was evaluated by induction of a tonic contraction with carbamylcholine in a-receptor-antagonized strips. The potency

458

/

Chapter 12

TABLE 4. Studies of spontaneous and stimulated

contractility of the ovary or follicle Species

Procedure

Rabbit Rabbit Rabbit Cat Human

In In In In In

Human Rabbit

In vitro In vitro

Monkey

In vivo In vitro

Rabbit Human

In vivo In vitro In vitro

Sheep

In vitro

vitro vivo vivo vitro vivo

Cat In vitro Guinea pig In vitro Cow In vitro Human In vitro Guinea pig In vitro Human

In vitro

Human

In vitro

Cow Rabbit

In In In Human In Guinea pig In Rat In Rat In

vitro vivo vitro vitro vitro vitro vitro

Rabbit

In vivo

Rabbit

In vivo

Cow

Treatment

Pharmacologic agents Transplanted/hCG Spontaneous Spontaneous Prostaglandins F2a and E2 Spontaneous Prostaglandins F2a and E2 Spontaneous Prostaglandins a-Adrenergic blockers 3-Adrenergic blockers Prostaglandins Electrical stimulation Neurotransmitters Prostaglandins Spontaneous contractions Catecholamines Prostaglandin F2a Spontaneous Cholinergic agonists Adrenergic agonists Adrenergic agonists Cholinergic agonists Spontaneous contractions Oxytocin Prostaglandin F2a Neurotransmitter Prostaglandins Adrenergic agonists Electrical stimulation Adrenergic agonists Cholinergic agonists Neurotransmitters Prostaglandins Oxytocin Spontaneous Prostaglandins Oxytocin Spontaneous

Denervated Spontaneous a-Adrenergic agonists In vitro Electrical stimulation Inhibitors In vitro Electrical stimulation

Reference 202 295 157 420 116 383 482 484

130 360 358 357 373

489

188

188 189 379 490 126 303 303 419 450

480-482, 512 500

494

rank of the test compounds was isoproterenol > norepi¬ nephrine > terbutaline > epinephrine. Relaxation of the follicle wall is mediated through (3-receptors. The relaxation is antagonized by propranolol but not by practolol (3,-an¬ tagonist), indicating the relaxation effect is probably caused by a (32-receptor (493). Both acetylcholine and carbamylcholine induce contrac¬ tions of the follicle strips, and after addition of atropine, contraction is elicited only at higher concentrations of the

agonists. These observations suggest that both the parasym¬ pathetic and the sympathetic divisions of the autonomic nervous system can induce a contractile response in the follicle (379). Dose-response curves generated in the pres¬ ence of acetylcholine and carbamylcholine alone or in the presence of atropine indicate that muscarinic receptors are probably on the smooth-muscle cells (492). Electrical stimulation of bovine follicle wall strips induces a frequency-dependent contraction that is abolished in the presence of bretylium, phentolamine, and reserpine, indi¬ cating that adrenergic nerves and a-receptors are involved in the response (494) (Table 4). A possible role for the adrenoreceptor agonists in ovu¬ lation has been examined in an in vitro perfusion system utilizing rabbit ovaries (427). Terbutaline, a 32-adrenore¬ ceptor agonist, increased the incidence of ovulations in ova¬ ries from hCG-treated rabbits. The mechanism for this effect may be dependent on 32-receptor activation of cAMP for¬ mation, with the latter, in turn, increasing progesterone secretion (vide supra). Blockade of a-receptors with phen¬ tolamine and phenoxybenzamine (a,-receptor antagonists) reduced the incidence of follicular rupture induced with hCG. Clonidine, an a-agonist, somewhat stimulated the ovulatory process, and piperoxan, an a2-antagonist, blocks follicular contractility. It thus appears that both a- and 3adrenoreceptor agonists enhance the response to gonadotro¬ pin-induced ovulation. The sites of their actions probably involve the follicular wall (a-adrenergic response) and an enhancement of hormonal action in the theca interna and membrana granulosa (427). Introduction of norepinephrine, terbutaline, and 4-aminopyridine into the bursa around the ovary increased the number of ovulations in pregnant-mare-serum-gonadotropin (PMSG)-primed LH-triggered immature rats (235). In this study, phentolamine partially blocked the norepinephrineenhanced response and decreased the number of ovulations when administered alone. Terbutaline enhanced the number of ovulations, and propranolol counteracted the effect, whereas the nonspecific 3-receptor agonist isoproterenol was inef¬ fective. These data further support the argument that acti¬ vation of the a-receptors exerts a follicle-wall effect and that 3-receptors exert an effect via a hormonal mechanism (235). The specific role of the cholinergic receptors is undefined, but Ojeda et al. (356) note that, in the rat, abdominal va¬ gotomy delays onset of puberty and disturbs ovarian func¬ tion. Abdominal vagotomy also depresses LH and FSH se¬ cretion in unilaterally ovariectomized rats (73). In rats, section of the ovarian artery has no effect on the integrity of the ovary because of an extensive anastomosis of the uteroovarian and ovarian blood vessels. Section of the ovarian artery resulted, however, in the disappearance of histofluorescence and specific acetylcholinesterase-positive nerves. Chemical sympathectomy with 6-hydroxydopamine caused loss of histofluorescence but some retention of acetylcho¬ linesterase-positive nerves in the hilar and medullary re-

Mammalian Ovulation

gions. Pelvic neurectomy or abdominal vagotomy had no effect on ovarian adrenergic or acetycholinesterase-positive nerves. Combined pelvic neurectomy, abdominal vagot¬ omy, and chemical sympathectomy resulted in loss of ad¬ renergic nerves but persistence of acetycholinesterase-pos¬ itive nerves in the hilar and medullary areas. These observations result in the suggestion that the small incidence of acetycholinesterase-positive fibers is derived from the vagus but that most of the acetylcholinesterase is in the adrenergic nerves (74) (Table 3). The discharge of norepinephrine at adrenergic terminals is accompanied by other substances, which may modulate further norepinephrine release. These include adenosine, prostaglandins, histamine, 5-hydroxytryptamine, acetylcho¬ line (83), and perhaps (3-endorphin (288). The concept that acetylcholine enhances the secretion of norepinephrine was initially introduced by Burn (80-82,256,408). The ability of atropine to block contractile activity in the ovarian strip (489) may be a result of mechanisms other than blockade of a muscarinic receptor on the smooth-muscle cell because “. . . with few exceptions sympathetic neurons are endowed with two presynaptic receptor systems for cholinergic drugs. Nicotinic agonists depolarize the nerve endings and evoke a calcium-dependent release of noradrenaline, whereas mus¬ carinic agonists inhibit calcium-dependent release processes such as release evoked by electrical pulses, high potassium concentrations and nicotinic drugs” (448). The consensus is that the neural innervation of the follicle probably modulates the mechanism of ovulation and that the smooth muscle in the follicle wall plays no critical role in ovulation (75,131), but a contrary opinion implies a major role (380).

THE OVARIAN BLOOD SUPPLY Morphology The blood supply to the ovary was first studied in detail by Clark (108) and was reviewed by Reynolds (416), who described the embryogenesis and the adult vasculature of the ovary, and by Ellinwood et al. (143), who reviewed the various methods available for measuring blood flow through the ovary and paid particular attention to the probable role of the uteroovarian veins in control of corpus luteal function. The blood and lymph supply of the ovary has also been extensively reviewed by Gillet et al. (185). In most mammals the ovarian arteries arise from the ab¬ dominal aorta inferior to the renal arteries and reach the ovaries through the mesovarium. The ovarian artery at the hilus gives rise to many primary and secondary spiral ar¬ teries. The latter give origin to a rich capillary plexus (237,416), which surrounds the follicle with a dense basketwork of capillaries. In rabbits the venules draining the capillary plexus are more numerous and have a larger di¬ ameter and thinner wall than the arterioles (237). The post¬ capillary venules merge into secondary and primary veins

/

459

and take a direct route to emerge from the ovary at the hilus (84). Growth of the capillary plexus in the rabbit ovarian fol¬ licle has been examined by determining the incidence of labeled cells in the membrana granulosa and in the endo¬ thelial cells of the theca interna after stimulation with hCG. The labeling index for both sets of cells is correlated with follicle size and increases in parallel with it. Because the relative number of capillaries constituting the capillary plexus remains constant as the cohort of tertiary follicles (250-900 (jim in diameter) grows in response to hCG stimulation, the size of the plexus must increase to keep pace with the growth of the follicle (262). Growth of the capillary plexus in the preovulatory follicle may be under the control of an angi¬ ogenic factor, whose presence has been demonstrated in extracts of nonluteal porcine ovarian tissue (306) and in extracts of rat follicles (260). At the level of the preovulatory follicle, the initiation of ovulation may be said to be presaged by a hyperemia (523), later followed by an edema (49,84,86) involving the theca externa and interna. Espey (153) has hypothesized that ovu¬ lation may be an inflammatory process. Consequently a number of workers have examined the morphology of the microvasculature of the rabbit preovulatory follicle and the changes in its permeability (236,237,363-368) and that of the capillary plexus of the human follicle (361). The vascular supply to primary follicles is represented by a simple capillary network whose complexity grows with growth of the follicle, assuming the appearance of a mul¬ tilayered capillary plexus. The capillaries take origin di¬ rectly from, and frequently at right angles to, the arterioles and drain abruptly into venules. This pattern makes possible a high arterial pressure throughout the microvasculature. The venules are readily distinguishable from the arterioles by their larger diameter and by the larger number of cap¬ illaries entering into them (361). After treatment with hCG, preovulatory follicles enlarge without apparent change in the capillaries. Prior to rupture of the apex, the capillaries become dilated and the injection resin leaks into the interstitial space (236,237). Increase in capillary permeability is manifested by increased numbers of pinocytotic vesicles, endothelial fenestrations, and interendothelial gaps (49,366). The pinocytotic vesicles are present without change in number, size, or distribution throughout the preovulatory phase. Fenestrations increase in number, reaching maximum at 4 hr, and interendothelial gaps large enough to allow carbon particles to pass through appear late, at 10 and 12 hr after administration of hCG (365). That the interendothelial gaps were large enough to allow carbon particles to pass through implies that at a much earlier stage they were large enough to allow macromole¬ cules to pass through; this is especially likely because in¬ crease in the accumulation of iodinated serum albumin by PMSG-stimulated hCG-triggered immature rat ovarian fol¬ licles is linear from the time of the initial bolus of hCG to ovulation (296).

460

/

Chapter 12

Quantitation of the Preovulatory Blood Flow That hyperemia is a prelude to ovulation has long been appreciated. Its presence was noted by Zondek et al. (523), although Heape (209) had noted the congested state of the blood vessels of the Graafian follicle. In rabbit follicles autotransplanted to the anterior chamber of the eye, the onset of the hyperemia was described as occurring within minutes after administration of hCG (295). This observation has been quantitated both in the whole ovary and in follicles in a number of studies with several different techniques. Ellis (144), Lipner and Smith (296), and Wada (486) measured radioiodinated serum albumin content of rat ovaries; Wurtman (513) employed a 42K indicator fractionation technique. The thermocouple technique for blood flow measurements (143) was used by Makinada (305) for hemodynamic study of blood flow through the ovarian cortex. Direct outflow of blood was measured in rats (401), in sheep (112,219,315), and in pigs (112). The method of choice for quantitating blood flow through the ovary and follicles with the least number of artifacts, and the method most frequently em¬ ployed, is the radioactive microsphere technique (RMS) introduced by Janson (225) for ovarian studies (143). Physiological Changes in Blood Flow Wurtman (513), using the indicator method, measured an increased ovarian blood flow within 6 sec after administra¬ tion of LH, whereas Janson (225) recorded, using the RMS technique, an increased blood flow within 2 min. The in¬ creased blood flow occurs in the absence of any change in arterial pressure and is therefore attributable to a decrease in vascular resistance, most probably as a result of arteriolar vasodilation (225). Blasco et al. (57), also using the RMS technique in the rabbit, observed that blood flow through the ovary during the preovulatory phase increased fourfold by 4 hr after hCG administration, whereas blood flow (ex¬ pressed as percent of cardiac output) to other organs (ovi¬ duct, uterus, brain, kidney) did not change. Lee and Novy (280), using the RMS technique, confirmed that blood flow in the rabbit ovary increased after administration of either 10 pg or 100 pg of LH and that the increase was independent of the dose of LH. The increase in absolute blood flow through the ovaries was at peak at 10 min (the earliest time measured after the control measurement) and had begun to decline by 60 min. To measure the blood flow to the dom¬ inant preovulatory follicle by the RMS technique, Murdoch et al. (342) excised the follicle and divided it into apical and basal portions and follicular fluid, then correlated the radioactivity with change in serum LH. Blood flow to the follicle (apex + base) was elevated after the rise in LH began during the 0-to-12-hr period, declined during the 12to-16-hr period, and continued to decline at 20 hr and until after ovulation at 24 hr or later. No preferential blood flow to apex or base was observed, and no leakage of labeled microspheres into the follicular fluid was noted (342). The

hyperemia initiated by LH is rapid in onset and persists for the rest of the rising limb of the LH surge, after which it either reaches a plateau or declines, suggesting that the blood flow, having reached a maximum, becomes constant for that period of the life of the follicle. Although blood flow is constant, the amount of protein passing through the walls of the capillary plexus increases (296) commensurate with the increasing edema and increasing follicular fluid volume (292). Vasodilatory Substances and Ovarian Blood Flow Histamine The earliest response of the capillary plexus of the pre¬ ovulatory follicle, occurring within seconds of the onset of the LH surge or administration of a bolus of LH or hCG, As a hyperemia (523), which persists and develops into an edematous reaction (153). The mechanism for the rapid response is undefined but may be caused by a rapid release of histamine (229,291,331,461,462,513) by mast cells lo¬ cated around the blood vessels in the hilus (230,264,491). The effect is most apparent in the preovulatory follicles, probably because of their extensive vascularization. The capillary and venule endothelia are the most likely sites for the changed permeability leading to edema. The formation of edema is primarily a result of increased macromolecular (protein) efflux, with consequent decrease in the lymph/plasma total-protein ratio thus decreasing the transmural colloid osmotic pressure gradient (192). Contractile elements in the endothelial cell and the pericytes activated by Ca2+ are hypothesized to enlarge the intercellular junctions, enlarging pores through which the protein can pass. Activation of the contractile elements requires the interaction of vasodilator substances with cellular receptors and an augmentation of Ca2+ influx, which also induces formation of prostaglan¬ dins. The latter facilitate influx of more Ca2+ and contrac¬ tion of the pericyte (321). The major site of action of most substances that increase macromolecular permeability is the postcapillary venule. The permeability of the postcapillary venule is increased by the mediators listed in Table 5. A number of inhibitors of postcapillary venule perme¬ ability have been described that also antagonize effects in¬ duced by histamine (Table 6). Among the early attempts to manipulate the level of histamine in the rat ovary and to block its action was that by Lipner (291). He used an Hj blocker and also attempted to deplete the histamine levels; however, the rats ovulated, indicating either inadequate deTABLE 5. Mediators of postcapillary venule permeability3 Histamine, 5-hydroxytryptamine Bradykinin, substance P ADP, adenosine, inosine Prostaglandins E2, F2 Leukotrienes C4, D4, E4, B4 Modified from Svensjo and Grega (456).

Mammalian Ovulation

TABLE 6. Inhibitors of histamine-induced permeability3 p2-Receptor agonists (isoprenaline, terbutaline, salbutamol) Calcium antagonist (Verapamil) Glucocorticoids (budesonide, dexamethasone, methylprednesoline) Ht-blocker antihistamine (pyrilamine) Phosphodiesterase inhibitors (theophylline, xanthines) Arginine vasopressin aModified from Svensjo and Grega (456). pletion of histamine and blockade or that hCG could drive the ovulation cascade independently of histamine. Blockade of ovulation with Hi antihistamines has been observed in vivo in rabbits (253) and in vitro in a perfused rabbit ovary (488); however, histamine may not be essential for ovula¬ tion, because other vasodilatory substances are probably also involved. In a study using the in vitro perfused rabbit ovary preparation, neither H, blockade with chlorphenira¬ mine nor H2 blockade with cimetidine, nor combination of the two, could block hCG-induced ovulation (255), even though in this preparation histamine alone could induce a low incidence of follicle rupture. Questions about the role of histamine in the cascade of events leading to ovulation are raised that still require resolution. Especially pertinent is the observation that the hCG-stimulated ovaries, even with exposure to Hi and/or H2 blockade, still show the edema so well summarized by Espey (153). Estrogens are capable of inducing a vasodilation in re¬ productive tissues (of most mammals) (423,461). The factor usually associated with the estrogenic response is histamine, although other vasodilatory substances are unexplored. Be¬ cause the estrogenic effect has a 30-min latency period and the response to LH or hCG is almost instantaneous (513), estrogen is probably not an immediate intermediate but may be involved in later maintenance of the hyperemia. Bradykinin Plasmin acting on prekallikrein generates the protease kallikrein, which releases from a high-molecular-weight plasma kininogen the nonapeptide bradykinin; low-molec¬ ular-weight plasma kininogen is converted by a tissue pro¬ tease, kallikrein, to the decapeptide kallidin, which by the action of an aminopeptidase is converted to bradykinin. The kinins act on the arterioles to induce vasodilation and on the small venules to increase the permeability of the mi¬ crocirculation, leading to edema. Kinins also release his¬ tamine from mast cells, which then augment the kinin vas¬ cular effect (134). Bradykinin is present in follicular fluid (407); its con¬ centrations in the follicle increase as the preovulatory follicle approaches the time of ovulation, and the levels reached are depressed by large doses of indomethacin, which block ovu¬ lation (161). The importance of this observation is obscure because the experiment does not distinguish between the actions of prostaglandins and bradykinin. Bradykinin re¬

/

461

leases arachidonic acid from cell membranes (266), and it is probable that in this study (161) the indomethacin exerted a dual effect. In vitro, neither bradykinin, histamine, nor angiotensin II had any effect on the rate of prostaglandin synthesis (259). Therefore, although increased synthesis of bradykinin parallels the increased synthesis of the prosta¬ glandins, it has a minimal relationship to the early hyperemic reaction and to the induction of ovulation. Bradykinin may, however, contribute to the edema subsequent to the LH surge. Because plasmin synthesis is blocked by indometh¬ acin (92), the postindomethacin increase in follicle volume is probably not caused by bradykinin, but other vasoactive substances that may still be active.

Prostaglandins Although the prostaglandins studied in follicular function are generally limited to PGF2a and PGE2, the use of the nonsteroidal anti-inflammatory drugs (NSAIDs) results in the inhibition of the synthesis of prostacyclin (PGI2), a va¬ sodilator; PGE2, an inducer of erythema and edema; PGE2a, in many tissues a vasodilator; and thromboxane (TXA2), a vasoconstrictor. Furthermore, NSAIDs may inhibit chemotaxis of cells involved in the inflammatory process, in¬ hibit labilization of membrane by lysosomes, antagonize the action of histamine and bradykinin, inhibit biosynthesis of mucopolysaccharides, uncouple oxidative phosphorylation, activate fibrinolytic activity, stabilize and block sulfhydryldisulfide reaction, inhibit production of collagenase, and suppress lymphocyte function (346). The effect of indomethacin on ovarian and follicular blood flow has been studied because the mechanism for the es¬ sential role of the prostaglandin in ovulation is still consid¬ ered to be in doubt in some laboratories. At dose levels that block ovulation, Lee and Novy (280) observed a reduction in absolute blood flow to 1.03 ml/min/g of ovary from an unstimulated control level of 1.68, a decrease of 40%. The increase of blood flow in indomethacin-treated rabbits after administration of LH at 10 min was 2.4-fold; at 30 min it was 3.0-fold, and by 60 min it was 3.4-fold. In the group treated with LH only, the comparable increases were 3.4-, 3.5-, and 2.5-fold for similar intervals. Systemic treatment with indomethacin causes a reduction in prostaglandin levels in most vascular beds, a reduction in blood flow, and an enhanced vasoconstrictor response; by analogy, it probably attenuates the ovarian response to vasodilator stimuli, and PGF2a induces an increase in ovarian blood flow (349). The importance of the prostaglandin vasodilator mechanism in the generation of the early hyperemic response is question¬ able in any case, because the peak of prostaglandin synthesis occurs hours after the initiation of the maximum vascular response to LH. Since LH exerts its cellular effects by activating adenylate cyclase in its target cells, it is possible that either cAMP or progesterone might effect the vascular response (339); however, there is no latency between the

462

/

Chapter 12

onset of progesterone secretion and augmented ovarian blood flow in ewes during the estrous cycle, as determined with Doppler ultrasonic transducers (347). That the effect of pros¬ taglandins on the vascular supply to the follicle is not related to ovulation was further demonstrated in ewes by Murdoch and Myers (340). They found, after administration of indomethacin, that the amount of blood in the follicles was greatly increased, that the follicles were increased in di¬ ameter (an observation also made by others; see ref. 243) because of an increased volume of follicular fluid, and that the amount of PGE2ot in the follicle wall was reduced, but that the amount of blood in the vessels of the wall was very noticeable. In order for fluid to accumulate in the follicle, blood flow must persist (292), and it becomes questionable that inhibition of ovulation by blockade of prostaglandin synthesis is mediated by controlling follicle blood flow.

THE ROLE OF SMOOTH MUSCLE IN OVULATION Presence of smooth muscle in both the stroma of the ovary and the follicle wall (Table 7) has generated a search for its role in ovulation. The presence of smooth muscle in ovarian stroma and preovulatory follicle, coupled with its contractility, initiated the suggestion, first introduced by Rouget (425), that ovulation is either induced or facilitated by the contractions. Because no increase in pressure in the preovulatory follicle has been demonstrated, it is unlikely that the smooth muscle contributes to an increase in pressure

TABLE 7. Distribution of smooth muscle in ovaries Species Rat Rat

Monkey Sheep Human Rabbit

Cat Monkey Cat Rabbit Guinea pig Hamster

Gerbil

Hamster Cow

Location

Reference

Theca externa Corpora lutea Theca externa Interstitial tissue Atretic follicles Atretic follicles Theca externa Theca externa Theca externa Corpora lutea Cortical stroma Cortical stroma Theca externa Theca externa Cortical stroma Theca externa Theca externa Theca externa Corpora lutea Interstitial tissue Theca externa Corpora lutea Interstitial tissue Theca externa Theca externa

369 377

377 370 360 360 53 18 181 68

68 318

318

391 490

(56,65,156,292). However, it is still possible that the con¬ traction of the smooth muscle causes the wall of the follicle to collapse, facilitating the evacuation of the contents of the follicle (313,481) at the rupture point. The hamster rupture point tends to be small and may require contraction of the smooth muscle to facilitate discharge of the ovum (390,463); however, this mechanism is not generally observed. Despite the many efforts to demonstrate a role of smooth muscle in ovulation (313,430,464), no cause-and-effect data have been presented. The conclusion at present is that smooth-muscle contractions play no role in follicular rupture, but perhaps facilitate evacuation of follicle contents. FOLLICULAR FLUID The growing follicle undergoes a transformation from a solid mass of cells enclosing the ovum to a fluid-filled fol¬ licle containing a centrally located antrum into which pro¬ jects a column of cells, the cumulus oophorus, continuous with the membrana granulosa. The ovum, surrounded by the corona radiata, sits attached to the cumulus oophorus and is positioned eccentrically within the antrum (397). Initially there appear, among the multiplying granulosa cells, small fluid-filled areas, which become confluent and give rise to a single fluid-filled antrum. This follicular configu¬ ration is characteristic of many, but not all, mammals (334). The composition of the follicular fluid (liquor folliculi) changes as the follicle matures and the antral volume in¬ creases. Young antral follicles contain a primary fluid largely composed of proteoglycans. The proteoglycans become di¬ luted by a gradual influx of fluid derived from the plasma enriched by the steroids and minute amounts of proteins synthesized by the theca interna and granulosa cells. Except in the case of large mammal follicles (human, cow, and pig), data on the composition of the fluid are frequently based on analyses of pools of fluid collected from small, medium, and large follicles. However, by the defi¬ nition developed by Peters (397), all the antral follicles belong to the category of large follicles because their oocytes are large, and only the membrana granulosa and theca in¬ terna increase in cell number while the antrum increases in volume. Physical Characteristics The follicular fluid in the preovulatory follicle is very similar to plasma. It is straw-colored, although it may at times appear yellow, and has a variable viscosity, an os¬ molality equal to that of plasma, and an electrolyte content, with minor variations, almost equal to that of plasma [rabbit (79,122,436)]. The gas content of human follicular fluid has been compared to that of plasma with differing results; the Po2 is highly variable and not correlated with follicular histology, but the Pco2 is correlated approximately with pH 7.3 (435). Fraser et al. (178) note that the Po2 is higher in ovarian venous blood than in peripheral blood in women

Mammalian Ovulation

and suggest that arteriovenous shunts may account for the higher Po2 levels and that therefore the follicular fluid Po2 should be lower than the Po2 of ovarian and peripheral venous blood. Unfortunately these studies are not compa¬ rable because Shalgi et al. (435) examined the follicular fluid, whereas Fraser et al. (178), studied only ovarian ve¬ nous blood and peripheral venous blood. In the rabbit ovary, no evidence of arteriovenous shunts was found in studies based on in situ perfusion with microspheres (8). Although we could not record a potential difference across the follicle wall in rats or rabbits (292), a small positive potential (+1.2 ± 0.3 mV) has been reported across the wall of the mouse follicle in cycling mice, with an increase of +3.8 ± 0.8 mV immediately before ovulation. These effects are modulated by sodium pentobarbital, sodium cy¬ anide, and ouabain, inducing a positive potential, whereas PGE, and PGF2c( caused negative potentials; the changes involved are extremely small, and their meanings are dif¬ ficult to interpret (526).

Proteins The protein content and composition of follicular fluid have been examined in cows (22,94,128,301,485,520), pigs (316,440), rabbits (122), rats (434), and women (249,307,396,437). The total protein content of the follic¬ ular fluid is equal to or lower than that of serum (Table 8). It may vary with stage of the estrous cycle (387), but the protein content in all the follicles is similar regardless of size (22). The specific proteins show some variability; fi¬ brinogen is present in lowest concentration, or absent, in pigs (440), cows (128), and women (307) and only 60% excluded (437). Rat follicles contain all the protein present in plasma (41) (Table 9).

TABLE 8. Comparison of protein concentrations in plasma and ovarian follicular fluid in several species Serum Follicular (g/100 ml) fluid (g/100 ml) Cow Cow Cow Cow Cow Pig Pig Rabbit Woman Woman Woman Woman

5.6 6.6 9.1

8.35 6.94 7.4 7.4C 7.6 6.8 5.8

4.7-5.6 4.5 5.6 7.0 6.57 7.19a 7.0b 6.8b 5.5 5.5 3.6 3.6 5.8

Fluid/serum ratio Reference

0.80 0.85 0.77 0.86 0.76 0.86 0.99 1.02 0.74 0.47 0.53 1.0

aMean of small and large follicles. ^Arterial and venous plasma, respectively. cPlasma value.

301 94 520 128 22 440 316 122 396 249 307 307 436

/

463

Attempts to demonstrate the presence of proteins specific to the follicular fluid of pigs (440), cows (128), rats (41), and humans (307,436,437) by immunizing rabbits with fol¬ licular fluid have been uniformly unsuccessful, but immu¬ noglobulin has been observed in the female rabbit repro¬ ductive tract (460). The failure to demonstrate the presence of antigenic components in follicular fluid foreign to plasma, however, indicates that the immunologic technique is in¬ sufficiently sensitive to detect such substances, because rat granulosa cells secrete inhibin (146), and inhibin has been demonstrated to occur in the follicular fluids of cows, pigs, horses, and humans (121,231,322,432). Various factors suspected to be present in follicular fluid have been noted (132,174,251,279,325) and summarized (101,201,207, 208,418). The control of the permeability of the capillaries of all antral follicles, small or large, must be subject to the same factors because the protein contents of the fluid collected from small and large follicles are, with minor variation, similar (22). Furthermore, the permeability of the capillary wall must be high, because the protein content of the fol¬ licular fluid is very similar to that of serum (Table 8), and only the distributions of specific proteins differ (Table 9). Although a blood-follicle wall barrier has been suggested (437), the exclusion of molecules with molecular weights greater than 1,300,000 and partial exclusion of fibrinogen, with a molecular weight of 340,000, suggests a passive phenomenon of molecular sieving; however, even the larg¬ est molecules are found in the follicular fluid (441). Hormones The concentrations of hormones in the follicular fluid have been summarized by McNatty (317) and more recently by Guraya (201). The largest body of information on the gonadotropin content is based on studies of women. In gen¬ eral the levels of FSH and LH reflect the circulating blood levels, but prolactin levels are more closely and inversely related to follicular fluid volumes. The steroid concentra¬ tions reflect the stimulation of biosynthesis and are depen¬ dent on the rate of synthesis and diffusion from the theca interna and membrana granulosa and on the presence of steroid-binding proteins that cause enrichment of the steroid concentration in the follicular fluid (317). The integrity of the follicle is dependent on the sequential exposure of its constituent cell population to FSH and LH, with prolactin serving to modulate the synthesis of steroids and the response to FSH and to affect follicle growth and development (147). Enzymes Schochet (428) initiated the search for the enzymatic mechanism of ovulation by demonstrating that follicular fluid could attack fibrin, boiled connective tissue, muscle, and ovarian tissue. The spectrum of enzymes identified in

464

/

Chapter 12

TABLE 9. Distribution of plasma proteins in serum and follicular fluid (F.F.)a Women (396) Molecular weight

Protein Total protein

Albumin arGlycoprotein a-Globulin arGlobulin a2-Globulin 3-Globulin PrGlobulin 32-Globulin IgG Transferrin Fibrinogen Haptoglobulin IgA a2-Macroglobulin IgM 3rLipoprotein

Women (249)

Women (437)

Cow (22,520)

Serum

F.F.

Serum

F.F.

Serum

F.F.

Serum

F.F.

7.17

5.66

7.4

5.5

5.8

5.8

6.6

5.6

61.4

64.8

12.2

7.7

10.3 8.8 73

9.8 10.5 84

69,000

45.1 6.6

6.8 13.8

150,000

27.8

% of Protein in serum or follicular fluid 56.5 63.5 52 69 3.5 2.8

9.7 11.4

19.2

6.1 10.6

15.2

340,000

3.7

900,000 1,300,000

Present(441)

1 1

Undetectable

aAndersen et al. (22) identified 37 individual proteins common to plasma and bovine follicular fluid by precipitate and 40 by crossed immunoelectrophoresis, including fibrinogen and its split products. the follicular fluid from women, cows, and sows is listed in Table 10. Except for plasmin and collagenase, it is doubt¬ ful that these enzymes play a direct role in ovulation. It is also questionable that the enzyme in rabbit and sow follicular fluid that attacked the synthetic collagenase substrate carbobenzoxy-Gly-Pro-Gly-Gly-Pro-Ala (158,159,362) is a true mammalian collagenase, because this synthetic substrate is not attacked by animal collagenase, and enzymes that cleave this substrate have no action on collagen (327). Plasminogen

activator in the follicular fluid (43,44,452) may be the sig¬ nificant enzyme that initiates the proteolytic cascade that ultimately degrades the collagen and leads to rupture of the follicle (vide infra) (Table 10). Proteoglycans Metachromasia in tissues is a well-established phenom¬ enon, as are its association with the follicular fluid of young

TABLE 10. Enzymes in follicular fluid Enzyme Endopeptidase Aminopeptidase Proteinase Plasminogen activator Plasmin Dipeptidase Acid phosphatase Alkaline phosphatase Adenosine triphosphatase Fructose diphosphate aldolase Lactate dehydrogenase Hyaluronidase Amino transferase-aspartate Amino transferase-alanine Pyrophosphatase Kallikrein Nucleotidase Thromboplasmin Collagenase (151) Adenosine triphosphatase

Rat (42)

Porcine (238,292)

Bovine (317)

+ + + +

Human (97-99)

+ + +

+

+ + + +

+ + +

+ +

+ + +

+ + + +

Rabbit (161)

Mammalian Ovulation /

follicles, its decreasing intensity of staining with maturation of the follicle, and its digestibility with hyaluronidase (509). These properties led to the suggestion that the substance was composed of mucopolysaccharides (509). Administra¬ tion of 35S04 results in autoradiographic labeling of the follicular fluid because of secretion by the granulosa cells, during antrum formation (191,518,520), of a material iden¬ tified as acid mucopolysaccharides (now called proteogly¬ cans) (467). The follicular fluid in follicles with newly formed antra is jelly-like (61,204,352,518) and becomes more fluid but remains viscous (141,142) as the follicles mature. The antra of mature follicles, prepared from rat ovaries fixed in al¬ dehyde and stained with chromic acid and phosphotungstic acid, contain a complex network of fine reticular configu¬ ration (233); however, at lower magnification, the follicular fluid appears as a homogeneous granular precipitate (24). Proteoglycans are composed of a core of protein, to which is attached polysaccharide chains called glycosaminoglycans (GAGs). Seven classes of GAGs with common character¬ istics have been identified. They contain long heteropoly¬ saccharide chains composed of largely repeating units of disaccharides, in which one sugar is a hexosamine and the other is a uronic acid with sulfate groups. The carboxyl and sulfate groups are among the factors that make the GAGs highly charged polyanions. Because they hold cations and water within their domains, the proteoglycans give tissues their resiliency, form solutions of high elasticity and vis¬ cosity, and stabilize the fibrous and cellular elements of tissues. Antrum formation in estrogen-primed hypophysectomized immature female rats occurs in response to admin¬ istration of FSH (190). In a similar model, labeled sulfate is incorporated into proteoglycans linearly with the log of the dose of FSH (338). The predominant glycosamino¬ glycans of the follicular fluid are chondroitin sulfate and heparan sulfate (29,30,46,183,338,514,515), with chon¬ droitin sulfate the major glycosaminoglycan of the proteo¬ glycan found in the rat (514), pig (29,515), and cow (35,194,195). The predominant glycosaminoglycan se¬ creted in vitro by rat granulosa cells (514,516) and bovine granulosa cells (46,47,285) is chondroitin sulfate. Healthy follicles have elevated concentrations of estrogen and low concentrations of progesterone and chondroitin sulfate, whereas atretic follicles have low amounts of estrogen and elevated concentrations of progesterone and chondroitin sul¬ fate (45). The accompaniments of follicular maturation are rising concentrations of estrogen and declining concentra¬ tions of chondroitin sulfate and heparan sulfate (85). Ele¬ vated levels of cAMP increase the synthesis of proteogly¬ cans by rat ovarian granulosa cells, because incorporation of 35S04 into proteoglycans is stimulated by ovine FSH and LH, human chorionic gonadotropin, PGE], PGE2, N,0'~ dibutyryl cAMP, theophylline, and testosterone (516). Treatment of bovine granulosa cells by using phenothiazine drugs such as trifluoroperazine, chlorpromazine, and chlor-

465

promazine sulfate to inhibit calmodulin or by decreasing the concentration of calcium in the medium with EGTA inhibits the synthesis of proteoglycans in response to stimulation with either ovine or rat FSH. The synthesis of proteoglycans in response to stimulation by FSH requires activation of adenylate cyclase and the presence of calcium ions (285). The formation of the secondary follicular fluid in the preovulatory follicle occurs with the onset of the LH surge and reflects the series of events associated with changes in the follicle wall. There is an almost instantaneous increase in follicular blood flow manifested by the vasodilation. Somewhat later (6 hr after the LH surge) the increase in capillary and venule permeability is accompanied by in¬ creased volumes of interstitial fluid containing high con¬ centrations of plasma proteins. The gradually rising con¬ centrations of the prostaglandins and leukotrienes induce contraction of the smooth muscle in the postcapillary ven¬ ules (456), thus increasing the intracapillary pressure further and enhancing the rate of transudation of plasma (321). There is an increase in tone of the smooth muscle of the follicle wall in response to sympathetic nerve terminal se¬ cretion of norepinephrine and co-transmitter substances. Later the (32-adrenergic effects are exerted on the membrana gran¬ ulosa, causing further increase of prostaglandin secretion and thus maintaining the increased smooth muscle tone and venule permeability (456). The consequence of these pro¬ cesses is to raise the free fluid pressure (as measured in the antrum) to a positive value, indicating that the factors that prevent accumulation of fluid (low interstitial-fluid protein concentration, lymphatic protein removal, and low venule resistance) are no longer effective and that conditions for accumulation of increased extracellular fluid volumes now exist (192). The importance of the proteoglycans in the follicle is probably dependent on the negative charges of the glycos¬ aminoglycans, which create Donnan osmotic and hydro¬ mechanical forces (62). These act as a sink and probably are responsible for the slow initial influx of fluid and gradual expansion of the antral space. PREOVULATORY MORPHOLOGICAL CHANGES ASSOCIATED WITH OVULATION Heape (209) thought that the cause of follicular rupture was obscure. Eighty years later the cause still under inves¬ tigation is less obscure, but is still far from being revealed in its totality. The morphological changes have been ex¬ amined in much detail, but only in a relatively small number of species, and though there appears to be much similarity in these changes, many more species need to be examined. In the rabbit and rat the volume of the preovulatory follicle after a coital stimulus or hCG increases linearly (296,495). Concurrent with the increase in follicular volume, there is an increase in collagen synthesis indicated by a threefold increase in the content of hydroxyproline in ovaries of PMSGprimed immature rats. After the endogenous surge of LH,

466

/

Chapter 12

there is a further twofold increase in ovarian hydroxyproline and a subsequent abrupt decline (328). Preovulatory follicles therefore undergo a period of enhanced collagen synthesis followed by a degradation of collagen. The appearance of the preovulatory follicle as it ap¬ proaches rupture has been described in great detail. In mice (86) and rabbits (104) the follicle protrudes above the surface of the ovary as its volume increases, the capillaries in the theca appear packed with erythrocytes, and the endothelium is flattened; with time, perforations appear in the capillary walls. As time of rupture approaches, the thecal cells at the base of the follicle break through the lamina propria and begin to intermingle with the parietal granulosa cells, which show a marked increase in smooth endoplasmic reticulum, signaling their transformation to lutein cells. At the apex, the germinal epithelium lies on (a) a basement membrane containing fibroblasts (the theca) and (b) a basement mem¬ brane, the lamina propria, of the membrana granulosa. The germinal cells flatten as the follicle protrudes and gives rise to the stigma and then disappear as rupture becomes im¬ minent. The underlying fibroblasts are no longer present, but collagen fibers are seen. The theca cells beneath the fragmented epithelial layer appear necrotic. At rupture, the theca cells are lost, as is the lamina propria of the membrana granulosa. The latter cells protrude through the developing rupture point and appear intact. Similar changes have been described in the rabbit and human, with the added obser¬ vation that the collagen in the apex becomes sparse (49— 54,148,359). As the time of ovulation approaches, the preovulatory follicle develops a protrusion called the stigma. In the rat the stigma assumes one of two different configurations. Stigmas may appear as extensively bulging vesicles, called bleb-type, or as small flat avascular areas, called flat-type. The bleb-type stigmas lose their epithelial cover, exposing densely arranged fibroblasts, with multivesicular bodies (150) protruding but without collagen fibers present, surrounding a pore. The surface epithelium sloughs off the avascular area of the flat-type stigmas, revealing an underlying stroma composed of coarse longitudinal and smaller circular fibers. Beneath these are fibroblasts surrounding a small opening in the stroma, through which granulosa cells protrude (477). Throughout the estrous cycle, cell replication does not occur in the theca externa of mature rat follicles, but the granulosa cells and theca interna incorporate [3H]thymidine into nuclear DNA during diestrus and stop taking up [3H]thymidine during estrus (478). Thus, although folliclewall synthetic activities are present, they derive from a constant cell population. Increased extravasation of fluid in the follicle wall is frequently described (49-54,86,148,209,495) and is sum¬ marized by Espey (153). Edema and follicle enlargement occur even when ovulation is blocked with indomethacin (25,162,517). The determinants of ovulation therefore are neither edema nor inflammation per se but, instead, syn¬ thetic mechanisms inherent to the preovulatory follicle and initiated by LH.

PREOVULATORY CHEMICAL CHANGES ASSOCIATED WITH OVULATION The surge of LH that occurs at midcycle in women and subhuman primates, in proestrus in spontaneously cycling mammals, or subsequent to coitus in reflex ovulators is followed by a series of changes in the follicle wall that lead to its disruption and the discharge of the ovum (431). These changes are initiated with the rising concentration of LH and, after 1 hr of exposure to LH, become independent of subsequent hormonal stimulation (292). The cascade of changes may be interrupted by blockade of protein synthesis with cycloheximide or actinomycin D (17,38,403,404), by blockade of steroid synthesis with cyanoketone or aminoglutethimide (294,297), with antiprogesterone antiserum (329,458), or with antitestosterone antiserum (330), by blockade of prostaglandin synthesis with indomethacin (25), or by interruption of the plasminogen activator-plasminogen cascade (92).

Protein Synthesis The rabbit, because it is a reflex ovulator, is an interesting model for the study of ovulation. The interval from the stimulus, either coital or hormonal (LH or hCG), to the time of ovulation is 10 to 12 hr (209). During this 10- to 12hour interval, all the important changes initiated by LH must occur. A role for protein synthesis in ovulation was first explored by means of the intrafollicular injection of acti¬ nomycin D (an inhibitor of transcription, blocking synthesis of mRNA) and cycloheximide (an inhibitor of translation, blocking peptide-chain initiation and chain elongation) (403,404). Both compounds block ovulation when intro¬ duced into the follicle up to 5 hr post-coitally. This obser¬ vation made with actinomycin D was confirmed in both mature and immature hamsters (38). Diphtheria toxin and cycloheximide, both of which block translation by inhibiting peptide chain elongation, inhibited ovulation in hamsters (17). Increasing the dose of either inhibitor extended the duration of inhibition, probably indicating that protein syn¬ thesis is an ongoing process throughout the crucial period. Alieva et al. (17) note that, even though ovulation is blocked by diphtheria toxin or cycloheximide, behavioral estrus that is progesterone dependent is unaffected. They conclude that either the blocked protein synthesis is most probably as¬ sociated with enzymes involved in follicle-wall degradation (and not with steroid synthesis) or behavioral estrus is con¬ trolled by smaller amounts of progesterone than is ovulation.

Steroidogenesis The biosynthesis of steroids by the theca interna and granulosa cells has been reviewed by Erickson et al. (147), with special emphasis on the ovarian production of andro¬ gen. The best explanation for estrogen production in the follicle is based on the two-cell two-hormone hypothesis.

Mammalian Ovulation

FSH serves to induce LH receptors, and the LH causes the theca interna to shift from a progesterone- to an androgensecreting tissue. In the presence of granulosa cells, the an¬ drogen is converted to estrogen, and follicular growth and development ensue. The preovulatory surge of LH termi¬ nates estrogen synthesis by the preovulatory follicle, and the theca cells change from an androgen- to a progesteronesecreting tissue (147). Blockade of ovulation occurs when immature rats treated with PMSG/hCG or PMSG/LH are pretreated with cyanoketone (2a-cyano-4,4,17a-trimethylandrost-5-en-17(3-ol-3one), a 3(3-hydroxysteroid dehydrogenase inhibitor (297); aminoglutethimide, a 20a-hydroxycholesterol dehydrogen¬ ase inhibitor; or Su 10603, a 17a-hydroxylase inhibitor (294). The authors suggested that progesterone was a precursor molecule essential for ovulation. Bullock and Kappauf (66), using immature rats primed with PMSG/hCG and treated with aminoglutethimide or cyanoketone, found that ovula¬ tion was unaffected by aminoglutethimide, even though serum progesterone was depressed, and that with cyanoketone at the largest dose, both depression of serum progesterone and blockade of ovulation occurred. They concluded that neither progesterone nor steroids were involved in the ovulatory mechanism. This conclusion was seemingly reinforced by Hamada et al. (205), who noted that a slightly higher number of perfused rabbit ovaries ovulated when exposed to hCG and progesterone than when perfused with hCG alone. More recently, apparent dissociation of steroid biosynthesis and ovulation was again observed. Perfusion of immature rat ovaries with LH 48 hr after they were primed with PMSG resulted in follicular rupture and an increase in the concen¬ tration of estradiol in the medium and in the follicular fluid. Addition of 4-hydroxyandrostene-3,17-dione prevents the

/

467

increase in estradiol but does not prevent ovulation (261). Although the concentrations of both progesterone and es¬ trogen were reduced, the continued synthesis of neither ste¬ roid was completely blocked, and it is not known what levels of steroid, if any, are necessary to allow ovulation. There is, however, an interesting body of literature sup¬ porting the observation that progesterone, testosterone, and, by implication, estradiol are involved in ovulation. Intrafollicular injection of normal rabbit serum (Fig. 1) does not affect the incidence of ovulation (Fig. 2), but ovine pro¬ gesterone antiserum reduces the incidence of ovulation to 20% (Fig. 3), whereas rabbit estrogen antiserum or pro¬ gesterone antiserum adsorbed with progesterone are inef¬ fective in blocking or even reducing the incidence of ovu¬ lation (Fig. 4). Furthermore, hamster ovaries explanted into progesterone-containing media have multiple ovulations (36,37). Rondell (421) noted that estradiol and progesterone increase the distensibility of follicle-wall strips; however, there is no direct link between this phenomenon and ovu¬ lation. A relationship between steroids and ovulation has been established by the use of antitestosterone antiserum and antiprogesterone antiserum in rats (329,330) and in humans and ewes (33). In these studies (329,330), ovulation was blocked in PMSG/hCG-treated immature rats with antiserum against either progesterone or testosterone, and the effect of the antiprogesterone antiserum was reversed by admin¬ istration of progesterone within 6 hr after the hCG treatment. Antiprogesterone antiserum was ineffective if administered more than 6 hr after the hCG treatment (257). This obser¬ vation in the rat parallels that by Swanson and Lipner (458) (Fig. 3) and supports that concept that steroids may be involved in ovulation.

FIG. 1. Appearance of the operative field with preovulatory follicle receiving an in¬ jection in an exteriorized ovary. The tissue is kept warm and moist with a continuous 37°C drip of mammalian Ringer’s solution. In the foreground are a fat body and the oviduct. [From Swanson and Lipner (1977), unpublished report. ]

468

/

Chapter 12

O

O

FIG. 2. Follicles (numbers above col¬ umns) injected with 0.1 pi of normal rab¬ bit serum at various intervals post-coi¬ tus. No significant difference is noted between the columns, and the range of ovulations was from 60 to 100%. On average, only 80% of follicles ovulate. [From Swanson and Lipner (1977), un¬ published report. ]

CO LlJ —

I o _J

o 2 O UJ

cr

LU

Q_

POSTCOITAL

HOUR

OF

INJECTION

Beilin and Ax (45) noted that healthy follicles have el¬ evated levels of estradiol. The path to estradiol synthesis includes testosterone from sources extrinsic to the estrogensecreting cell; therefore, the most probable sites for inhi¬ bition of the ovulatory mechanism are at stages preceding the synthesis of estrogen (294,297,329,330). This finding explains the failure of MER-25 (297) and of antiestrogen antiserum introduced directly into the follicle to inhibit ovu¬ lation (Fig. 4). In immature rats primed with PMSG/hCG, both indomethacin and cycloheximide reduce the levels of estradiol and progesterone in the ovaries. In the ovaries of rats not treated with the inhibitors, the concentrations of estradiol and progesterone are inversely related, and the

steroid changes precede the rise in prostaglandins. Thus one may postulate that the steroids contribute to the regulation of prostaglandin production, which in turn may regulate enzyme production (154).

Prostaglandins Arachidonic acid is converted by prostaglandin cycloox¬ ygenase to the endoperoxide intermediate PGH2, which is then converted by the action of isomerases to a number of biologically active molecules, among which prostaglandin E2 (PGE2), prostaglandin F2 alpha (PGF2a), and prostacyclin (PGI2«) are the most abundant (455). PGI2a is unstable, and

FIG. 3. Follicles (numbers above col¬ umns) were injected with 0.1 p.1 of ovine progesterone antiserum (produced in sheep against progesterone-BSA conjugate, with a low cross-reactivity against estrogens and progestagens, other than progester¬ one; a gift from Dr. J. D. Neill and Dr. M. Freeman). There is no significant difference between the columns that have the same letters. Columns with different letters were significantly dif¬ ferent at p < 0.01. [From Swanson and Lipner (1977), unpublished re¬

port.]

POSTCOITAL

HOUR

OF

INJECTION

Mammalian Ovulation

o Z) > o

100

PROG. ADSORBED PROG. A/S

E2

A/S

80

V) LU

60 o u.

40 LlI

o cc

LU CL

20

POSTCOITAL HOUR OF INJECTION

FIG. 4. Follicles (numbers above columns) injected with ovine progesterone antiserum and adsorbed with progesterone (left side) or injected with rabbit estrogen antiserum (batch E27a-H was a gift from Dr. K. Wright and Dr. D. C. Collins, prepared in rabbits with estradiol-170-carboxymethoxine thyroglobulin). No significant difference was found from the ex¬ pected incidence of ovulation with either treatment. [From Swanson and Lipner (1977), unpublished report.]

its presence is manifested by that of the stable molecule 6keto-PGFla. Prostaglandin cyclooxygenase is subject to in¬ hibition by NSAIDs such as aspirin, ibuprofen, piroxicam, and indomethacin. Inhibition of prostaglandin cyclooxy¬ genase effectively blocks the synthesis of all the prosta¬ glandins of the PGH2 series. An alternative pathway for the oxygenation of arachidonic acid is provided by lipoxygenase enzymes, which are insensitive to NSAIDs. The products of the lipoxygenase enzymes are hydroperoxyeicosatetranoic acids (HPETEs). Among the latters’ degradation prod¬ ucts are 5-hydroperoxyeicosatetranoic acid, which can, by epoxide formation, give rise to the leukotrienes (266). The prostaglandins have numerous and varied actions, among which are smooth-muscle contractile (PGF2a) or relaxant (PGE2) activity, activation of adenylate cyclase, participa¬ tion in the inflammatory reaction, and activation of proteo¬ lytic enzymes. One of the important actions of the leuko¬ trienes is in their chemotactic effect on polymorphoneutrophils and their involvement in the inflammatory reaction. Inhibition of cyclooxygenase with indomethacin blocks ovulation (25), and blockade of lipoxygenase activity with nordihydroguaiaretic acid (a nonspecific inhibitor of both prostaglandin cyclooxygenase and lipoxygenase; see ref. 384), 3-amino-l-(3-trifluoromethylphenyl)-2-pyrazoline hydrochloride (BW 755C), and FPL-55712 (lipoxygenase inhibitors; see ref. 95) results in partial blockade of ovulation (413). Furthermore, rat ovarian and follicular homogenates

/

469

both possess lipoxygenase activity that increases after in vivo administration of hCG (414). These observations im¬ plicate the leukotrienes as well as the prostaglandins in follicular rupture. Rat granulosa cells obtained from PMSG-treated imma¬ ture rats form 6-keto-PGFla and PGE2, with maximum stim¬ ulation occurring in the presence of arachidonic acid and LH. Although arachidonic acid, LH, and lueteinizing hor¬ mone releasing hormone (LHRH)-ethylamide (a potent LHRH agonist) stimulate prostaglandin formation, it has been found that histamine, bradykinin, and angiotensin II have no ef¬ fects on the rate of prostaglandin synthesis (259). The sig¬ nificance of the ability of the granulosa cells to synthesize 6-keto-PGFla resides in the observation that prostacyclin (PGI2a, a major arachidonic acid metabolite; see ref. 405) and the 6-keto-PGFla that appears as a stable oxidative product of the PGI2a decrease the incorporation of [3H]proline into human cervical tissue during the follicular phase and increase the incorporation of the labeled amino acid during the luteal phase of the menstrual cycle (503). The steroid milieu of the tissue apparently influences the action of the prostaglandin. A similar control of collagen synthesis prob¬ ably holds in the Graafian follicle, because incorporation of [3H]proline into the tunica albuginea of human preovulatory follicles is depressed by PGE2 (127). Prostaglandins acting on cultured follicles mimic the ef¬ fects exerted by LH. PGE2 can induce adenylate cyclase activity, activate protein kinase (269,270) and ornithine de¬ carboxylase (270,376), increase glucose oxidation (395), and induce luteinization and resumption of meiotic division of the oocyte (289,474). PGF2a is less effective than PGE2 in increasing cAMP accumulation and in inducing the first meiotic division of the oocyte (290). Any substance capable of increasing cAMP accumulation is effective at inducing maturation of follicle-enclosed ova. Among these com¬ pounds are LH, FSH, PGE2, and PGF2cx. Whereas FSH induces secretion of progesterone, LH and PGE2 induce the secretion of progesterone, androstenedione, and estradiol (289). The action of LH on the biosynthetic activities of cultured follicles is independent of concurrent synthesis of prosta¬ glandins. In the presence of flufenamic acid [N-(a,a,atrifluoro-m-tolyl) anthranilic acid], indomethacin, or aspirin (prostaglandin cyclooxygenase inhibitors), LH stimulates cAMP accumulation and progesterone release from the fol¬ licles (289). Furthermore, after prolonged exposure to LH, a subsequent stimulation with LH elicits no further acti¬ vation of adenylate cyclase (282), but PGE2 provokes ad¬ ditional adenylate cyclase activation (289). The actions of LH and PGE2 are independent and parallel, and PGE2 mim¬ ics the LH effects (290). Although whole rabbit or rat follicles respond to exposure to LH with increased prostaglandin synthesis (40,281), dem¬ onstration that isolated follicle components secrete prosta¬ glandins under gonadotropic stimulation has been achieved only occasionally. Rabbit granulosa cells secrete PGF2a but

470

/

Chapter 12

are unstimulated by human menopausal gonadotropin (hMG with LH/FSH activity) (100). Rabbit follicles dissected into thecal wall and granulosa components secrete prostaglan¬ dins, but neither follicle component is stimulated by LH (474). The granulosa cells from PMSG-treated immature rats do, however, show a response to LH with an increase in PGE synthesis (109). There is an increase in PGF in pieces of follicles of human ovarian tissue after incubation for 72 hr and continuous exposure to hCG and hMG (402); in human granulosa cells, hCG or hCG and hMG stimulate PGE production (388). Rat ovarian theca incubated with FSH increases PGE formation 15-fold over basal levels and threefold over the stimulated rate in granulosa from the same follicle (524). Theca and granulosa from gilts treated with PMSG and hCG and then cultured with LH or FSH are differently affected. FSH exerts no additional stimulation, but LH stimulates the granulosa to increase production of PGE2 (166). In a study of dispersed granulosa cells and theca interna cells obtained from prepubertal gilts treated with PMSG and hCG, both cell types produce PGF2a and PGE2; the latter was the major product produced. Neither FSH nor LH induced responses in these cell types (11). Either the appropriate in vitro conditions have yet to be achieved, there is a step (or steps) yet to be elucidated, or the interaction of the theca interna and granulosa is required for the gonadotropin stimulus to be effective. Furthermore, the greatest success with LH stimulation occurs after pre¬ vious exposure to PMSG. These observations are very sim¬ ilar to those describing the control of the production of plasminogen activator and suggest that maturation of the follicle subsequent to exposure to FSH must precede the response to LH (see subsection entitled “Plasminogen”). In the absence of prostaglandins the final phase of ovu¬ lation—the follicular rupture—does not occur (281). Thus, although LH induces the process of ovulation, prostaglan¬ dins must be present for the final phase. Inhibition of pros¬ taglandin synthesis by administration of indomethacin blocks ovulation in rats (25,324,475), rabbits (26,196,353), mar¬ moset monkeys (304), and gilts (10). The concentrations of PGE and PGF in the ovaries, fol¬ licles, and follicular fluid are indicated in Table 11. In rats, rabbits, humans, and swine the prostaglandin levels rise as time of ovulation approaches. In rabbits, PGF2a increases 10-fold and PGE2 increases fivefold 5 hr after an ovulatory dose of hCG, and by 10 hr these prostaglandin concentra¬ tions have increased 60-fold and 15-fold, respectively (284). PGE2 is present in higher concentrations initially than is PGF2c(, and only by 9 hr does the ratio of PGE2 to PGF2 approach unity. In rats and sheep the increase in follicular prostaglandin is initiated by the LH surge and reaches max¬ imum concentration as time of ovulation approaches (39,289,341). Prepubertal pigs treated with PMSG and hCG ovulate approximately 116 hr after the administration of the hCG. PGF in the follicular fluid of preovulatory follicles increases to a maximum by 90 to 92 hr (9). That prosta¬ glandins are probably involved in ovulation in humans is

indicated by the increased synthesis of prostaglandins by human ovaries exposed to human menopausal gonadotropin (402) (Table 11). Prostaglandins have been implicated in a number of di¬ verse effects on the ovarian follicle. PGF2a may be involved in maturation of guinea-pig (465), and prostaglandins in maturation of mouse (137), ovarian follicles. PGE2 inhibits the synthesis of collagen within the tunica albuginea of preovulatory human follicles (127) but stimulates the syn¬ thesis of proteoglycans in this tissue (468). In mice, PGE2 induces cumulus expansion associated with synthesis of hy¬ aluronic acid (145). In rabbits, blockade of prostaglandin synthesis with indomethacin prevents the appearance of pro¬ teolytic enzymes (collagenases) specifically associated with collagen degradation (215,216,246). PGF2a injected into rabbit Graafian follicles induces the expulsion of the ovum along the path through the follicle wall created by the can¬ nula (27), and PGF2a injected intraaortically in indomethacin-treated rabbits increases ovarian contractility (130). Follicle walls prepared from human ovarian follicles are spontaneously contractile and manifest a significant increase in tone when exposed to PGF2a, but a small decrease in tone when exposed to PGE2 (357,358). In a similar study performed on sow ovarian follicles, PGF2ct decreased the tension generated in early and late preovulatory strips, whereas PGE2 increased the basal tension (187,188). Guinea-pig ovaries develop a marked increase in isometric tension when exposed to either oxytocin or PGF2a during late proestrus (189). The effect of the prostaglandins on contractility of whole rabbit ovaries has also been examined both in vivo and in vitro. PGF2a causes an increase in intraovarian pressure and contractions of human ovarian stroma in situ, whereas PGE2 has no effect on the intraovarian pressure, but the combi¬ nation of the two prostaglandins in equal quantities is stim¬ ulatory (116). In vitro study of human ovarian tissue pre¬ pared from proliferative-phase follicles indicates that PGE2 causes a decrease, and PGF2a an increase, in tonic con¬ tractions (117). In situ ovaries are spontaneously contractile even without prior exposure to hCG. PGF2a administered via the aorta increases tone and amplitude of contractions, and PGE2 similarly administered induces a reduction in tone and contractile activity. In vitro measurements yield similar responses to the two prostaglandins. In hCG-treated rabbits the ovaries both in vivo and in vitro were spontaneously active, and PGF2ct stimulates increases in tone and amplitude of contraction (483). Although the prostaglandins affect the smooth muscle of the ovary and probably the follicle, a relationship to ovulation is still not established. The effect of PGF2a on ovaries of most animals examined is to increase tone and contractility of the smooth muscle in the follicle wall. PGE2, on the other hand, causes a reduction in the tonic contraction and obliterates contractile activity. The number of actions elicited by the prostaglandins on tissues are numerous and diverse, but the mechanism by which they act on the ovary to elicit ovulation is now be-

Mammalian Ovulation /

471

TABLE 11. Prostaglandin levels in ovaries, follicles, or follicular fluid Prostaglandin (pg/mg) Species Rat

Rabbit

Treatment

Tissue

Estrous Graafian cycle follicle Proestrus 8:00 a.m. 12:00 p.m. 4:00 p.m. 8:00 p.m. 12:00 a.m. Estrus 4:00 a.m. 8:00 a.m. None

PGE

Prostaglandin (pg/follicle or pg/ml)

PGF

PGE

11+1.0 10 + 1.0 14 + 3.0 370 + 210 746 + 275 49 + 43 25 + 7.0

18 19 22 247 369

+ + + + +

2.5 3.0 6.0 102 63

69 + 50 27 + 6.0

37 37 1,050 + 51 2,570 + 509 118 + 100 66 + 21

26

284 70.0 + 27.9 15.2 + 5.9

(100 1 hr 5 hr 9 hr

(4) (4) (5)

107 + 45 335 + 191 816 + 246

19 + 4.3 296 + 321 763 + 247

Whole follicle, preovulatory Late proliferative follicle Theca granulosa PMSG, hCG Time after hCG (hr) 0 4 30 35 38 40

Sheep0

161 + 132 70 + 17

742 + 174

Graafian follicle (5)a

Human

Swine (pre¬ pubertal)

77 77 746 + 2,701 1,380 + 263

20

50 g NIH-LHB7 (9i hr post-LH)

None hCG IU) after after after

Reference 283

Graafian follicle

Rabbit

PGF

3,160

70.6 n.d.b

840

87 n.d 476

Pooled follicular fluid

(4) (4) (5) (6) (4) (4)

Estrous cycle Follicle wall Pre LH rise Hours after LH rise 0 4 8 12 16-20

aNumbers in parentheses indicate the number of animals. bn.d., not detected. cValues are approximate, based on Fig. 5 [Murdoch et al. (341)].

388

1,200+200 160+20 1,100 + 100 140 + 30 6,300 + 800 6,100 + 1,100 37,600 + 26,300 38,600 + 1,800 176,000 + 26,000 231,000 + 55,000 196,000 + 192,000 + 161,000 341 2,300

5,700

2,600 2,600 13,000 15,000 5,000

6,600 6,600 11,000 12,600 10,600

472

/

Chapter 12

coming more apparent (92,412). NSAIDs block the syn¬ thesis of prostaglandins and inhibit ovulation, but dexamethasone, a very effective steroidal anti-inflammatory drug, has little or no effect on ovulatory rate (161). NSAIDs also inhibit the activation of neutrophils and inhibit calcium ion (Ca2 + ) movement into them; they therefore prevent the re¬ lease of other inflammation-provoking substances (2). These observations can be extended to the ovarian follicle; NSAIDs may prevent the release of enzymes from polymorphoneutrophils as well as from other cells that participate in ovu¬ lation. However, it should be noted that PGE2 affects ovu¬ lation in indomethacin-blocked animals (25,473) and thus plays a major role in ovulation. PGE2 causes vasodilation and smooth-muscle relaxation, and PGF2 causes venomotor smooth muscle contraction and elicits this effect on ovarian follicle smooth muscle (358). PGE2 is the predominant pros¬ taglandin in the follicle, but the PGF2a may exert an effect on the smooth-muscle elements of the follicle wall (130). Furthermore, antiserum to PGF2c( injected into preovulatory follicles of LH-primed rabbits blocks ovulation of the anti¬ serum-treated follicles only (26); in indomethacin-blocked rats, ovulation can be induced by administration of PGE2 (473,475). These diverse observations have now become interpretable as a result of more recent studies (see subsec¬ tions entitled “Plasminogen” and “Collagenolysis”).

Step 1 Preovulatory follicle

FSH, FSH

FIG. 5. A hypothesis describing the proteolytic cascade in¬ volved in degradation of the collagen in the preovulatory follicle wall. Step 1: The preovulatory follicle stimulated by gonado¬ tropins secretes urokinase-type plasminogen activator. Step 2: Urokinase-type plasminogen activator converts plasmino¬ gen to plasmin, and the production of plasminogen antiactivator

Plasminogen Enzymatic degradation of the follicle wall is probably the best hypothesis explaining follicular rupture (42,44, 151,152,292). Espey (152) is a proponent of the concept that activation of the fibroblast and its release of an activator (Fig. 5, step 3) to convert latent collagenase is the mech¬ anism controlling ovulation. The plasminogen-activatorplasminogen hypothesis (Fig. 5) is the most recent and most likely explanation of the mechanism initiating the cascade that leads to follicular rupture. The study of this system has resulted in an intensive evaluation of the various steps in the postulated cascade. The plasminogen-activator-plasminogen system may be important in situations where controlled proteolysis and tis¬ sue degradation are necessary, as in involution of the mam¬ mary gland at termination of lactation (374), rupture of the ovarian follicle at the time of ovulation (42,44,452), and embryonic reorganization (311,453). The advantages of this system are based on the ability of the cell to secrete small amounts of a specific protease (plasminogen activator) to generate a second protease (plasmin) with a broad range of substrates. The second protease (plasmin) has its pH opti¬ mum at the pH of extracellular fluid. The proenzyme (plas¬ minogen) is present in plasma in high concentrations (0.5

testosterone ->

urokinase-type plasminogen

is decreased under the influence of the gonadotropins. Step 3: Latent collagenase is activated by the plasmin, and active collagenase is generated. Step 4: The collagenase attacks the collagen, giving rise to telopeptide-free collagen. Step 5: Telopeptide-free collagen is degraded by nonspecific proteases.

Mammalian Ovulation

mg/ml), and the space of action of plasmin is limited by the presence of plasma protease inhibitors (311). Secretion of plasminogen activator produced by bovine aortic endo¬ thelial cells (438) and macrophages (479) is subject to in¬ hibition by similar compounds (Table 12). Plasminogen ac¬ tivator production by granulosa cells is subject to stimulation by a number of substances (Table 13). Plasminogen is a glycoprotein contained in the plasma and is converted to a serine protease by proteolytic modi¬ fication. Circulating plasminogen is present as two nonin¬ terconvertible isozymic forms differing only in their state of glycosylation. In adult animals, they are principally syn¬ thesized in the liver and are continuously secreted (59). Plasminogen is converted to the active serine protease by two different plasminogen activators (PAs): urokinase (uPA), found in urine (504); and tissue-type activator (tPA), found in animal tissue homogenates (28) and human tissue extracts (16). The two PAs are immunologically unrelated, and each is immunoprecipitated by a specific antiserum. Urokinase (MW 40,000-52,000) is secreted by endothelial cells; tPA (MW 74,000) is contained in endothelial cells (287) and is probably the vascular PA that is released upon adequate stimulation and that binds to and requires fibrin for its ac¬ tivity (111). Cultured granulosa cells produce both uPA and tPA (91,92,252,351,412). Plasminogen activators are present in human ovarian tis¬ sue extracts (16), in bovine follicular fluid, and in the su¬ pernatant fluid of follicle-wall homogenates (42). The fol¬ licular fluid also contains plasma protease inhibitors that are destroyed in acid media (42) and which may be derived from the plasma and anti-plasminogen activator whose se¬ cretion by granulosa cells is inhibited by gonadotropins (351). Granulosa cell lysates, prepared from ovarian preovulatory follicles of immature rats treated with PMSG/LH or from

TABLE 12. Inhibitors of the secretion of plasminogen activator activity Compound Cycloheximide Colchicine Hydrocortisone Dibutyryl cAMP Theophylline Cholera toxin Colchicine Dibutyryl cAMP Epinephrine Isoproterenol Prostaglandin Et Prostaglandin E2 Steroid hormones (dexamethasone) Vinblastine

Test tissue

Reference

Bovine endothelial cells

438

Peritoneal macrophages

479

/

473

TABLE 13. Substances increasing granulosa cell production of plasminogen activator Substance Dibutyryl cAMP Dibutyryl cAMP + theophylline LH Prostaglandin E! Prostaglandin E2 FSH FSH + testosterone Cholera toxin

Reference 44 44 44 44 44 452 298 496

mature proestrous rats, contain PA, which increases in con¬ centration as time of ovulation approaches (44). Granulosa cells cultured with FSH or LH in vitro produce PA, but FSH in this system is more effective than LH. Cyclic AMP and its analogs also stimulate production of PA, and theophylline potentiates the response of the gran¬ ulosa cells to LH. PGE! and PGE2 stimulate production of granulosa-cell PA; the largest responses are obtained in the range between 10-8 and 10-6 m. Because the gonadotropins and the prostaglandins stimulate the adenylate cyclase path¬ way of the cell, cAMP probably activates the protein syn¬ thetic system, leading to increased production of PA (452). Peak levels of PA activity in PMSG-stimulated immaturerat granulosa cells in culture are obtained at concentrations of 30 ng FSH/ml, with the response beginning after 2 hr of exposure to the gonadotropin (43,92). An extensive analysis of the hormones stimulating PA activity of immature-rat granulosa cells found rat, equine, human, porcine, ovine, bovine, and rabbit FSH to be effective (314) at culture concentrations of less than 10 ng/ml. This study is note¬ worthy because 32 hormones were examined, and only FSH from mammalian species was effective at stimulating PA activity. In contrast, Too et al. (469,470), in a similar sys¬ tem, i.e., PMSG-primed immature-rat ovarian granulosa cells in culture, found that 5 |xg/ml FSH and 5 |xg/ml relaxin stimulated PA activity after 20 hr of incubation. These work¬ ers also found that relaxin had no effect on cAMP levels of the granulosa cells but that FSH did, confirming the findings of Strickland and Beers (452). They also noted that, in response to relaxin, the granulosa cells released both latent and active collagenase as well as proteoglycanase. Although the latter enzyme may be released, collagenase has been identified in follicular fluid only by Espey and Rondell (158,159) and Okazaki et al. (362) (vide supra). The hormonal treatment of rats prior to collection of their ovaries and culture of the granulosa cells determines the effectiveness with which LH can elicit a PA response. The granulosa cells of hypophysectomized or intact immature rats treated with diethylstilbestrol (DES) are unaffected by LH but are highly responsive to FSH (252,298,351,496). Subsequent to incubation with FSH, the undifferentiated

474

/ Chapter 12

granulosa cells become responsive to LH (351,496). The granulosa cells obtained from PMSG-primed immature rats produce PA in response to addition of FSH to the culture medium after a 2-hr induction period, but in response to LH only after a longer lag period (92). However, whole proestrous preovulatory follicles cultured for 6 hr in the presence of LH, FSH, or prolactin (PRL) are fivefold more responsive to LH than to FSH, and PRL is ineffective in inducing PA production (411). Because an early response of the preovulatory follicle to LH is increased steroid synthesis, and blockade of steroid synthesis prevents ovulation, a relationship between steroids and PA production has been examined. Cyanoketone in¬ duces suppression of plasminogen production in the pre¬ ovulatory ovaries of immature rats primed with PMSG/hCG, and progesterone reverses this effect (355). Granulosa cell cultures, however, fail to show enhanced PA production when exposed to steroids (298,452), but, although proes¬ trous preovulatory follicles incubated with steroids (preg¬ nenolone, progesterone, testosterone, estradiol) fail to show enhanced follicular PA, addition of estradiol to culture me¬ dium containing LH-stimulated follicles further enhances PA production. In this same model, aminoglutethimide phosphate inhibits LH-stimulated PA production, and the inhibition is reversed by addition of progesterone, testos¬ terone, or estradiol but not by dihydrotestosterone (411). An explanation for these effects, though tentative, is that the steroid cascade is involved in the production of PA. Hypotheses on the production of two types of plasmin¬ ogen activator by the follicle and their relationship to ovu¬ lation have arisen as extensions of the early observation of the presence of PA in the follicle (42). Criticisms of the plasminogen-activator-plasminogen hypothesis (152) are that tissue-type plasminogen activator requires fibrin to be ac¬ tivated, that fibrinogen is present in follicular fluid in lower concentrations than in plasma, and that anticoagulant activ¬ ity is present in follicular fluid (242,381,447). However, urokinase-type plasminogen activator does not have these limitations. Analysis of the two types of plasminogen ac¬ tivator produced by the major component of the preovulatory follicle, the granulosa cells, by means of antibodies specific for each, indicates that the tissue-type enzyme is produced predominantly; however, whole follicles produce primarily the urokinase-type plasminogen activator (91). Unstimulated, undifferentiated granulosa cells secrete the urokinase-type plasminogen activator, and addition of FSH induces the secretion of tissue-type enzyme as indicated by sodium dodecyl sulfate polyacrylamide gel electrophoresis (351). After 2 days of priming with low concentrations of FSH, either FSH or LH induces the secretion of the tissuetype plasminogen activator. Thus, either gonadotropin in¬ duces the formation of tissue-type plasminogen activator in differentiated granulosa cells. That the production of plasminogen activator represents induction of protein synthesis in granulosa cells is indicated

by the response to the presence of protein-synthesis inhib¬ itors in the culture medium. FSH induces cellular plasmin¬ ogen activator, as measured by lysis of a chromogenic sub¬ strate, within 2 hr in undifferentiated granulosa cells; within 6 hr, 80% of maximal enzyme activity i)S contained in the cells; negligible plasminogen activator activity is released into the culture medium. Both actinomycin D and cycloheximide suppress cellular plasminogen-activator produc¬ tion when added to the culture medium early in the incu¬ bation period, and reduced secretion occurs when they are added at 44 to 48 hr of the incubation (252). The formation of plasmin or other proteases in the pre¬ ovulatory follicle has also been evaluated. Fibrinolytic ac¬ tivity in the external area of the preovulatory follicle reaches a maximum about 2 hr before ovulation in the rat (12), and rrans-aminomethyl cyclohexane carboxylic acid (t-AMCHA), an inhibitor of fibrinolysis, suppresses the appear¬ ance of fibrinolytic activity, decreases the appearance of ova, and postpones the time of ovulation. t-AMCHA ad¬ ministered to PMSG/hCG-treated immature rats or to proes¬ trous rats suppresses ovulation almost completely (12,13). Treatment with protease inhibitor also reduces the incidence of ovulation, suggesting that nonspecific proteolytic en¬ zymes may also be involved (220,392,413). Serine protease inhibitors or indomethacin block ovula¬ tion without inhibition of ovarian plasminogen activator, and only at high doses of indomethacin is blockade of both ovarian plasminogen-activator production and ovulation achieved (163,439); however, Reich et al. (411) found that indomethacin had no effect on production of plasminogen activators. The type of plasminogen activator is probably the key determinant with regard to plasminogen-activator production and ovulation. Indomethacin added to follicles in vitro prevents the LH-stimulated increase in plasminogen activator; however, when indomethacin is administered in vivo to PMSG/hCG-primed rats and the preovulatory fol¬ licles and granulosa cells are cultured, the situation is strik¬ ingly changed. At doses of indomethacin that induce block¬ ade of ovulation, the granulosa cells still produce and secrete tissue-type plasminogen activator, but whole-follicle pro¬ duction of urokinase-type plasminogen activator is reduced to basal levels (Table 14) (92). Correlated with this last observation are the reports that prostaglandins reverse the effect of indomethacin, that PGE2 is more involved in reversal of ovum maturation and cu¬ mulus mucification (138), and that PGF2a is more involved in the reversal of the indomethacin-imposed blockade of ovulation (135,136,426). Antiserum to PGE2 is a less ef¬ fective inhibitor of ovulation than is antiserum to PGF2a (26,275). Fibroblast proliferation is stimulated by PGF2a (125,211), and both prostaglandins stimulate fibroblasts to produce latent collagenase (123,139,300,398). Latent collagenase thus may be activated by urokinase, by plasmin¬ ogen activator, by the plasmin generated by the PA, or by fibrobasts.

Mammalian Ovulation / TABLE 14. Effect of indomethacin or ovarian PA production and ovulation in rats treated with hCGa Indomethacin (mg) 0 1 2 5

Granulosa tPA 37.2 57.4 19.2 17.6

± 26.2 ±11.4 ± 24.3 ± 11.4

Follicular fluid tPA

No. Follicular ovulating/No. uPA treated

2.4 4.0 2.3 1.9

5.0 2.4 1.0 0.6

± ± ± ±

1.4 0.6 3.8 0.8

± ± ± ±

3.7 3.4 1.0 0.3

15/20 6/19 3/20 1/15

aPMSG-primed 26-day-old rats treated with hCG and in¬ domethacin solvent or hCG and indomethacin. The data are expressed as a percentage of solubilized substrate. For ex¬ perimental details, see original report. Modified from Canipari and Strickland (92).

Collagenolysis Injection of collagenolytic enzyme (bacterial collagenase) into preovulatory follicles initiates the preovulatory changes leading to rupture of the follicle but at an accelerated pace (157). Early observations of collagenolytic activity in ova¬ ries (158,159,362) using the synthetic collagen substrate Ncarbobenzoxy-Gly-Pro-Gly-Gly-Pro-Ala were invalidated when it was noted that this substrate is not attacked by true mammalian collagenase (327). Collagenolytic activity in rat preovulatory follicles was demonstrated with a microassay; its presence in a latent form was noted, as was its increase in activity on treatment of immature rats with PMSG/hCG (327). Collagenolytic activity in the rabbit ovary (246,247) and in the human ovary was studied with a collagenase-specific peptide, and three other noncollagenolytic peptidases were studied with other synthetic peptides (179). All the pepti¬ dases had a marked preovulatory decrease in activity at the apex of the follicle. The activity in the granulosa cell layer increased near the time of ovulation (179), suggesting a role for collagenase in ovulation. In a study of rat-follicle col¬ lagenolytic activity, in which Type I collagen was used as the substrate and disappearance of collagen was determined by disappearance of hydroxyproline, collagen disappeared from the follicle and collagenase activity, activated with trypsin or aminophenylmercuric acetate, was present up to the time of ovulation. The presence of collagenolytic activity at the apex of the follicle (and that it is responsible for breakdown of the follicular collagen) was not, however, demonstrated (328). Collagenolytic activity in the ovaries of immature rats primed with PMSG/hCG rises to a peak at 8 hr after the injection of the hCG and remains high at 12 hr, correlating with maximal release of ova (120). Collagenolysis was also demonstrated with two different methods by Reich et al. (412). These workers labeled ovaries with L[3H]proline,

475

introduced into the ovarian bursa, and observed the disap¬ pearance of [3H]hydroxyproline from ovaries of adult rats through the estrous cycle, after nembutal was administered at proestrus (blocking the LH surge) or in response to ad ministered hCG; this effect was also observed in immature rats treated with PMSG/hCG. Excellent correlation was ob¬ tained between increased disappearance of labeled amino acid and either endogenous LH surge or exogenous hCG. That collagenase was the probable enzyme involved was also demonstrated by blockade of ovulation by introduction of cysteine (blockade of metalloprotease) into the ovarian bursa of immature rats treated with PMSG/hCG. The in¬ vestigators also demonstrated that the addition of p-ami¬ nophenylmercuric acetate or plasmin to [3H]proline-labeled collagen extracts of preovulatory ovaries from adult rats, or from PMSG/hCG, induced high levels of collagenolytic activity, as demonstrated by solubilization of [3H]hydroxyproline. These data further support the hypotheses that ovulation is dependent on activation of latent collagenase and that plasmin is probably the enzyme involved in its activation. Utilizing a variety of proteinase inhibitors, Ichi¬ kawa et al. (220) found that two different proteolytic en¬ zymes are involved in the blockade of ovulation in explanted hamster preovulatory follicles. Talopeptin, a metallopro¬ teinase inhibitor, inhibits mammalian collagenase and blocks ovulation (221), and microbial alkaline proteinase inhibitor blocks ovulation at the late stage of the ovulatory process, indicating that other proteinases are probably involved in further degradation of the telopeptide-free collagen formed by collagenase. These observations are summarized in a five-step cascade leading to the degradation of the collagen in the follicle wall and its rupture (Fig. 5). The smooth muscle in the wall may contribute to the expulsion of the antral contents subsequent to the degradative changes.

SUMMARY Ovulation is initiated by the surge of LH (431). Almost immediately the preovulatory follicle becomes hyperemic and subsequently edematous, even in the presence of in¬ domethacin, while progesterone and other steroid syntheses persist unaffected. Protein synthesis (17,32,403,404) ini¬ tiated by LH is responsible for cellular differentiation of the membrana granulosa to lutein cells and secretion of steroids and plasminogen activator (42,298,349,351). Activation of adenylate cyclase initiates many of these responses, includ¬ ing prostaglandin secretion (109,269,270,281,476). The theca interna also responds to LH stimulation by increased secre¬ tion of progesterone (257) and androgens (147) as well as prostaglandins (11,40,388) and plasminogen activator (252). The secretion of other substances by the follicle probably also occurs, but these are as yet uncharacterized. The ad¬ renergic neurons in the follicle wall are activated either by the LH or neurogenically and secrete norepinephrine. His-

476

/ Chapter 12

tamine released from mast cells (264,291,461) and the aadrenergic agonist effects may be to enhance the hyperemia (513,523) by affecting the contractility of the endothelial cells, the pericytes, and the postcapillary venules (192,321,349,364). The ^-adrenergic agonist effect may be to enhance secretion of progesterone (245,525). The pros¬ taglandins (42,92,411) increase plasminogen-activator pro¬ duction, as does progesterone. The enhanced secretion of plasminogen activator (13) then converts the plasminogen

in the follicular fluid and extracellular edema fluid to plasmin (43); the latter acts on latent collagen attached to the collagen fibers (343). Induced collagenolysis (221) and serine pro¬ teases (220) then complete the proteolysis of the collagen (120,327,328,362,412). The net effect is to decrease the tensile strength of the follicle wall to the point at which rupture occurs under the existing intrafollicular pressure of 15 to 20 mm Hg (56,65) (Fig. 6).

Latent collagenase

Active collagenase

Collagen

vk

Telopeptide-free collagen

Serine proteases

Follicle rupture

FIG. 6. Multifactor hypothesis of ovulation (see text of “Summary”).

Mammalian Ovulation

ACKNOWLEDGMENTS The references contained in this review extend only to early 1986 and are far from inclusive. There are many re¬ ports omitted largely because I am unaware of them or because they were peripheral to the areas reviewed. To those omitted, I apologize. I wish to thank my wife Janet for her forebearance. Spe¬ cial notes of appreciation go to Linda Mathews, who plowed through my rough drafts and made the writing and com¬ pletion possible, and to Dr. Anne Thistle for her editorial efforts.

REFERENCES 1. Abel, W., and Mcllroy, A. L. (1912): The arrangement and distri¬ bution of nerves in certain mammalian ovaries. Proc. R. Soc. Med. Obstet. Gynecol., 6:240-247. 2. Abramson, S., Korchak, H., Ludewig, R., Edelson, H., Haines, K., Levin, R. I., Herman, R., Rider, L., Kimmel, S., and Weissmann, G. (1985): Modes of action of aspirin-like drugs. Proc. Natl. Acad. Sci. USA, 82:7227-7231. 3. Adashi, E. Y., and Hsueh, A. J. W. (1981): Stimulation of beta-2adrenergic responsiveness by follicle stimulating hormone in rat gran¬ ulosa cells in-vitro and in-vivo. Endocrinology, 108:2170-2178. 4. Aeby, C. (1859): Uber glatten Muskelfassen in Ovarium und Mesovarium von Wirbelthieren. Arch. Anat. Physiol. Wissensch. Med., 675-676. 5. Aeby, C. (1861): Die glatten Muskelfassen in den Eierstocken der Wirbelthiere. Arch. Anat. Physiol. Wissensch. Med., 1:635-645. 6. Aguado, L. I., and Ojeda, S. R. (1984): Pre-pubertal ovarian function is finely regulated by direct adrenergic influences: Role of noradren¬ ergic innervation. Endocrinology, 114:1845-1853. 7. Aguado, L. I., and Ojeda, S. R. (1984): Ovarian adrenergic nerves play a role in maintaining preovulatory steroid secretion. Endocri¬ nology , 114:1944-1946. 8. Ahren, K., Janson, P. O., and Selstam, G. (1974): Search for arterio¬ venous shunts in the rabbit ovary in situ using perfusion of micro¬ spheres. J. Reprod. Fertil., 41:133-142. 9. Ainsworth, L., Baker, R. D., and Armstrong, D. T. (1975): Pre¬ ovulatory changes in follicular fluid prostaglandin F levels in swine. Prostaglandins, 9:915-925. 10. Ainsworth, L., Tsang, B. K., Downey, B. R., Baker, R. D., Marcus, T. J., and Armstrong, D. T. (1979): Effects of indomethacin on ovulation and luteal function in gilts. Biol. Reprod., 21:401-412. 11. Ainsworth, L., Tsang, B. H., Marcus, G. J., Downey, B. R. (1984): Prostaglandin production by dispersed granulosa and theca interna cells from porcine preovulatory follicles. Biop. Reprod., 31:115— 121. 12. Akazawa, K., Matsuo, O., Kosugi, T., Mihara, H., and Mori, N. (1983): The role of plasminogen activator in ovulation. Acta Physiol. Lot. Am., 33:105-110. 13. Akazawa, K., Mori, N., Kosugi, T., Matuso, O., and Mihara, H. (1983): Localization of fibrinolytic activity in ovulation of the rat follicle as determined by the fibrin slide method. Jpn. J. Physiol., 33:1011-1018. 14. Albertini, D. F., and Anderson, E. (1974): The appearance and structure of intercellular connections during the ontogeny of the rabbit ovarian follicle with particular reference to gap junctions. J. Cell Biol., 63:234-250. 15. Albertini, D. F., Fawcett, D. W., and Olds, P. J. (1975): Morpho¬ logical variation in gap junctions of ovarian granulosa cells. Tissue Cell, 7:389-405. 16. Albrechtsen, O. K. (1957): The fibrinolytic activity of human tissues. Br. J. Haematol., 3:284-291. 17. Alieva, J. J., Bonventre, P. F., and Lamanna, C. (1979): Inhibition of ovulation in hamsters by the protein synthesis inhibitors diphtheria toxin and cycloheximide. Proc. Soc. Exp. Biol. Med., 162:170-174.

/

477

18. Amenta, F., Allen, D. J., Didio, L. J. A., and Motta, P. (1979): A transmission electronmicroscopic study of smooth muscle cells in the ovary of rabbits, cats, rats, and mice. J. Submicrosc. Cytol., 11:3952. 19. Amenta, F., Cavallotti, C., De Rossi, M., and Evangelisti, F. (1981): Acetylcholinesterase containing nerve fibers in guinea-pig ovary. J. Neural Transm., 52:295-302. 20. Amman, K. (1936): Histologie des Schweine-Eierstockes unter besonderer Beruksichtigung des Ovarialzylus. Druck von Humber & Co. Aktiengesellschaft, Fravenfeld. 21. Amsterdam, A., Koch, Y., Lieberman, M. E., and Lindner, H. L. (1975): Distribution of binding sites for human chorionic gonado¬ tropin in the preovulatory follicle of the rat. J. Cell. Biol., 67:894902. 22. Andersen, M. M., Kroll, J., Byskov, A. G., and Faber, M. (1976): Protein composition in the fluid of individual bovine follicles. J. Reprod. Fertil., 48:109-118. 23. Anderson, E. (1979): Follicular morphology. In: Ovarian Follicular Development, edited by A. R. Midgley, Jr., and W. A. Sadler, pp. 91-105. Raven Press, New York. 24. Apkarian, R., and Curtis, J. C. (1981): SEM cryofracture study of ovarian follicles of immature rats. Scan. Electron. Miscrosc., 4:165172. 25. Armstrong, D. T., and Grinwich, D. L. (1972): Blockade of spon¬ taneous and LH-induced ovulation in rats by indomethacin, an in¬ hibitor of prostaglandin biosynthesis. Prostaglandins, 1:21-28. 26. Armstrong, D. T., Grinwich, D. L., Moon, Y. S., and Zanecnik, J. (1974): Inhibition of ovulation in rabbits by intrafollicular injection of indomethacin and PGF2 anti-serum. Life Sci., 14:129-140. 27. Armstrong, D. T., Moon, Y. S., and Zanecnik, J. (1974): Evidence for a role of ovarian prostaglandins in ovulation. In: Gonadotropins and Gonadal Function, edited by N. R. Moudgal, pp. 345-356. Academic Press, New York. 28. Astrup, T., and Permin, P. M. (1947): Fibrinolysis in the animal organism. Nature, 159:681-682. 29. Ax, R. L., and Ryan, R. J. (1978): The porcine ovarian follicle. IV. Mucopolysaccharides at different stages of development. Biol. Re¬ prod., 20:1123-1132. 30. Ax, R. L., and Ryan, R. J. (1979): FSH stimulation of ^-glucos¬ amine-incorporation into proteoglycans by porcine cells in vitro. J. Clin. Endocrinol. Metab., 49:646-648. 31. Bagavandoss, P., Midgley, A. R., Jr., and Wicha, M. (1983): De¬ velopmental changes in the ovarian follicular basal lamina detected by immunofluorescence and electron microscopy. J. Histochem. Cytochem., 31:633-640. 32. Bahr, J., Kao, L., and Nalbandov, A. V. (1974): The role of cate¬ cholamines and nerves in ovulation. Biol. Reprod., 10:273-290. 33. Baird, D. T. (1983): Factors regulating the growth of the preovulatory follicle in the sheep and human. J. Reprod. Fertil., 69:343-352. 34. Balboni, G. C. (1983): Structural changes: Ovulation and luteal phase. In: The Ovary, edited by G. B. Serra, pp. 123-141. Raven Press, New York. 35. Ball, G. D., Beilin, M. E., Ax, R. L., and First, N. L. (1982): Glycosaminoglycans in bovine cumulus-oocyte complexes: Mor¬ phology and chemistry. Mol. Cell. Endocrinol., 28:113-122. 36. Baranczuk, R. J., and Fainstat, T. (1976): Progesterone-induced ovulation of the hamster ovary in vitro. J. Endocrinol., 70:317-318. 37. Baranczuk, R. J., and Fainstat, T. (1976): In vitro ovulation from adult hamster ovary. Am. J. Obstet. Gynecol., 124:517-522. 38. Barros, C., and Austin, C. R. (1968): Inhibition of ovulation by systemically administered actinomycin D in the hamster. Endocri¬ nology, 83:177-179. 39. Bauminger, S., Lieberman, M. E., and Lindner, H. R. (1975): Ste¬ roid-independent effect of gonadotropins on prostaglandin synthesis in rat graafian follicles in vitro. Prostaglandins, 9:753-764. 40. Bauminger, S., and Lindner, H. R. (1975): Periovulatory changes in ovarian prostaglandin formation and their hormonal control in the rat. Prostaglandins, 9:737-751. 41. Beck, L. R., and Shelden, R. M. (1972): Antigenicity of rat follicular fluid. Fertil. Steril., 23:910-914. 42. Beers, W. H. (1975): Follicular plasminogen and plasminogen ac¬ tivator and the effect of plasmin on ovarian follicle wall. Cell, 6:379386.

478

/ Chapter 12

43. Beers, W. H., Strickland, S., and Reich, E. (1975): Ovarian plas¬ minogen activator: Relationship to ovulation and hormonal regula¬ tion. Cell, 6:387-394. 44. Beers, W. H., and Strickland, S. (1978): A cell culture assay for follicle-stimulating hormone. J. Biol. Chem., 253:3877-3881. 45. Beilin, M. E., and Ax, R. L. (1984): Chondroitin sulfate: An in¬ dicator of atresia in bovine follicles. Endocrinology, 114:428-434. 46. Beilin, M. E., Hinshelwood, M. M., Robinson, G. M., Ax, R. L., and Hauser, E. R. (1983): Estrogen, heparan sulfate, and chondroitin sulfate in relation to morphology of individual bovine follicles. In: Factors Regulating Ovarian Function, edited by G. S. Greenwald and P. F. Terranova, pp. 45-48. Raven Press, New York. 47. Beilin, M. E., Lenz, R. W., Steadman, L. E., and Ax, R. L. (1983): Proteoglycan production by bovine granulosa cells in vitro occurs in response to FSH. Mol. Cell. Endocrinol., 29:51-65. 48. Bjersing, L. (1978): Maturation, morphology, and endocrine function of the follicular wall in mammals. In: The Vertebrate Ovary, edited by R. E. Jones, pp. 181-214. Plenum Press, New York. 49. Bjersing, L., and Cajander, S. (1974): Ovulation and the mechanism of follicle rupture. I. Light microscopic changes in rabbit ovarian follicles prior to induced ovulation. Cell Tissue Res., 149:287-300. 50. Bjersing, L., and Cajander, S. (1974): Ovulation and the mechanism of follicle rupture. II. Scanning electron microscopy of rabbit ger¬ minal epithelium prior to induced ovulation. Cell Tissue Res., 149:301— 312. 51. Bjersing, L., and Cajander, S. (1974): Ovulation and the mechanism of follicle rupture. III. Transmission electron microscopy of rabbit germinal epithelium prior to induced ovulation. Cell Tissue Res., 149:313-327. 52. Bjersing, L., and Cajander, S. (1974): Ovulation and the mechanism of follicle rupture IV. Ultrastructure of membrana granulosa of rabbit graafian follicles prior to induced ovulation. Cell Tissue Res., 153:114. 53. Bjersing, L., and Cajander, S. (1974): Ovulation and the mechanism of follicle rupture. V. Ultrastructure of tunica albuginea and theca externa of rabbit graafian follicles prior to induced ovulation. Cell Tissue Res., 153:15-30. 54. Bjersing, L., and Cajander, S. (1974): Ovulation and the mechanism of follicle rupture. VI. Ultrastructure of theca interna and the inner vascular network surrounding rabbit graafian follicles prior to induced ovulation. Cell Tissue Res., 153:31-44. 55. Bjorkman, N. (1962): A study of the ultrastructure of the granulosa cells of the ovary. Acta Anat., 51:125-147. 56. Blandau, R. J., and Rumery, R. E. (1963): Measurements of intrafollicular pressure in the ovulatory and preovulatory follicles of the rat. Fertil. Steril., 14:330-341. 57. Blasco, L., Wu, C. H., Flickinger, G. L., Pearlmutter, D., and Mikhail, G. (1975): Cardiac output and genital blood flow distribution during the preovulatory period in rabbits. Biol. Reprod., 13:581— 586. 58. Bodkhe, R. R., and Harper, M. J. K. (1972): Changes in the amount of adrenergic neurotransmitter in the genital tract of untreated rabbits and rabbits given reserpine or ipronizid during the time of egg trans¬ port. Biol. Reprod., 6:288-299. 59. Bohmfalk, J. F., and Fuller, G. M. (1980): Plasminogen is synthe¬ sized by primary cultures of rat hepatocytes. Science, 209:408-410. 60. Bomsel-Helmrich, O., Gougeon, A., Thebault, A., Saltarelli, D., Milgron, E., Frydman, R., and Papiemik, E. (1979): Healthy and artretic human follicles in the preovulatory phase: Differences in evolution of follicular morphology and steroid content of follicular fluid. J. Clin. Endocrinol. Metab., 48:686-694. 61. Bostrom, H., and Odeblad, O. (1952): Autoradiographic observation on the uptake of S35 in the genital organs of the female rat and rabbit after injection of labelled sodium sulphate. Acta Endocrinol. (Copenh.), 10:89-96. 62. Brace, R. A. (1981): Progress toward resolving the controversy of positive vs negative interstitial fluid pressure. Circ. Res., 49 281297. 63. Brill, W. (1915): Untersuchungen uber die Nerven des Ovariums. Arch. Mikr. Anat., 86(Sect. l):338-376. 64. Brink, C. E., and Grob, H. S. (1973): Response of the denervated mouse ovary to exogenous gonadotropins. Biol. Reprod., 9:108 (ab¬ stract No. 120).

65. Bronson, R. A., Bryant, G., Balk, M., Emanuels, N. (1979): Intrafollicular pressure within preovulatory follicles of the pig. Fertil. Steril., 31:205-213. 66. Bullock, D. W., and Kappauf, B. H. (1973): Dissociation of go¬ nadotropin-induced ovulation and steroidogenesis in immature rats. Endocrinology, 92:1625-1628. 67. Bulmer, D. (1965): A histochemical study of ovarian cholinesterases. Acta Anat., 62:254-265. 68. Burden, H. W. (1972): Ultrastructural observations on ovarian peri¬ follicular smooth muscle in the cat, guinea pig, and rabbit. Am. J. Anat., 133:125-142. 69. Burden, H. W. (1972): Adrenergic innervation in ovaries of the rat and guinea pig. Am. J. Anat., 133:455-462. 70. Burden, H. W. (1973): The distribution of smooth muscle in the cat ovary with a note on its adrenergic innervation. J. Morphol., 140:467476. 71. Burden, H. W. (1978): Ovarian innervation. In: The Vertebrate Ovary, edited by R. E. Jones, pp. 615-638. Plenum Press, New York. 72. Burden, H. W., and Lawrence, I. E. (1977): The effects of de¬ nervation on the localization of A5-3(3-hydroxysteroid dehydrog¬ enase activity in the rat ovary during pregnancy. Acta Anat., 97: 286-290. -73. Burden, H. W., and Lawrence, I. E., Jr. (1977): The effect of denervation on compensatory ovarian hypertrophy. Neuroendocri¬ nology, 23:368-378. 74. Burden, H. W., and Lawrence, I. E., Jr. (1978): Experimental studies on the acetylcholinesterase positive nerves in the ovary of the rat. Anat. Rec., 190:233-242. 75. Burden, H. W., and Lawrence, I. E. (1980): Nerve supply of the ovary. In: Biology of the Ovary, edited by P. M. Motta and E. S. E. Hafez, pp. 99-105. Martinus Nijhoff, The Hague. 76. Burghardt, R. C., and Anderson, E. (1979): Hormonal modulation of ovarian interstitial cells with particular reference to gap junctions J. Cell Biol., 81:104-114. 77. Burghardt, R. C., and Anderson, E. (1981): Hormonal modulation of gap junctions in rat ovarian follicles. Cell Tissue Res., 214:181 — 194. 78. Burghardt, R. C., and Matheson, R. L. (1982): Gap junction am¬ plification in rat ovarian granulosa cells. Dev. Biol., 94:206-215. 79. Burgoyne, P. S., Borland, R. M., Biggers, J. D., and Lechene, C. P. (1979): Elemental composition of rabbit control fluid during preovulatory follicular swelling. J. Reprod. Fertil., 57:515-523. 80. Bum, J. H. (1961): A new view of adrenergic nerve fibers explaining the action of reserpine, bretylium and guanethidine. Br Med J 1:1623-1627. 81. Bum, J. H. (1966): Adrenergic transmission introductory remarks. Pharmacol. Rev., 18:459-470. 82. Bum, J. H., and Rand, M. J. (1965): Acetylcholine in adrenergic transmission. Annu. Rev. Pharmacol., 5:163-182. 83. Bumstock, G. (1986): The changing face of autonomic neurotrans¬ mission. Acta Physiol. Scand., 126:67-92. 84. Burr, J. H., and Davis, J. R. (1951): The vascular system of the rabbit ovary and its relationship to ovulation. Anat. Rec., 111:273297. 85. Bushmeyer, S. M., Beilin, M. E., Brantmeier, S. A., Boehm, S. K., Kubajak, C. L., and Ax, R. L. (1985): Relationships between bovine follicular fluid glycosaminoglycans and steroids. Endocri¬ nology, 117:879-885. 86. Byskov, A. G. S. (1969): Ultrastructural studies on the preovulatory follicle in the mouse ovary. Z. ZellForsch. Mikroskop. Anat., 100 285299. 87. Cajander, S. (1976): Structural alterations of rabbit ovarian follicles after mating with special reference to the overlying surface epithe¬ lium. Cell Tissue Res., 173:437-449. 88. Cajander, S., and Bjersing, L. (1975): Fine structural demonstration of acid phosphatase in rabbit germinal epithelium prior to induced ovulation. Cell Tissue Res., 164:279-289. 89. Cajander, S., and Bjersing, L. (1976): Further studies of the epi¬ thelium covering preovulatory rabbit follicles with special reference to lysosomal alterations. Cell Tissue Res., 169:129-141. 90. Canipari, R., and Strickland, S. (1985): Hormonal regulation of plasminogen activator in the rat ovary. Biol. Reprod., 32(Suppl. 1): 183 (abstract No. 292).

Mammalian Ovulation

91. Canipari, R., and Strickland, S. (1985): Plasminogen activator in the rat ovary. Production and gonadotropin regulation of the enzyme in granulosa and thecal cells. J. Biol. Chem., 260:5121-5125. 92. Canipari, R., and Strickland, S. (1986): Studies on the hormonal regulation of plasminogen activator production in the rat ovary. En¬ docrinology, 118:1652-1659. 93. Capps, M. L., Lawrence, I. E., Jr., and Burden, H. W. (1981): Cellular functions in perifollicular contractile tissue of the rat ovary during the preovulatory period. Cell Tissue Res., 219:133-141. 94. Caravaglios, R., and Cilotti, R. (1957): A study of the proteins in the follicular fluid of the cow. J. Endocrinol., 15:273-278. 95. Casey, F. B., Appleby, B. J., and Buck, D. C. (1983): Selective inhibition of lipoxygenase metabolic pathway of arachodonic acid by the SRS-A antagonist, FPL 55712. Prostaglandins, 25:1-11. 96. Castren, O., Airaksinen, M. M., Fristrom, S., and Saarikoski, S. (1973): Decrease of litter size and fetal monoamines by 6-hydroxydopamine in mice. Experientia, 29:576-578. 97. Caucig, H., Friedrich, F., Breitenecker, G., and Golob, E. (1972): Enzyme activity in the fluid of the human ovarian follicle. Gynecol. Obstet. Invest., 3:215-220. 98. Caucig, PL, Friedrich, F., Hager, R., and Golob, E. (1971): Enzym untersuchungen in Follikel- und Zystenflussigkeit des Menchlichen Ovars. Acta Endocrinol. (Copenh.) (Suppl.), 152:52 (abstract No. 52). 99. Cerletti, P., and Zichella, L. (1961): Nucleotidases, nucleotides, vitamins and coenzymes in the follicular fluid of human ovary. Clin. Chim. Acta, 6:581-582. 100. Challis, J. R. G., Erickson, G. F., and Ryan, K. J. (1974): Pros¬ taglandin F production in vitro by granulosa cells from rabbit pre¬ ovulatory follicles. Prostaglandins, 7:183-193. 101. Channing, C. P., Schaerf, F. W., Anderson, L. D., and Tsafriri, A. (1980): Ovarian follicular and luteal physiology. In: Reproductive Physiology III, International Review of Physiology, Vol. 22, edited by R. O. Greep, pp. 117-201. University Park Press, Baltimore. 102. Channing, C. P., Tanabe, K., Chacon, M., and Tildon, J. T. (1984): Stimulatory effects of FSH and luteinizing hormone upon secretion of progesterone and inhibin activity by cultured infant human ovarian granulosa cells. Fertil. Steril., 42:598-605. 103. Channing, C. P., and Tsafriri, A. (1977): Mechanism of action of luteinizing hormone and follicle-stimulating hormone on the ovary in vitro. Metabolism, 26:413-468. 104. Chemey, D. D., Didio, L. J. A., and Motta, P. (1975): The devel¬ opment of rabbit ovarian follicles following copulation. Fertil. Steril., 26:251-211. 105. Chihal, H. J. W., Weitsen, H. A., Stone, S. C., and Peppier, R. D. (1976): Autonomic innervation and plasma estradiol-17 and progesterone levels in rats with subcutaneous ovarian autografts. Cell Tissue Res., 175:113-121. 106. Christiansen, J. A., Jensen, C. E., and Zachariae, F. (1958): Studies on the mechanism of ovulation. Some remarks on the effect of de¬ polymerization of high-polymers on the preovulatory growth of fol¬ licles. Acta Endocrinol., 29:115-117. 107. Claesson, L. (1947): Is there any smooth musculature in the wall of the Graafian follicle? Acta Anat., 3:295-311. 108. Clark, J. G. (1899): Origin, development and degeneration of the blood vessels of the ovary. Bull. J. Hopkins Hosp., 94, 95, 96:4044. 109. Clark, M. R., Marsh, J. M., and Lemaire, W. J. (1978): Stimulation of prostaglandin accumulation in preovulatory rat follicles by aden¬ osine 3'-5' monophosphate. Endocrinology, 102:39-44. 110. Clinton, M., Long, W. F., Williamson, F. B., Hutchinson, J. S., and Seddon, B. (1983): Incorporation of 35S into glycosaminoglycans of ovarian follicular and luteal tissue isolated during the guinea pig estrous cycle. Biochem. Biophys. Res. Commun., 111:574-580. 111. Collen, D. (1980): On the regulation and control of fibrinolysis. Thromb. Haemost., 43:77-85. 112. Cook, B., Kaltenback, C. C., Niswender, G. D., Norton, H. W., and Nalbandov, A. V. (1969): Short-term ovarian responses to some pituitary hormones infused in vivo in pigs and sheep. J. Animal Sci., 29:711-718. 113. Coons, L. W., and Espey, L. L. (1977): Quantitation of nexus junctions in the granulosa cell layer of rabbit ovarian follicles during ovulation. J. Cell Biol., 74:321-325.

/

479

114. Comer, G. W. (1919): On the origin of the corpus luteum of the sow from both granulosa and theca interna. Am. J. Anat., 26:117183. 115. Comer, G. W. (1919): Cyclic changes in the ovaries and uterus of the sow, and their relation to the mechanism of implantation. In: Publications of the Carnegie Institute, No. 276 (contribution to Em¬ bryology, No. 64), pp. 119-145. 116. Coutinho, E. M., and Maia, H. (1971): The contractile response of the human uterus, fallopian tubes and ovary to prostaglandins in vivo. Fertil. Steril., 22:539-542. 117. Coutinho, E. M., Maia, H., and Maia, H., Jr. (1974): Ovarian contractility. Basic Life Sci., 4:127-137. 118. Cruikshank, W. C. (1797): Experiments in which on the third day after impregnation the ova of rabbits were found in the fal¬ lopian tubes, on the fourth after, in the uterus with the first appear¬ ances of the fetus. Philos. Trans. R. Soc. Lond., (Biol.), 87:197214. 119. Curry, T. E., Jr., Lawrence, I. E., Jr., and Burden, H. W. (1984): Ovarian sympathectomy in the guinea-pig. 2. Effects on follicular development during the prepubertal period and following exogenous gonadotropin stimulation. Cell Tissue Res., 236:593-596. 120. Curry, T. E., Dean, D. D., Woessner, J. F., Jr., and LeMaire, W. J. (1985): The extraction of a tissue collagenase associated with ovulation in the rat. Biol. Reprod., 33:981-991. 121. Daume, E., Chari, S., Hopkinson, C. R. N., Sturm, G., and Hirschhauser, C. (1978): Nachweis von Inhibin-aktivitat in der Follikelflussigkeit menschlicher Ovarien. Klin. Wochenschr., 56:369-370. 122. David, A., Frenkel, G., and Kraicer, P. F. (1973): Chemical com¬ position of rabbit follicular fluid. Fertil. Steril., 24:227-229. 123. Dayer, J. M., Krane, S. M., Russel, R. G. G., and Robinson, D. R. (1976): Production of collagenase and prostaglandins by iso¬ lated adherent rheumatoid synovial cells. Proc. Natl. Acad. Sci. USA, 73:945-949. 124. Deanesly, R. (1956): Cyclic function in ovarian grafts. J. Endo¬ crinol., 13:211-220. 125. DeAsua, L. J., Clingan, D., and Rudland, P. S. (1975): Initiation of cell proliferation in cultured mouse fibroblasts by prostaglandin F2c- Proc. Natl. Acad. Sci. USA, 12:2124-2128. 126. De La Cruz, A., Wright, K. H., and Wallach, E. E. (1976): The effects of cholinergic agents on ovarian contractility in the rabbit. Obstet. Gynecol., 47:272-278. 127. Dennefers, B., Tjugum, J., Norstrom, A., Janson, P. O., Nilsson, L., Hamberger, L., and Wilhelmsson, L. (1982): Collagen synthesis inhibition by PGE2 within the human follicular wall—One possible mechanism underlying ovulation. Prostaglandins, 24:295-302. 128. Desjardins, C., Kirton, K. T., and Hafs, H. D. (1966): Some chem¬ ical, immunochemical and electrophoretic properties of bovine fol¬ licular fluid. J. Reprod. Fertil., 11:237-244. 129. deVos, J. (1894): Etude sur Pinnervation de l’ovaire. Bull Acad. Med. Beige Ser. 4, 8:552-558. 130. Diaz-Infante, A., Jr., Wright, K. H., and Wallach, E. E. (1974): Effects of indomethacin and prostaglandin F2o on ovulation and ovar¬ ian contractility in the rabbit. Prostaglandins, 5:567-579. 131. Didio, L. J. A., Allen, D. J., Corner, S., and Motta, P. M. (1980): Smooth muscle in the ovary. In: Biology of the Ovary, edited by P. M. Motta and E. S. E. Hafez, pp. 107-118. Martinus Nijhoff, The Hague. 132. di Zerega, G. S., Goembelsmann, U., and Nakamura, R. M. (1982): Identification of protein(s) secreted by the preovulatory ovary which suppresses the follicle response to gonadotropins. J. Clin. Endocri¬ nol. Metab., 54:1091-1096. 133. Dominques, R., and Riboni, L. (1971): Failure of ovulation in autografted ovary of hemispayed rat. Neuroendocrinology, 7:164-170. 134. Douglas, W. W. (1980): Polypeptides—Angiotensin, plasma kinins, and others. In: The Pharmacological Basis of Therapeutics, 6th ed., edited by A. G. Gilman, L. S. Goodman, and A. Gilman, pp. 659663. Macmillan, New York. 135. Downey, B. R., and Ainsworth L. (1980): Reversal of indomethacin blockade of ovulation in gilts by prostaglandins. Prostaglandins, 19:17-22. 136. Downs, S. M., and Longo, F. J. (1982): Effects of indomethacin on preovulatory follicles in immature superovulated mice. Am. ./. Anat., 164:265-274.

480

/

Chapter 12

137. Downs, S. M., and Longo, F. J. (1983): Prostaglandins and pre¬ ovulatory follicular maturation in mice. J. Exp. Zool., 228:99-108. 138. Downs, S. M., and Longo, F. J. (1983): An ultrastructural study of preovulatory apical development in mouse ovarian follicles: Effects of indomethacin. Anat. Rec. 205:159-168. 139. Dowsett, M., Eastman, A. R., Easty, D. M., Easty, G. C., Powles, T. J., and Neville, A. M. (1976): Prostaglandin mediation of collagenase-induced bone resorption. Nature, 263:72-74. 140. Dunaif, A. E., Zimmerman, E. A., Friesen, H. G., and Frantz, A. G. (1982): Intracellular localization of prolactin receptors and prolactin in the rat ovary by immunocytochemistry. Endocrinology, 110:1465-1471. 141. Edwards, R. G. (1974): Follicular fluid. J. Reprod. Fertil., 37:189219. 142. Edwards, R. G., Steptoe, P. C., Fowler, R. E., and Baillie, J. (1980): Observations on preovulatory human ovarian follicles and their as¬ pirates. Br. J. Obstet. Gynaecol., 87:769-779. 143. Ellinwood, W. E., Nett, T. M., and Niswender, G. D. (1978): Ovarian vasculature: Structure and function. In: The Vertebrate Ovary: Comparative Biology and Evolution, edited by R. E. Jones, pp. 583— 416. Plenum Press, New York. 144. Ellis, S. (1961): Bioassay of luteinizing hormone. Endocrinology, 68:334-340. 145. Eppig, J. J. (1981): Prostaglandin E2 stimulates cumulus expansion and hyaluronic acid synthesis by cumuli oophori isolated from mice. Biol. Reprod., 25:191-195. 146. Erickson, G. F., and Hsueh, A. J. W. (1978): Secretion of “inhibin” by rat granulosa cells in vitro. Endocrinology, 103:1960-1963. 147. Erickson, G. F., Magoffin, D. A., Dyer, C. A., and Hofeditz, C. (1985): The ovarian androgen producing cells: A review of struc¬ ture/function relationships. Endocr. Rev., 6:371-399. 148. Espey, L. L. (1967): Ultrastructure of the apex of the rabbit Graafian follicle during the ovulatory process. Endocrinology, 81:267-276. 149. Espey, L. L. (1967): Tenacity of porcine Graafian follicle as it ap¬ proaches ovulation. Am. J. Physiol., 212:1397-1401. 150. Espey, L. L. (1971): Decomposition of connective tissue in rabbit ovarian follicles by multivesicular structures of thecal fibroblasts. Endocrinology, 88:437^144. 151. Espey, L. L. (1974): Ovarian proteolytic enzymes and ovulation. Biol. Reprod., 10:216-235. 152. Espey, L. L. (1978): Ovulation. In: The Vertebrate Ovary, edited by R. E. Jones, pp. 503-532. Plenum Press, New York. 153. Espey, L. L. (1980): Ovulation as an inflammatory reaction: a hy¬ pothesis. Biol. Reprod., 22:73-106. 154. Espey, L. L. (1986): Simultaneous determination of ovarian pros¬ taglandin E2, prostaglandin F2„, 6-keto-prostaglandin F,a, (3-estradiol and progesterone during ovulation in the PMSG/hCG-primed im¬ mature rat. Biol. Reprod., 34 (Suppl. 1), 151 (abstract). 155. Espey, L. L., and Coons, P. J. (1976): Factors which influence ovulatory degradation of rabbit ovarian follicles. Biol. Reprod., 14:233245. 156. Espey, L. L., and Lipner, H. (1963): Measurement of intrafollicular pressures in the rabbit ovary. Am. J. Physiol., 205:1067. 157. Espey, L. L., and Lipner, H. (1965): Enzyme-induced rupture of rabbit Graafian follicle. Am. J. Physiol., 208:208-213. 158. Espey, L. L., and Rondell, P. (1967): Estimation of mammalian collagenolytic activity with a synthetic substrate. J. Appl. Physiol., 23:757-761. 159. Espey, L. L., and Rondell, P. (1968): Collagenolytic activity in the rabbit and sow Graafian follicle during ovulation. Am. J. Physiol., 214:326-329. 160. Espey, L. L., and Stutts, R. H. (1972): Exchange of cytoplasm between two cells of the membrana granulosa in rabbit ovarian fol¬ licles. Biol. Reprod., 6:168-175. 161. Espey, L., Miller, D., and Margolius, H. S. (1985): Indomethacin and cycloheximide inhibition of ovarian kinin-generating activity during ovulation in the PMSG/hCG-primed immature rat. Biol. Re¬ prod., 32(Suppl. 1) (abstract No. 291). 162. Espey, L. L., Coons, P. J., Marsh, J. M., and LeMaire, W. J. (1981): Effect of indomethacin on preovulatory changes in the ul¬ trastructure of rabbit Graafian follicles. Endocrinology, 108:10401048. 163. Espey, L., Shimada, H., Okamura, H., and Mori, T. (1985): Effect of various agents on ovarian plasminogen activator activity during

ovulation in pregnant mare’s serum gonadotropin-primed immature rats. Biol. Reprod., 32:1087-1094. 164. von Euler, U. S., and Hammarstrom, S. (1937): Ueber Vorkommen und Wirkung^von Adrenalin in Ovarien. Scand. Arch. Physiol., 77:163178. 165. Evans, D. H. L., and Murray, J. G. (1954): Histological and func¬ tional studies on the fibre composition of the vagus nerve of the rabbit. J. Anat., 88:320-337. 166. Evans, G., Dobias, M., King, C. J., and Armstrong, D. T. (1983): Production of prostaglandins by porcine preovulatory follicular tis¬ sue and their roles in intrafollicular function. Biol. Reprod., 28:322328. 167. Eyre, D. R. (1980): Collagen: Molecular diversity in the body’s protein scaffold. Science, 207:1315-1322. 168. Falck, B. (1962): Observations on the possibilities of the cellular localization of monoamines by a fluorescence method. Acta Physiol. Scand., 56:1-25. 169. Falck, B., and Owman, C. (1965): A detailed methodological de¬ scription of the fluorescence method for the cellular demonstration of biogenic monoamines. Acta Univ. Lundensis Sect. II, 7:5-23. 170. Fessler, J. H., and Fessler, L. I. (1978): Biosynthesis of procollagen. Annu. Rev. Biochem., 47:129-162. J71. Fink, G., and Schofield, G. C. (1971): Experimental studies on the innervation of the ovary in cats. J. Anat., 109:115-126. 172. Fletcher, W. H. (1975): Assembly of an “enclosed gap junction” by the granulosa cells of developing ovarian follicles in sexually im¬ mature rats. J. Cell Biol., 67:116a(abstract no. 232). 173. Fletcher, W. H. (1979): Intercellular junctions in ovarian follicles: A possible functional role in follicle development. In: Ovarian Fol¬ licular Development and Function, edited by A. R. Midgley and W. A. Sadler, pp. 113-120. Raven Press, New York. 174. Fletcher, P. W., Dias, J. A., Sanzo, M. A., and Reichert, L. E., Jr. (1982): Inhibition of FSH action on granulosa cells by low mo¬ lecular weight components of follicular fluid. Mol. Cell. Endocrinol., 25:303-316. 175. Fletcher, W. H., and Everett, J. W. (1973): Ultrastructural reorgan¬ ization of rat granulosa cells on the day of proestrus. Anat. Rec., 175:320a. 176. France, E. S. (1969): Reversal by pargyline of reserpine block of induced ovulation—Direct ovarian effects. Neuroendocrinology, 6:7789. 177. Frankenhauser. R. (1867): Die Nerven der Gebarmuter und ihre Endegigungen in den glatten Muskelfasern. F. von Mauke, Jena. 178. Fraser, I., Baird, D. T., and Cockbum, F. (1973): Ovarian venous blood P02, PC02 and pH in women. J. Reprod. Fertil., 33:11-17. 179. Fukumoto, M., Yajima, Y., Okamura, H. and Midorikawa, O. (1981): Collagenolytic enzyme activity in human ovary: An ovulatory enzyme system. Fertil. Steril., 36:746-750. 180. Fukushimo, M. (1977): Intercellular junctions in the human devel¬ oping preovulatory follicle and corpus luteum. Int. J. Fertil., 22:206216. 181. Fumagalli, Z., Motta, P., and Calvieri, S. (1971): The presence of smooth muscular cells in the ovary of several mammals as seen under the electron microscope. Experientia, 27:682-683. 182. von Gawronsky, N. (1894): Ueber verbreitung und Endigung der Nerven in den weiblichen Genitalien. Arch. Gynecol., 47:271-283. 183. Gebauer, H., Lindner, H. R., and Amsterdam, A. (1978): Synthesis of heparin-like glycosamine-glycans in rat ovarian slices. Biol. Re¬ prod., 18:350-358. 184. Gelderd, J. G., and Peppier, R. D. (1979): Effect of spinal cord transection on the reproductive system in the female rat. Neuroen¬ docrinology, 29:293-299. 185. Gillet, J. Y., Maillet, R., and Gautier, C. (1980): Blood and lymph supply of the ovary. In: Biology of the Ovary, edited by P. M. Motta and E. S. E. Hafez, pp. 86-98. Martinus Nijhoff, The Hague. 186. Gilula, N. B., Epstein, M. L., and Beers, W. H. (1978): Cell-tocell communication and ovulation. A study of the cumulus-oocyte complex. J. Cell Biol., 78:58-75. 187. Gimeno, M. F., Borda, E., Sterin-Borda, L., Vidal, J. H., and Gimeno, A. L. (1976): Pharmacologic influences on human ovarian contractions. Obstet. Gynecol., 47:218-222. 188. Gimeno, M. F., Gimeno, A. L., and Rettori, V. B. (1975): Phys¬ iologic and pharmacologic studies on the motility of isolated guinea pig ovaries. Fertil. Steril., 26:422-426.

Mammalian Ovulation

189. Gimeno, M. F., Sterin-Speziale, N., Landa, A., Bonacossa, A., and Gimeno, A. L. (1977): Drug-induced motility of sow (Sus scrofa) Graafian follicles isolated during pre- and postovulatory periods of the sex cycle. Act Physiol. Lat. Am., 27:321—331. 190. Goldenberg, R. L., Vaitukaitis, J. L., and Ross, G. T. (1972): Estrogen and follicle stimulating hormone interactions on follicle growth in rats. Endocrinology, 90:1492-1498. 191. Gothie, S. (1954): Etude comparee de la repartition du P32 et du 35S dans l’organisme de lapire, specialement dans l’ovaire. C. R. Soc. Biol., 148:1210-1213. 192. Grega, G. J., Adamski, S. W., and Dobbins, D. E. (1986): Phys¬ iological and pharmacological evidence for the regulation of per¬ meability. Fed. Proc., 45:96-100. 193. Grenedai, I., Marchetti, B., Maugeri, S., Roxas, M. A., and Scapagnini, V. (1978): Prevention of compensatory ovarian hypertrophy by local treatment of the ovaries with 6-OHDA. Neuroendocrinology, 27:272-278. 194. Grimek, H. J., and Ax, R. L. (1982): Chromatographic compari¬ sons of chondroitin-containing proteoglycans from small and large bovine ovarian follicles. Biochem. Biophys. Res. Commun., 104: 1401-1406. 195. Grimek, H. J., Beilin, M. E., and Ax, R. L. (1984): Characteristics of proteoglycans isolated from small and large bovine ovarian fol¬ licles. Biol. Reprod., 30:397-409. 196. Grinwich, D. L., Kennedy, T. G., and Armstrong, D. T. (1972): Dissociation of ovulatory and steroidogenic actions of luteinizing hormone in rabbits with indomethacin, an inhibitor of prostaglandin biosynthesis. Prostaglandins, 1:89-96. 197. Grab, H. S. (1969): Effects of denervation of mouse ovary. Am. Zool, 9:1086 (abstract No. 141). 198. Grab, H. S. (1972): Effects of abdominal ovarian denervation on vaginal opening, estrus and ovarian histology. Fed. Proc., 31:265 (abstract No. 296). 199. Grab, H. S., and Brink, C. E. (1973): Effects of exogenous norepineprine on the ovaries of hypophysectemized mice. Fed Proc., 32:213 (abstract No. 10). 200. Grohe, F. (1863): Uber den Bau und das Wachstum des menschlichen Eierstocks. Arch. Pathol. Anat. Physiol. 26:271-306. 201. Guraya, S. S. (1985): Biology of Ovarian Follicles in Mammals. Springer-Verlag, Berlin. 202. Guttmacher, M. S., and Guttmacher, A. F. (1921): Morphological and physiological studies on the musculature of the mature Graafian follicle of the sow. Bull. J. Hopkins Hosp., 32:394-399. 203. Guyton, A. C. (1986): Textbook of Medical Physiology, 7th ed., p. 693. W. B. Saunders, Philadelphia. 204. Hadek, R. (1963): Electron microscope study on primary liquor folliculi secretion in the mouse ovary. J. Ultrastruct. Res., 9:445-458. 205. Hamada, Y., Bronson, R. A., Wright, K. H., and Wallach, E. E. (1977): Ovulation in the perfused rabbit ovary: The influence of prostaglandins and prostaglandin inhibitors. Biol. Reprod., 17:58— 63. 206. Hammond, J., and Marshall, F. H. A. (1925): Reproduction in the Rabbit. Oliver and Boyd, London. 207. Hammond, J. M. (1981): Peptide regulators in the ovarian follicle. Aust. J. Biol. Sci., 34:391-504. 208. Hammond, J. M., Barrando, L. S., Skaleris, D., Knight, A. B., Romanus, J. A., and Rechler, M. M. (1985): Production of insulin¬ like growth factors by ovarian granulosa cells. Endocrinology, 117:2553-2555. 209. Heape, W. (1905): Ovulation and degeneration of the ova in the rabbit. Proc. R. Soc. Fond. (Biol.), 76B:260-268. 210. von Herff, O. (1892): Ueber den fienesen Verlauf der Nerven im Eierstock des Menschen. Z. Geburtschilfe Gynakol., 24:289-308. 211. Hial, V., DeMello, M. C. F., Horakova, Z., and Beaven, M. A. (1977): Antiproliferative activity of antiinflammatory drugs in two mammalian cell culture lines. J. Pharmacol. Exp. Ther., 202:446454. 212. Hill, R. T. (1949): Adrenal cortical physiology of spleen grafted and denervated ovaries in the mouse. Exp. Med. Surg., 7:86-98. 213. Hill, R. T. (1962): Paradoxical effects of ovarian secretions. In: The Ovary, Vol. II, edited by S. Zuckerman, pp. 231-261. Academic Press, New York. 214. Hill, R. T., Allen, E., and Kramer, T. C. (1935): Cinemicrographic studies of rabbit ovulation. Anat. Rec., 63:239-245.

/

481

215. Himeno, N., Kawamura, N., Okamura, H., Mori, T., Fukumoto, M., and Midorikawa, O. (1984): Collagen synthetic activity in rabbit ovary during ovulation and its blockage by indomethecin. Acta Obstet. Gynaecol. Jpn., 36:1930-1934. 216. Himeno, N., Kawamura, N., Okamura, H., Mori, T., Fukumoto, M., and Midorikawa, O. (1984): The effect of prostaglandin F-2alpha on collagen synthesis in rabbit ovary during the ovulatory process. Acta Obstet. Gynaecol. Jpn. 36:2494-2495 (in Japanese). 217. Hinsey, J. C., and J. E. Markee. (1932): A search for neurological mechanisms in ovulation. Proc. Soc. Exp. Biol. Med., 30:136-138. 218. His, W. (1865): Beobachtungen ueber den Bau des Saugethiereierstockes. Arch. Mikr. Anat., 1:97-105. 219. Hixon, J. E., and Clegg, M. T. (1969): Influence of the pituitary on ovarian progesterone output in the eye: Effects of hypophysectomy and gonadotropic hormones. Endocrinology, 84:828-834. 220. Ichikawa, S., Morioka, H., Oda, M., Oda, K., andMurao, S. (1983): Effects of various proteinase inhibitors on ovulation of explanted hamster ovaries. J. Reprod. Fertil., 68:407^412. 221. Ichikawa, S., Ohta, M., Morioka, H., and Murao, S. (1983): Block¬ age of ovulation in the explanted hamster ovary by a collagenase inhibitor. J. Reprod. Fertil., 68:17-19. 222. Jackson, D. S. (1980): The substrate collagen. In: Collagenase in Normal and Pathological Connective Tissues, edited by D. E. Woolley and J. M. Evanson, pp. 1-10. John Wiley & Sons, New York. 223. Jacobowitz, D., and Laties, A. M. (1970): Adrenergic reinnervation of the cat ovary transplanted to the anterior chamber of the eye. Endocrinology, 86:921-924. 224. Jacobowitz, D., and Wallach, E. E. (1967): Histochemical and chem¬ ical studies of the autonomic innervation of the ovary. Endocrinology, 81:1132-1139. 225. Janson, P. O. (1975): Effects of luteinizing hormone on blood in the follicular rabbit ovary as measured by radioactive microspheres. Acta Endocrinol., 79:122-123. 226. Jensen, C. E., and Zachariae, F. (1958): Studies on the mechanism of ovulation. Isolation and analysis of acid mucopolysaccharides in bovine follicular fluid. Acta Endocrinol., 27:356-368. 227. Jocelyn, H. D., and Setchell, B. P. (1972): Translation of Regnier de Graaf. On Human Reproductive Organs from the Latin Text Tractus de Vivorum Organis Generationi Inservientibus (1668) and DeMulierum Organis Generationi Inservientibus Tractatus Novus (1672), Chapter XII, pp. 131-135. Reprod. Fertil., (Suppl.), 17:131135. 228. Johns, A., Chlumecky, J., Cottle, M., and Paton, D. M. (1975): Effect of chemical sympathectomy and adrenergic agonists on the fertility of mice. Contraception, 11:563-570. 229. Jonassen, F., Granerus, G., and Wetterquist, H. (1976): Histamine metabolism during the menstrual cycle. Acta Obstet. Gynecol. Scand., 55:297-304. 230. Jones, R. E., Duvall, D., and Guillette, L. J., Jr. (1980): Rat ovarian mast cells: Distribution and cyclic changes. Anat. Rec., 197:489493. 231. deJong, F. H., and Sharpe, R. M. (1976): Evidence for inhibin-like activity in bovine follicular fluid. Nature, 263:71-72. 232. Jordan, S. M. (1970): Adrenergic and cholinergic innervation of the reproductive tract and ovary in the guinea pig and rabbit. J. Physiol. (Lond.), 210:115-117. 233. Kang, H. H., Anderson, W. A., Chang, S. C., and Ryan, R. J. (1979): Studies on the structure of the extracellular matrices of the mammalian follicles as revealed by high-voltage electromicroscopy and cytochemistry. In: Ovarian Follicular Development and Func¬ tion, edited by A. R. Midgley and W. A. Sadler, pp. 121-135. Raven Press, New York. 234. Kannisto, P., Owman, C., Rosengren, E., and Walles, B. (1984): Intraovarian adrenergic nerves in the guinea pig: Development from fetal life to sexual maturity. Cell Tissue Res., 238:235-240. 235. Kannisto, P., Owman, C., and Walles, B. (1985): Involvement of local adrenergic receptors in the process of ovulation in gonadotro¬ phin-primed immature rats. J. Reprod. Fertil., 75:357-362. 236. Kanzaki, H. (1981): Scanning electronmicroscopic study on corro¬ sive casts for rabbit ovarian follicle microvasculature during the ovu¬ latory and luteinizing process. Acta Obstet. Gynecol. Jpn., 33:19251933. 237. Kanzaki, H., Okamura, H., Okuda, Y., Takenaka, A., Morimoto, K., and Nishimura, T. (1982): Scanning electron microscopic study

482

/ Chapter 12

of rabbit ovarian follicle microvasculature using resin injection-cor¬ rosion casts. J. Anat., 134:697-704. 238. Kapri, A., and Aratei, H. (1965): The biochemistry of the follicular fluid. Rev. Roum. Biochim., 2,3:243-248. 239. Kamovsky, M. J., and Roots, L. (1964): A “direct-coloring” thicoholine method for cholinesterases. J. Histochem. Cytochem., 12:219—

221. 240. Kasson, B. G., and Hsueh, A. J. (1985): Cholinergic inhibition of follicle-stimulating hormone-induced progesterone production by cultured rat granulosa cells. Biol. Reprod., 33:1158-1167. 241. Kasson, B. G., Meidan, R., Davoren, J. B., and Hseuh, A. J. (1985): Identification of subpopulations of rat granulosa cells: Sedimentation properties and hormonal responsiveness. Endocrinology, 117: 1027-1034. 242. von Kaulla, K. N., and Shettles, L. B. (1956): Thromboplastic ac¬ tivity of human cervical mucus and ovarian follicular and seminal fluids. Fertil. Steril., 7:166-169. 243. Kawai, Y., Satoh, K., Mitsuhashi, N., Hasumi, K., Sakakibara, K., Kinoshita, K., Wu, T., and Sakamoto, S. (1981): Prostaglandin and luteinized unruptured follicles: The follicles of gilts treated with the inhibitors of prostaglandin production. Folia Endocrinol. Jpn 57:1475-1488. 244. Kawakami, M., Kubo, K., Uemura, T., and Nagase, M. (1979): Evidence for the existence of extra-hypophyseal neural mechanisms controlling ovarian steroid secretion. J. Steroid Biochem., 11:10011005. 245. Kawakami, M., Kubo, K., Uemura, T., Nagase, M., and Hayashi, R. (1981): Involvement of ovarian innervation in steroid secretion. Endocrinology, 109:136-145. 246. Kawamura, N., Himeno, N., Okamura, H., Mori, T., Fukumoto, M., and Midorikawa, O. (1984): Effect of indomethacin on collagenolytic enzyme activities in rabbit ovary. Acta Obstet. Gynaecol Jpn., 36:2099-2105. 247. Kawamura, N., Yajema, Y., Huang, C. H., Okuda, Y., Fukumoto, M., Okamura, H., and Nishimura, T. (1981): DNP-peptidase activity in the rabbit ovary. Acta Obstet. Gynecol. Jpn. 33:1684-1688. 248. Kelly, G. L. (1931): Direct observation of rupture of the Graafian follicle in a mammal. J. Fla. Med. Assoc., 17:422-423. 249. Kiekhofer, W., Holmen, G. J., and Peckham, B. (1962): Some chemical characteristics of ovarian and parovarian cystic fluids. Ob¬ stet. Gynecol., 20:471^183. 250. Klebs, E. (1861): Die Eierstockseier der Wirbelthiere. Arch. Pathol. Anat. Physiol., 21 and 28:362-366. (quoted from Nagel). 251. Kling, O. R., Roche, P. C., Campeau, J. D., Nishimura, K., Nak¬ amura, R. M., and di Zerega, G. S. (1984): Identification of a porcine fluid fraction which suppressed follicular response to gonadotropins. Biol. Reprod., 30:564-572. 252. Knecht, M. (1986): Production of a cell-associated and secreted plasminogen activator by cultured rat granulosa cells. Endocrinology, 118:348-353. 253. Knox, E., Lowry, S., and Beck, L. (1979): Prevention of ovulation in rabbits by antihistamines. In: Ovarian Follicular Development and Function, edited by A. R. Midgley and W. A. Sadler, pp. 159-161. Raven Press, New York. 254. Kobayashi, Y., Sjoberg, N. O., Walks, B., Owman, C., Wright, K. H., Santulli, R., and Wallach, E. E. (1983): Effect of adrenergic agents on the ovulatory process in the in-vitro perfused rabbit ovary. Am. J. Obstet Gynecol., 145:857-864. 255. Kobayashi, Y., Wright, K. H., Santulli, K., Kitai, H., and Wallach, E. G. (1983): Effect of histamine and histamine blockers on the ovulatory process in vitro perfused rabbit ovary. Biol. Reprod., 28 385392. 256. Koelle, G. B. (1962): A new general concept of the neurohumoral functions of acetylcholine and acetylcholinesterase. J. Pharm. Phar¬ macol., 14:65. 257. Kohda, H., Mori, T., Ezaki, Y., Nishimura, T., and Kambegawa, A. (1980): A progesterone-dependent step in ovulation induced by human chorionic gonadotrophin in immature rats primed with pregnant mare serum gonadotrophin. J. Endocrinol., 87:105107. 258. von Kolliker, A. (1849): Beitrage zur kenntniss der glatten Muskeln. Abhandl. Wiss. Zool., 1:48-87. 259. Koos, R. D., and Clark, M. R. (1982): Production of 6-keto-prostaglandin F-l-alpha by rat granulosa cells in vitro. Endocrinology 11:1513-1518.

260. Koos, R. D., and LeMaire, W. J. (1983): Evidence for an angiogenic factor from rat follicles. In: Factors Regulating Ovarian Functions, edited by G. S. Greenwald and P. F. Terranova, pp. 191-196. Raven Press, New York. 261. Koos, R. D., Feiertag, M. A., Brodie, A. M. H., and LeMaire, W. J. (1984): Inhibition of estrogen synthesis does not inhibit lu¬ teinizing hormone-induced ovulation. Am. J. Obstet. Gynecol., 148:939-945. 262. Kranzfelder, D., Korr, H., Mestwerdt, W., and Maurer-Schultze, B. (1984): Follicle growth in the ovary of the rabbit after ovulationinducing application of human chorionic gonadotropin. Cell Tissue Res., 238:611-620. 263. Kraus, S. D. (1947): Observations on the mechanism of ovulation in the frog, hen and rabbit. West J. Surg. Obstet. Gynecol., 55:424437. 264. Krishna, A., and Terranova, P. F. (1985): Alterations in mast cell degranulation and ovarian histamine in the proestrous hamster. Biol. Reprod., 32:1211-1217. 265. Kristek, F., Tesarik, J., and Unzeiting, V. (1984): Ultrastructure of the human theca. Folia Morphol., 32:5-8. 266. Kuehl, F. A., Jr., and Egan, R. W. (1980): Prostaglandins, arachidonic acid and inflammation. Science, 210:978-984. “267. Kulkami, P. S., Wakade, A. R., and Kirpekar, S. M. (1976): Sym¬ pathetic innervation of guinea pig uterus and ovary. Am. J. Physiol., 230:1400-1405. 268. Kuntz, A. (1945): The Autonomic Nervous System, 3rd ed. Lea and Febiger, Philadelphia. 269. Lamprecht, S. A., Zor, U., Tsafriri, A., and Lindner, H. R. (1971): Action of prostaglandin E2 and luteinizing hormone on cyclic aden¬ osine 3'-5'-monophosphate production and protein kinase activity in fetal, early postnatal and adult rat ovaries. Isr. J. Med. Sci., 7:704705. 270. Lamprecht, S. A., Zor, U., Tsafriri, A., and Lindner, H. R. (1973): Action of prostaglandin E2 and of luteinizing hormone on ovarian adenylate cyclase, protein kinase, and ornithine decarboxylase ac¬ tivity during postnatal development and maturity in the rat. J. En¬ docrinol., 57:217-233. 271. Larsen, W. J. (1977): Structural diversity of gap junctions: A review. Tissue Cell, 9:373-394. 272. Larsen, W. J., and Tung, H. N. (1978): Origin and fate of cyto¬ plasmic gap junctional vesicles in rabbit granulosa cells. Tissue Cell, 10:585-598. 273. Larsen, W. J., Tung, H. N., Murray, S. A., and Swenson, C. A. (1979): Evidence for the participation of actin microfilaments and bristle coats in the internalization of gap junction membrane. J. Cell Biol., 83:576-583. 274. Larsen, W. J., Tung, H. N., and Polking, C. (1981): Response of granulosa cell gap junctions to human chorionic gonadotropin (hCG) at ovulation. Biol. Reprod., 25:1119-1134. 275. Lau, I. F., Saksena, S. K., and Chang, M. C. (1974): Prostaglandins F and ovulation in mice. J. Reprod. Fertil., 40:467-469. 276. Lawrence, I. E., and Burden, H. W. (1976): The autonomic inner¬ vation of the interstitial gland of the rat ovary during pregnancy. Am. J. Anat., 147:81-94. 277. Lawrence, T. S., Beers, W. H., and N. B. Gilula. (1978): Trans¬ mission of hormonal stimulation by cell-to-cell communication Na¬ ture, 272:501-506. 278. Lawrence, T. S., Ginzberg, R. D., Gilula, N. B., and W. H. Beers. (1979): Hormonally induced cell shape changes in cultured rat ovarian granulosa cells. J. Cell Biol., 80:21-36. 279. Ledwitz-Rigby, F., and Ribgy, B. W. (1983): The actions of follic¬ ular fluid factors on steroidogenesis by cultured ovarian granulosa cells. J. Steroid Biochem., 19:127-131. 280. Lee, W., and Novy, M. J. (1978): Effects of luteinizing hormone and indomethacin on blood flow and steroidogenesis in the rabbit ovary. Biol. Reprod., 18:799-807. 281. LeMaire, W. J., and Marsh, J. M. (1975): Interrelationships between prostaglandins, cyclic AMP and steroids in ovulation J Reprod Fertil. (Suppl.), 22:53-74. 282. LeMaire, W. J., Davies, P. J., and Marsh, J. M. (1976): The role of prostaglandins in the development of refractoriness to LH stim¬ ulation by Graafian follicles. Prostaglandins, 12:271-279. 283. LeMaire, W. J., Leidner, R., and Marsh, J. M. (1975): Pre- and postovulatory changes in the concentration of prostaglandins in rat Graafian follicles. Prostaglandins, 9:221-229.

Mammalian Ovulation

284. LeMaire, W. J., Yang, N. S. T., Behrman, H. R., and Marsh, J. M. (1973): Preovulatory changes in the concentration of prosta¬ glandins in rabbit Graafian follicles. Prostaglandins, 3:367. 285. Lenz, R. W., Ax, R. L., and First, N. L. (1982): Proteoglycan production by bovine granulosa cells in vivo is regulated by cal¬ modulin and calcium. Endocrinology, 110:1052- 1054. 286. LePere, R. H., Benoit, P. E., Hardy, R. C., and Goldzieher, J. W. (1966): The origin and function of the ovarian nerve supply in the baboon. Fertil. Steril., 17:68-75. 287. Levin, E. G., and Loskutoff, D. J. (1982): Cultured bovine endo¬ thelial cells produce both urokinase and tissue-type plasminogen ac¬ tivators. J. Cell Biol. 94:631-636. 288. Lim, A. T., Lolait, S., Barlow, J. W., Wai Sum, O., Zois, I., Toh, B. H., and Funder, J. W. (1983): Immunoreactive beta-endorphin in sheep. Nature, 303:709-711. 289. Lindner, H. R., Tsafriri, A., Lieberman, M. E., Zor, U., Koch, Y., Bauminger, S., and Bame, A. (1974): Gonadotropin action on cul¬ tured Graafian follicles: Induction of maturation division of the mam¬ malian oocyte and differentiation of the luteal cell. Rec. Prog. Hor¬ mone Res., 30:79-138. 290. Lindner, H. R., Zor, U., Kohen, F., Bauminger, S., Amsterdam, A. , Lahar, M., and Salomon, Y. (1980): Significance of prosta¬ glandins in the regulation of cyclic events in the ovary and uterus. Adv. Prostaglandin Thromboxane Leukotriene Res., 8:1371-1390. 291. Lipner, H. (1971): Ovulation from histamine depleted ovaries. Proc. Soc. Exp. Biol. Med., 136:111-114. 292. Lipner, H. (1973): Mechanism of mammalian ovulation. In: Hand¬ book of Physiology—Endocrinology II, Part 1, Chapter 18, pp. 409437. 293. Lipner, H., and Cross, N. L. (1968): Morphology of the membrana granulosa of the ovarian follicle. Endocrinology, 82:638-641. 294. Lipner, H., and Greep, R. O. (1971): Inhibition of steroidogenesis at various sites in the biosynthetic pathway and the relationship to induced ovulation. Endocrinology, 88:602-607. 295. Lipner, H., and Maxwell, B. A. (1960): Hypothesis concerning the role of follicular contractions in ovulation. Science, 131:1737-1738. 296. Lipner, H., and Smith, M. S. (1971): A method for determining the distribution and source of protein in preovulatory rat ovaries. J. Endocrinol., 50:1-14. 297. Lipner, H., and Wendelkin, L. (1971): Inhibition of ovulation by inhibition of steroidogenesis in immature rats. Proc. Soc. Exp. Biol. Med., 136:1141-1145. 298. Liu, W. K., Burleigh, B. D., and Ward, D. N. (1981): Steroid and plasminogen activator production by cultured rat granulosa cells in response to hormone treatment. Mol. Cell. Endocrinol. 21:63-73. 299. Loewenstein, W. R. (1981): Junctional intercellular communication: The cell-to-cell membrane channel. Physiol. Rev., 61:829-913. 300. Lupulescu, A. P. (1977): Cytologic and metabolic effects of pros¬ taglandins on rat skin. J. Invest. Dermatol., 68:138-145. 301. Lutwak-Mann, C. (1954): Note on the chemical composition of bo¬ vine follicular fluid. J. Agr. Sci., 44:477-480. 302. Macdonald, E. J., and Airaksinen, M. M. (1974): The effect of 6hydroxydopamine on the oestrus cycle and fertility of rats. J. Pharm. Pharmacol., 26:518-521. 303. Mai, H., Jr., Barbosa, I., and Coutinho, E. M. (1975): Effects of aminophylline, imidazole and indomethacin on spontaneous and prostaglandin induced ovarian contractions in vitro. Int. J. Fertil., 20:82-86. 304. Mai, H., Jr., Barbosa, I., and Coutinho, E. M. (1978): Inhibition of ovulation in marmoset monkeys by indomethacin. Fertil. Steril., 29:565-570. 305. Makinada, S. (1980): Hemodynamics and histological studies on ovarian blood flow during ovulation. Hokkaido J. Med. Sci., 55:521 526. 306. Makris, A., Ryan, K. H., Hasumizu, T., Hill, C. L., and Zetter, B. R. (1984): The nonluteal porcine ovary as a source of angiogenic activity. Endocrinology, 115:1672-1677. 307. Manarang-Pangan, S.,'and Menge, A. C. (1971): Immunologic stud¬ ies on human follicular fluid. Fertil. Steril., 22:367-372. 308. Mandl, L. (1895): Ueber Anordnung und Endigungsweise der Nerven im Ovarium. Arch. Gynecol., 48:376. 309. Marion, G. B., Gier, H. T., and Choudary, J. B. (1968): Micro¬ morphology of the bovine ovarian follicular system. J. Anim. Sci., 27:451^165.

/

483

310. Markee, J. E., and Hinsey, J. C. (1936): Observations on ovulation in the rabbit. Anat. Rec., 64:309-319. 311. Marotti, K. R., Belin, D., and Strickland, S. (1982): The production of distinct forms of plasminogen activator by mouse embryonic cells. Dev. Biol. 90:154-159. 312. Martin, G. G., and Miller-Walker, W. C. (1983): Visualization of the 3-dimensional distribution of collagen fibrils over pre-ovulatory follicles in the hamster Mesocricetus auratus. J. Exp. Zool., 225:311320. 313. Martin, G. G., and Talbot, P. (1981): The role of follicular smooth muscle cells in hamster ovulation. J. Exp. Zool., 216:469—482. 314. Martinat, N., and Combamous, Y. (1983): The release of plasmin¬ ogen activator by rat granulosa cells is highly specific for FSH ac¬ tivity. Endocrinology, 119:433^135. 315. McCracken, J. A., Baird, D. T., and Goding, J. R. (1971): Factors affecting the secretion of steroids from autotransplanted ovary in the sheep. Recent Prog. Hormone Res., 27:537-582. 316. McGaughey, R. W. (1975): A comparison of the fluids from small and large ovarian follicles of the pig. Biol. Reprod., 13:147-153. 317. McNatty, K. P. (1978): Follicular fluid. In: The Vertebrate Ovary, edited by R. E. Jones, pp. 215-259. Plenum Press, New York. 318. McReynolds, H. D., Siraki, C. M., Bramson, P. H., and Pollock, R. J., Jr. (1973): Smooth muscle-like cells in ovaries of the hamster and gerbil. Z. Zellforsch., 140:1-8. 319. Merk, F. B., Albright, J. T., and Botticelli, C. R. (1973): The fine structure of granulosa cell nexuses in rat ovarian follicles. Anat. Rec., 175:107-125. 320. Miller, E. J. (1985): Recent information on the chemistry of the collagens. In: The Chemistry and Biology of Mineralized Tissues, edited by W. T. Butler, pp. 80-93. EBSCO Media, Birmingham, AL. 321. Miller, F. N., and Sims, D. E. (1986): Contractile elements in the regulation of macromolecular permeability. Fed. Proc., 45: 84-88. 322. Miller, K. F., Wesson, J. A., and Ginther, O. J. (1979): Changes in concentration of circulating gonadotrophins following administra¬ tion of equine follicular fluid to ovariectomized mares. Biol. Reprod., 21:867-872. 323. Mitchell, G. A. G. (1938): Innervation of ovary, uterine tube, testis and epididymis. J. Anat.. 72:508-517. 324. Mitsuhashi, N. (1981): Studies on the mechanism and the significance of prostaglandin biosynthesis by the ovary, ovulation block by the indomethacin and incubation of the follicle. Acta Obstet. Gynaecol. Jpn., 33:479-488. 325. Montz, F. J., Ujita, E. L., Campeau, J. D., and diZerega, G. S. (1984): Inhibition of luteinizing hormone, human chorionic gonad¬ otropin binding to porcine granulosa cells by a follicular fluid protein. Am. J. Obstet. Gynecol., 148:436-441. 326. Moor, R. M., Hay, M. F., and Seamark, R. F. (1975): The sheep ovary: Regulation of steroidogenic, haemodynamic and structural changes in the largest follicle and adjacent tissue before ovulation. J. Reprod. Fertil., 45:595-604. 321. Morales, T. I., Woessner, J. F., Howell, D. S., Marsh, J. M., and LeMaire, W. J. (1978): A microassay for the direct demonstration of collagenolytic activity in Graafian follicles of the rat. Biochim. Biophys. Acta, 524:428-434. 328. Morales, T. I., Woessner, J. F., Jr., Marsh, J. M., and LeMaire, W. J. (1983): Collagen, collagenase and collagenolytic activity in rat Graafian follicles during follicular growth and ovulation. Biochim. Biophys. Acta, 756:119-122. 329. Mori, T., Suzuki, A., Nishimura, T., and Kambegawa, A. (1977): Inhibition of ovulation in immature rats by antiprogesterone anti¬ serum. J. Endocrinol., 73:185-186. 330. Mori, T., Suzuki, A., Nishimura, T., and Kambegawa, A. (1977): Evidence for androgen participation in induced ovulation in immature rats. Endocrinology, 101:623-626. 331. Morikawa, H., Okamura, H., Takenaka, A., Morimoto, K., and Nishimura, T. (1981): Histamine concentration and its effect on ovarian contractility in humans. Int. J. Fertil., 26:283-286. 332. Morimoto, K., Okamura, H., Kanzaki, H., Okuda, Y., Takenaka, A., and Nishimura, T. (1981): Adrenergic nerve supply to bovine ovarian follicles. Int. J. Fertil., 26:14-19. 333. Morimoto, K., Okamura, H., and Tanaka, C. (1982): Development and periovulatory changes of ovarian norepinephrine in the rat. Am. J. Obstet. Gynecol., 143:389-392.

484

/

Chapter 12

334. Mossman, H. W., and Duke, K. L. (1973): Comparative Morphology of the Mammalian Ovary. University of Wisconsin Press, Madison, Wis. 335. Motta, P.; Chemey, D. D., and Didio, L. J. A. (1971): Scanning and transmission electromicroscopy of the ovarian surface in mam¬ mals with special reference to ovulation. J. Submicr. Cytol. 3:85-

100. 336. Motta, P. M., and Familiari, G. (1981): Occurrence of a contractile tissue in the theca externa of atretic follicles in the mouse ovary. ActaAnat. (Basal), 109:103-114. 337. Motta, P. M., and Makabe, S. (1982): Development of the ovarian surface and associated germ cells in the human fetus: a correlated study by scanning electron microscopy and transmission electron microscopy. Cell Tissue Res., 226:493-510. 338. Mueller, P. L., Schreiber, J. R., Lucky, A. W., Schulman, J. D., Rodbard, D., and Ross, G. T. (1978): Follicle-stimulating hormone stimulates ovarian synthesis of proteoglycans in estrogen-stimulated hypopohysectomized immature female rats. Endocrinology, 102:824831. 339. Murdoch, W. J., and Dunn, T. G. (1982): Alterations in follicular steroid hormones during the preovulatory period in the ewe. Biol. Reprod., 27:300-307. 340. Murdoch, W. J., and Myers, D. A. (1983): Effect of treatment of estrous ewes with indomethacin on the distribution of ovarian blood to the periovulatory follicle. Biol. Reprod., 29:12291232. 341. Murdoch, W. J., Dailey, R. A., and Inskeep, E. K. (1981): Pre¬ ovulatory changes in prostaglandins E2 and F2 in ovine follicles. J. Animal Sci., 53:192—205. 342. Murdoch, W. J., Nix, K. J., and Dunn, T. G. (1983): Dynamics of ovarian blood supply to periovulatory follicles of the ewe. Biol. Reprod., 28:1001-1006. 343. Murphy, G., and Sellers, A. (1980): The extracellular regulation of collagenase activity. In: Collagenase in Normal and Pathological Connective Tissues, edited by D. E. Woolley and J. M. Evanson, pp. 65-81. John Wiley & Sons, New York. 344. Nagel, W. (1896): Ham und Geschlechtsorgane. In: Handbuck der Anatomie des Menschen. Bd. VII, edited by K. von Bardeleben, pp. 42-64. Verlag G. Fischer, Jena. 345. Neilson, D., Jones, G. S., Woodruff, J. D., and Goldberg, B. (1970): The innervation of the ovary. Obstet. Gynecol. Surv., 25:889-904. 346. Nickander, R., McMahon, F. G., and Ridolfo, A. S. (1979): Non¬ steroidal antiinflammatory agents. Annu. Rev. Pharmacol. Toxicol., 19:469-490. 347. Niswender, G. D., Reimers, T. J., Diekman, M. A., and Nett, T. M. (1976): Blood flow: A mediator of ovarian function. Biol. Reprod., 14:64-81. 348. Norjavaara, E., Selstam, G., and Ahren, K. (1982): Catecholamine stimulation of cyclic AMP and progesterone production in the rat corpora lutea of different ages. Acta Endocrinol. (Copenh.), 100:613— 622. 349. Novy, M. J., and Cook, M. J. (1973): Redistribution of blood flow by prostaglandin F2 in the rabbit ovary. Am. J. Obstet. Gynecol., 117:381-385. 350. Nowacki, R. (1977): Das zeitgenossische Bild desreifen ovarialfollikels der Saugetiere. Zbl. Vet. Med. C Anat. Hist. Embryol., 6:217— 239. 351. Ny, T., Bjersing, L., Hsueh, A. J. W., and Loskutoff, D. J. (1985): Cultured granulosa cells produce two plasminogen activators, each regulated by gonadotropins. Endocrinology, 116:1666-1668. 352. Odin, L. (1959): Studies on the chemistry of ovarian cyst contents. Acta Soc. Med. Upsala, 64:25-91. 353. O’Grady, J. P., Caldwell, B. V., Auletta, F. J., and Speroff, L. (1972): The effects of an inhibitor of prostaglandin synthesis (in¬ domethacin) on ovulation, pregnancy and pseudopregnancy in the rabbit. Prostaglandins, 1:97-106. 354. Ohlsson, K. (1980): Polymorphonuclear leucocyte collagenase. In: Collagenase in Normal and Pathological Connective Tissues, edited by D. E. Woolley and J. M. Evanson, pp. 209-222. John Wiley & Sons, New York. 355. Ohno, Y., and Mori, T. (1985): Correlation between progesterone and plasminogen activator in rat ovaries during the ovulatory process. Acta Obstet. Gynaecol. Jpn., 37:247-256.

356. Ojeda, S. R., White, S. S., Aguado, L. I., and Andersen, J. M. (1983): Abdominal vagotomy delays the onset of puberty and inhibits ovarian function in the female rat. Neuroendocrinology, 36:261-267. 357. Okamura, H. , Okazaki, T., andNakajima, A. (1974): Human ovarian contractility m vitro. Acta Obstet. Gynecol. Jpn., 21:89-96. 358. Okamura, H., Okazaki, T., and Nakajima, A. (1974): Effects of neurotransmitters and prostaglandins on human ovarian contractility. Obstet. Gynecol., 44:720-726. 359. Okamura, H., Takenaka, A., Yajima, Y., and Nishimura, T. (1980): Ovulatory changes in the wall at the apex of the human Graafian follicle. J. Reprod. Fertil., 58:153-155. 360. Okamura, H., Virutamasen, P., Wright, K. H., and Wallach, E. E. (1972): Ovarian smooth muscle in the human being, rabbit, and cat. Am. J. Obstet. Gynecol., 112:183-191. 361. Okamura, H., Okuda, Y., Kanzaki, H., Takenaka, A., Morimoto, K., and Nishimura, T. (1981): Ultrastructural observation of the ovulatory changes in the capillary of the human follicular apex. Acta Obstet. Gynecol. Jpn., 33:215—221. 362. Okazaki, T., Okamura, H., and Nishimura, T. (1973): Collagenolytic activity in rat ovary during ovulation. Acta Obstet. Gynecol. Jpn., 20:112-117. 363. Okuda, Y. (1983): An ultrastructural study on capillary permeability of the rabbit ovarian follicles during ovulation. J. Clin. Electron Microsc., 16:117-134. 364. Okuda, Y., Okamura, H., Kanzaki, H., and Takenaka, A. (1983): Capillary permeability of rabbit ovarian follicles prior to ovulation. J. Anat., 137:263-269. 365. Okuda, Y., Okamura, H., Kanzaki, H., Takenaka, A., Morimoto, K. , and Nishimura, T. (1980): An ultrastructural study of capillaries of rabbit follicles during ovulatory process. Acta Obstet. Gynecol. Jpn., 32:739-748. 366. Okuda, Y., Okamura, H., Kanzaki, H., Takenaka, A., Morimoto, K., and Nishimura, T. (1980): An ultrastructural study of capillary permeability of rabbit follicles during ovulation using carbon traces. Acta Obstet. Gynecol. Jpn., 32:859-867. 367. Okuda, Y., Okamura, H., Kanzaki, H., Takenaka, H., Morimoto, K., and Nishimura, T. (1982): An ultrastructural study of capillary permeability of rabbit ovarian follicles using horse radish peroxidase as a tracer. Acta Obstet. Gynecol. Jpn., 34:181-186. 368. Okuda, Y., Okamura, H., Kanzaki, H., Fujii, S., Takenaka, A., and Wallach, E. E. (1983): An ultrastructural study of ovarian peri¬ follicular capillaries in the indomethacin-treated rabbit. Fertil Steril., 39:85-92. 369. O’Shea, J. D. (1970): An ultrastructural study of smooth muscle¬ like cells in the theca externa of ovarian follicles in the rat. Anat Rec„ 167:127-140. 370. O’Shea, J. D. (1971): Smooth muscle-like cells in the theca externa of ovarian follicles in the sheep. J. Reprod. Fertil., 24:283-285. 371. O Shea, J. D. (1981): Structure-function relationships in the wall of the ovarian follicle. Aust. J. Biol. Sci., 34:379-394. 372. O’Shea, J. D., Cran, D. G., Hay, M. F., and Moor, R. M. (1978): Ultrastructure of the theca interna of ovarian follicles in sheep Cell Tissue Res., 187:457-472. 373. O Shea, J. D., and Phillips, R. E. (1974): Contractility in vitro of ovarian follicles from sheep, and the effects of drugs. Biol. Reprod 10:370-379. 374. Ossowski, L., Biegel, O., and Reich, E. (1979): Mammary plas¬ minogen activator: Correlation with involution, hormonal modulation and comparison between normal and neoplastic tissues. Cell 16 929940. 375. Osterholzer, H. O., Johnson, J. H., and Nicosia, S. V. (1985): An autoradiographic study of rabbit ovarian surface epithelium before and after ovulation. Biol. Reprod., 53:729-738. 376. Osterman, J., and Hammond, J. M. (1978): Prostaglandin stimulation of ovarian ornithine decarboxylase EC-4.1.1.17 in vitro. Biochem. Biophys Res. Commun., 83:794-799. 377. Osvaldo-Decima, L. (1970): Smooth muscle in the ovary of the rat and monkey. J. Ultrastruct. Res., 29:218-237. 378. Owman, C., Rosengren, E., and Sjoberg, N. O. (1967): Adrenergic innervation of the human female reproductive organs: A histochemical and chemical investigation. Obstet. Gynecol., 30:763-773. 379. Owman, C., Sjoberg, N. O., Svensson, K. G., and Walles, B. (1975): Autonomic nerves mediating contractility in the human graa¬ fian follicle. J. Reprod. Fertil., 45:553-556.

Mammalian Ovulation

380. Owman, C., Sjoberg, N. O., Wallach, E. E., Walles, B., and Wright, K. H. (1979): Neuromuscular mechanisms of ovulation. In: Human Ovulation: Mechanisms, Prediction, Detection and Regulation, edited by E. S. E. Hafez, pp. 57-100. Elsevier/North Holland, Am¬ sterdam. 381. Palla, V. (1952): Azione del solfata di protamina sulla sostanza eparinosimile del liquor folliculi. Minerva Ginecol., 4:164-174. 382. Palotie, A., Peltenen, L., Foidart, J. M., and Rajaniemi, H. (1984): Immunohistochemical localization of basement membrane compo¬ nents and interstitial collagen types in preovulatory rat ovarian fol¬ licles. Coll. Relat. Res., 4:279-287. 383. Palti, Z., and Freund, M. (1972): Spontaneous contractions of the human ovary in vitro. J. Reprod. Fertil., 28:113-115. 384. Panganamala, R. V., Miller, J. S., Gwebu, E. T., Sharma, H. M., and Cornwell, D. G. (1977): Differential inhibitory effects of vitamin E and other antioxidants on prostaglandin synthetase, platelet ag¬ gregation and lipoxidase. Prostaglandins, 14:261-271. 385. Parr, E. L. (1974): Histological examination of the rat ovarian follicle wall prior to ovulation. Biol. Reprod., 11:483-503. 386. Parr, E. L. (1975): Rupture of ovarian follicles at ovulation. J. Reprod. Fertil. (Suppl.), 22:1-22. 387. Pascu, T., Tudorascu, R., Stancioiu, N., and Lunca, H. (1971): Concentration des proteines totales et des fractions proteiques dans le liquide folliculaire normal, pendant les differentes phases due cycle oestral, et dans le liquide des kystes folicularies ovariens, ainse que dans le sang des mimes vaches. Reel. Med. Vet., 147:979-991. 388. Patwardhan, V. V., and Lanthier, A. (1981): Prostaglandin E and prostaglandin F in human ovarian follicles: Endogenous contents and in-vitro formation by theca and granulosa cells. Acta Endocrinol., 97:543-550. 389. Pendergrass, P. B. (1980): Ultrastructural comparison of smooth muscle from ovarian follicle and oviduct of the golden hamster. Cell Tissue Res., 209:43—48. 390. Pendergrass, P. B., and Reber, M. (1980): Scanning electron micros¬ copy of the Graafian follicle during ovulation in the golden hamster. J. Reprod. Fertil., 59:21-24. 391. Pendergrass, P. B., and Talbot, P. (1979): The distribution of con¬ tractile cells in the apex of the preovulatory hamster follicle. Biol. Reprod., 20:205-213. 392. Penkala, J. E., and Talbot, P. (1984): Inhibitors of proteinase activity block in vivo ovulation in the golden hamster. J. Cell Biol., 99:391 (abstract). 393. Peracchia, C. (1980): Structural correlates of gap junction permea¬ tion. Int. Rev. Cytol., 66:81-146. 394. Peracchia, C. (1984): Communicating junctions and calmodulin: In¬ hibition of electrical uncoupling in Xenopus embryo by calmidazolium. Membr. Biol., 81:49-58. 395. Perkiev, T., and Ahren, K. (1971): Effects of prostaglandins, LH and polyphloretin phosphate on the lactic acid production of the prepubertal rat ovary. Life Sci., 10:1387-1393. 396. Perloff, W. H., Schultz, J., Farris, E., and Balin, H. (1955): Some aspects of the chemical nature of human ovarian follicular fluid. Fertil. Steril., 6:11-17. 397. Peters, H. (1969): The development of the ovary from birth to ma¬ turity. Acta Endocrinol. (Copenh.)., 62:98-116. 398. Pettigrew, D. V., Ho, G. H., Sodek, J., Brunette, D. M., and Wang, H. M. (1978): Effect of oxygen tension and indomethacin on pro¬ duction of collagenase and neutral proteinase enzymes and their latent forms by porcine gingival explants in culture. Arch. Oral Biol., 23:767-777. 399. Pfluger, E. (1857): Ueber die Bewegungen der Ovarien. Arch. Anat. Physiol. Wissensch. Med., 1:30-32. 400. Pfluger, E. F. W. (1863): Ueber die Eierstocke der Saugethiere und des Menschen. W. Englemann, Leipzig. 401. Piacsek, B. E., and Huth, J. F. (1971): Changes in ovarian venous blood flow following cannulation: Effect of luteinizing hormone (LH) and antihistamine. Proc. Soc. Exp. Biol. Med., 138:1022-1024. 402. Plunkett, E. R., Moon, Y. S., Zamecnik, J., and Armstrong, D. (1975): Preliminary evidence of a role for prostaglandin F in human follicular function. Am. J. Obstet. Gynecol., 123:391-397. 403. Pool, W. R., and Lipner, H. (1965): Inhibition of ovulation in the rabbit by actinomycin D. Nature, 203:1385-1387. 404. Pool, W. R., and Lipner, H. (1966): Inhibition of ovulation by antibiotics. Endocrinology, 79:858-864.

/

485

405. Poyser, N. L., and Scott, F. M. (1980): Prostaglandin and thromboxyane production by the rat uterus and ovary in vitro during the estrous cycle. J. Reprod. Fertil., 60:33-40. 406. Priedkalns, J., Weber, A. F., and Zemjanis, R. (1968): Qualitative and quantitative morphological studies of the cells of the membranosa, theca interna and corpus luteum of the bovine ovary. Z. Zellforsch., 85:501-520. 407. Ramwell, P. W., Shaw, J. E., and Jessup, S. J. (1969): Follicular fluid kinin and its action on fallopian tube. Endocrinology, 84:931936. 408. Rand, M. J., and Trinker, F. R. (1966): Pharmacological agents affecting the release and activity of catecholamines. Br. J. Anaesth., 38:666-689. 409. Ratner, A., Weiss, G. R., and Sanborn, C. R. (1980): Stimulation by B2-adrenergic receptors of the production of cyclic AMP and progesterone in rat ovarian tissue. J. Endocrinol., 87:123-129. 410. Rawson, J. M., and Espey, L. L. (1977): Concentration of electron dense granules in the rabbit ovarian surface epithelium during ovu¬ lation. Biol. Reprod., 17:561-566. 411. Reich, R., Miskin, R., and Tsafriri, A. (1985): Follicular plasmin¬ ogen activator: Involvement in ovulation. Endocrinology, 116:516— 521. 412. Reich, R., Tsafriri, A., and Mechanic, G. L. (1985): The involve¬ ment of collagenolysis in ovulation in the rat. Endocrinology, 116:522527. 413. Reich, R., Kohen, F., Naor, Z., and Tsafriri, A. (1983): Possible involvement of lipoxygenase products of arachidonic acid pathway in ovulation. Prostaglandins, 26:1011-1020. 414. Reich, R., Kohen, F., Slager, R., and Tsafriri, A. (1985): Ovarian lipoxygenase activity and its regulation by gonadotropin in the rat. Prostaglandins, 30:581-590. 415. Retzius, G. (1893): Ueber die Nerven der Ovarium und Hoden. Biol. Untersuch., 5:31-34. 416. Reynolds, S. R. M. (1973): Blood and lymph vascular systems of the ovary. In: Handbook of Physiology, Vol. II, Section 7, pp. 261— 316, American Physiological Society, Washington, D.C. 417. Riese, H. (1891): Die feinsten Nervenfasem und ihre Endigungen in Ovarium der Saugethiere und des Menschen. Anat. Anz., 6:400-420. 418. Rigby, B. W., Ling, S. Y., and Ledwitz-Rigby, F. L. (1983): In search of the elusive follicular fluid factors. In: Factors Regulating Ovarian Function, edited by G. S. Greenwald and P. F. Terranova, pp. 179-183. Raven Press, New York. 419. Roca, R., Garofalo, E., Piriz, H., Martino, 1., Rieppi, G., and Sala, M. (1976): Effects of oxytocin on in vitro ovarian contractility during the estrous cycle of the rat. Biol. Reprod., 15:464-466. 420. Rocerto, T., Jacobowitz, D. and Wallach, E. (1969): Observations of spontaneous contractions of the cat ovary in vitro. Endocrinology, 84:1336-1341. 421. Rondell, P. (1964): Follicular pressure and distensibility. Am. J. Physiol., 207:590-594. 422. Rondell, P. (1970): Biophysical aspects of ovulation. Biol. Reprod. Suppl., 2:64-89. 423. Rosenfeld, C. R., Morriss, F. H., Jr., Battaglia, F. C., Mokowski, E. L., and Mescha, G. (1976): Effect of estradiol-17p on blood flow to reproductive and nonreproductive tissues in pregnant ewes. Am. J. Obstet. Gynecol., 124:618-629. 424. Rosengren, E., and Sjoberg, M. O. (1967): The adrenergic nerve supply to the female reproductive tract of the cat. Am. J. Anat., 121:271-284. 425. Rouget, C. (1858): Recherches sur les organes erectiles de la femme et sur l’appareil musculataire tubo ovarien dans luvs rapports avec l’ovulation et la menstruation. J. Physiol. (Paris), 1:320-343. 426. Sato, T., Taya, K., Jyiyo, T., and Igarashi, M. (1974): Ovula¬ tion block by indomethacin, an inhibitor of prostaglandin syn¬ thesis: A study of its site of action in rats. J. Reprod. Fertil., 39: 33-40. 427. Schmidt, G., Owman, C., Sjoberg, N. O., and Walles, B. (1985): Influence of adrenoreceptor agonists and antagonists on ovulation in the rabbit ovary perfused in vitro. J. Auton. Pharmacol., 5:241-250. 428. Schochet, S. S. (1916): A suggestion as to the process of ovulation and ovarian cyst formation. Anat. Rec. 10:447^157. 429. Schroder, R. (1930): Weibliche Genitalorgane. In: Handbuch mikroanatomische des Menschen. Vol. II, edited by W. von Mollendorff, pp. 1-329. Springer-Verlag, Berlin.

486

/ Chapter 12

430. Schroeder, P. C., and Talbot, P. (1982): Intrafollicular pressure decreases in hamster preovulatory follicles during smooth muscle cell contraction in vitro. J. Exp. Zool., 224:417-426. 431. Schwartz, N. B., and Channing, C. P. (1977): Evidence for ovarian inhibin: Suppression of the secondary rise in serum follicle stimu¬ lating hormone levels in proestrous rats by injection of porcine fol¬ licular fluid. Proc. Natl. Acad. Sci. USA, 74:5721-5724. 432. Schwartz, N. B., and McCormack, C. E. (1972): Reproduction: Gonadal function and its regulation. Annu. Rev. Physiol., 34:425472. 433. Semenova, I. I. (1969): Adrenergic innervation and distribution of cholinesterases in the human ovary. Bull. Eksp. Biol. Med., 68:103106. 434. Shalgi, R., Kaplan, R., and Kraicer, P. F. (1977): Proteins of fol¬ licular, bursal and ampullar fluids of rats. Biol. Reprod., 17:333— 338. 435. Shalgi, R., Kraicer, P. F., and Soferman, N. (1972): Gases and electrolytes of human follicular fluid. J. Reprod. Fertil., 28:335340. 436. Shalgi, R., Kraicer, P. G., and Soferman, N. (1972): Human fol¬ licular fluid. J. Reprod. Fertil., 31:515-516. 437. Shalgi, R., Kraicer, P., Rimon, A., Pinto, M., and Soferman, N. (1973): Proteins of human follicular fluid: The blood-follicle barrier. Fertil. Steril., 24:429-434. 438. Shepro, D., Schleef, R., and Hecktman, H. B. (1980): Plasminogen activator activity by cultured bovine aortic endothelial cells. Life Sci. 26:415^422. 439. Shimada, H., Okamura, H., Nada, Y., Suzuki, A., Tojo, S., and Takada, A. (1983): Plasminogen activator in rat ovary during the ovulatory process: Independence of prostaglandin mediation. J. En¬ docrinol., 97:201-205. 440. Shivers, C. A., Metz, C. B., and Lutwak-Mann, C. (1964): Some properties of pig follicular fluid. J. Reprod. Fertil., 8:115-120. 441. Simpson, E. R., Rochelle, D. B., Carr, B. R., Macdonald, P. C., Cecil, H., and Green, J. (1980): Plasma lipoproteins in follicular fluid of human ovaries. 7. Clin. Endocrinol. Metab., 51:1469-1471. 442. Sjoberg, N. O. (1968): Increase in transmitter content of adrenergic nerves in the reproductive tract of female rabbits after oestrogen treatment. Acta Endocrinol., 57:405-413. 443. Skinner, M. K., McKeracher, H. L., and Dorrington, J. H. (1985): Fibronectin as a marker of granulosa cell cytodifferentiation. En¬ docrinology, 117:886-892. 444. Smith, J. T. (1934): Some observations on the rupture of the Graafian follicles in rabbits. Am. J. Obstet. Gynecol., 27:728-730. 445. Smith, J. T. (1937): Rupture of Graafian follicles. Am. J. Obstet. Gynecol., 33:820-827. 446. Sporrong, B., Kannisto, P., Owman, C., Sjoberg, N. O., and Walles, B. (1985): Histochemistry and ultrastructure of adrenergic and acetylcholinesterase-containing nerves supplying follicles and endocrine cells in the guinea pig ovary. Cell Tissue Res., 240: 505-511. 447. Stangroom, J. E., and Weevers, R. deG. (1962): Anticoagulant ac¬ tivity of equine follicular fluid. J. Reprod. Fertil., 3:269-282. 448. Starke, K. (1979): Presynaptic regulation of release in the central nervous system. In: The Release of Catecholamines from Adrenergic Neurons, edited by D. M. Paton, pp. 143-184. Pergamon Press, New York. 449. Stefenson, A., Owman, C. M., Sjoberg, N. O., Sporrong, B., and Walles, B. (1981): Comparative study of the autonomic innervation of the mammalian ovary with particular regard to the follicular sys¬ tem. Cell Tissue Res., 215:47-62. 450. Sterin-Borda, L., Borda, E., Gimeno, M. F., and Gimeno, A. L. (1976): Spontaneous and prostaglandin or oxytocin induced motility of rat ovaries isolated during different stages of the estrous cycle: Effects of norepinephrine. Fertil. Steril., 27:319-327. 451. Stjemquist, M., Emson, P., Owman, C., Sjoberg, N. O., Sundler, F., and Tatemoto, K. (1983): Neuropeptide Y in the female repro¬ ductive tract of the rat. Distribution of nerve fibers and motor effects. Neurosci. Lett., 39:279-284. 452. Strickland, S., and Beers, W. H. (1976): Studies on the role of plasminogen activator in ovulation. In vitro response of granulosa cells to gonadotropins, cyclic nucleotides and prostaglandins. J. Biol. Chem., 251:5694-5702.

453. Strickland, S., Reich, E., and Sherman, M. I. (1976): Plasminogen activator in early embryogenesis. Enzyme production by trophoblast and parietal endoderm. Cell, 9:231-240. 454. Strulovici, B., Lindner, H. R., Shinitzky, M., and Zor, U. (1981): Elevation of apparent membrane viscosity in ovarian cells by follicle stimulating hormone. Biochim. Biophys. Acta, 640: 159-168. 455. Sun, F. F., Chapman, J. P., and McGuire, J. C. (1977): Metabolism of prostaglandin endoperoxide in animal tissues. Prostaglandins, 14:1055-1074. 456. Svensjo, E., and Grega, G. J. (1986): Evidence for endothelial cellmediated regulation of macromolecular permeability by post-capil¬ lary venules. Fed. Proc., 45:89-95. 457. Svensson, K. G., Owman, C., Sjoberg, N.-O., Sporrong, B., and Walles, B. (1975): Ultrastructural evidence for adrenergic innerva¬ tion of the interstitial gland in the guinea pig ovary. Neuroendocri¬ nology, 17:40-47. 458. Swanson, R. J., and Lipner, H. (1977): Mechanism of ovulation: Effect of intrafollicular progesterone antiserum. Fed. Proc., 36:390 (abstract). 459. Sweet, L. K., and Thorp, E. G. (1929): The effect of lower abdominal sympathectomy on the estrous cycle. Am. J. Physiol., 89:50-53. -460. Symons, D. B. A., and Herbert, J. (1971): Incidence of immuno¬ globulins in fluids of the rabbit tracts and the distribution of IgGglobulin in the tissues of the female tract. J. Reprod. Fertil., 24:5562. 461. Szego, C. M. (1965): Role of histamine in mediation of hormone action. Fed. Proc., 24:1343-1352. 462. Szego, C. M., and Giten, E. S. (1964): Ovarian histamine depletion during acute hyperaemic response to luteinizing hormone. Nature, 201:682-684. 463. Talbot, P., and Chacon, R. S. (1982): In vitro ovulation of hamster oocytes depends on contraction of follicular smooth muscle cells. J. Exp. Zool., 224:409-415. 464. Talbot, P., and Schroeder, P. C. (1982): 5-Hydroxytryptamine causes contraction of smooth muscle cells in preovulatory hamster follicles. J. Exp. Zool., 224:427-436. 465. Tam, W. H., and Roy, R. J. (1982): A possible role of prostaglandin F-2-a in the development of ovarian follicle in guinea pigs. J. Re¬ prod. Fertil., 66:277-282. 466. Thomson, A. (1919): The ripe human Graafian follicle, together with some suggestions as to its mode of rupture. J. Anat., 54:1^40. 467. Thorsoe, H. (1962): Hexuronic acid, hexosamine and radiosulphate uptake in ovaries of myxoedematous rabbits. Acta Endocrinol. (Copenh.), 41:613-618. 468. Tjugum, J., Norstrom, A., and Dennefors, B. (1983): Influence of prostaglandin E2 on proteoglycan synthesis in the human ovarian follicle wall. Prostaglandins, 25:71-77. 469. Too, C. K. L., Weiss, T. J., and Bryant-Greenwood, G. D. (1982): Relaxin stimulates plasminogen activator secretion by rat granulosa cells in vitro. Endocrinology, 111:1424-1426. 470. Too, C. K. L., Bryant-Greenwood, G. D., and Greenwood, F. C. (1984): Relaxin increases the release of plasminogen activator, collagenase and proteoglycanase from rat granulosa cells in vitro. En¬ docrinology, 115:1043-1050. 471. Tranzer, J. P., and Thoenen, H. (1968): An electron microscopic study of selective acute degeneration of sympathetic nerve terminals after administration of 6-hydroxy dopamine. Experientia 24-155156. 472. Triebwasser, W. F., Clark, M. R., LeMaire, W. J., and Marsh, J. M. (1978): Localization and in vitro synthesis of prostaglandins in components of rabbit preovulatory Graafian follicles. Prostaglan¬ dins, 16:621-632. 473. Tsafriri, A., Lindner, H. R., Zor, U., and Lamprecht, S. A. (1972): Physiological role of prostaglandins in the induction of ovulation. Prostaglandins, 2:1-10. 474. Tsafriri, A., Lindner, H. R., Zor, U., and Lamprecht, S. A. (1972): In vitro induction of meiotic division in follicle-enclosed rat oocytes by LH, cyclic AMP and prostaglandin E2. J. Reprod. Fertil., 31:39475. Tsafriri, A., Koch, Y., and Lindner, H. R. (1973): Ovulation rate and serum LH levels in rats treated with indomethacin or PGE2. Prostaglandins, 3:461-467.

Mammalian Ovulation

476. Tsang, B. K., Ainsworth, L., Downey, B. R., and Armstrong, D. T. (1979): Preovulatory changes in cyclic AMP and prostaglandin concentrations in follicular fluid of gilts. Prostaglandins, 17:141— 148. 477. Tsujimoto, D., Katayama, K., Tojo, S., and Mizoguti, H. (1982): Scanning electron microscopic studies on stigmas in rat ovaries. Acta Obstet. Gynecol. Scand., 61:269-273. 478. Tuohimaa, P., and Niemi, M. (1969): Cell renewal in the ovarian follicles of the rat during the oestrous cycle and in persistent estrus. Acta Endocrinol., 62:306-314. 479. Vassalli, J-D., Hamilton, J., and Reich, E. (1976): Macrophage plasminogen activator: Modulation of enzyme production by anti¬ inflammatory steroids, mitotic inhibitors and cyclic nucleotides. Cell, 8:271-281. 480. Virutamasen, P., Hickok, R. L., and Wallach, E. E. (1971): Local ovarian effects of catecholamines on human chorionic gonadotrophininduced ovulation in the rabbit. Fertil. Steril., 22:235-243. 481. Virutamasen, P., Smitarsiri, Y., and Fuchs, A. R. (1976): Intraovarian pressure changes during ovulation in rabbits. Fertil. Steril., 27:188-196. 482. Virutamasen, P., Wright, K. H., and Wallach, E. E. (1972): Effects of catecholamines on ovarian contractility in the rabbit. Obstet. Gy¬ necol., 39:225-236. 483. Virutamasen, P., Wright, K. H., and Wallach, E. E. (1972): Effects of prostaglandins E2 and F2a on ovarian contractility in the rabbit. Fertil. Steril., 23:675-682. 484. Virutamasen, P., Wright, K. H., and Wallach, E. E. (1973): Monkey ovarian contractility: Its relationship to ovulation. Fertil. Steril., 24:763-771. 485. Wada, H. (1978): The biochemical and biological properties of bo¬ vine follicular fluid compared with blood serum. Bull. Azabu Vet. Coll., 3:313-331. 486. Wada, H. (1981): Effects of gonadotropins steroid hormones and histamine on the blood follicle barrier of the ovary. Med. J. Kobe Univ., 42:57-66. 487. Waldeyer, W. (1870): Eierstock and Ei. von Wilhelm Englemann, Leipzig. 488. Wallach, E. E., Wright, K. H., and Hamada, Y. (1978): Investi¬ gation of mammalian ovulation with an in vitro perfused rabbit ovary preparation. Am. J. Obstet. Gynecol., 132:728-738. 489. Walks, B., Edvinsson, L., Falck, B., Nybell, G., Owman, C., Sjoberg, N. O., and Svensson, K. G. (1974): Modifications of ovar¬ ian and follicular contractility by amines. A mechanism involved in ovulation. Eur. J. Obstet. Gynecol. Reprod. Bio., 4/1 (Suppl.): S103-S107. 490. Walles, B., Edvinsson, L., Owman, C., Sjoberg, N. O., and Svens¬ son, K. G. (1975): Mechanical response in the wall of ovarian fol¬ licles mediated by adrenergic receptors. J. Pharmacol. Exp. Ther., 193:460—473. 491. Walles, B., Edvinsson, L., Falck, B., Owman, C., Sjoberg, N. O., and Svensson, K. G. (1975): Evidence for a neuromuscular mech¬ anism involved in the contractility of the ovarian follicular wall. Fluorescence and electron microscopy and effects of tyramine on follicle strips. Biol. Reprod., 12:239-248. 492. Walles, B., Edvinsson, L., Owman, C., Sjoberg, N. O., and Sporrong, B. (1976): Cholinergic nerves and receptors mediating con¬ traction of the graafian follicle. Biol. Reprod., 15:565-572. 493. Walles, B., Falck, B., Owman, C., and Sjoberg, N. O. (1977): Characterization of autonomic receptors in the smooth musculature of human Graafian follicle. Biol. Reprod., 17:423—431. 494. Walles, B., Owman, C., and Sjoberg, N. O. (1982): Contraction of the ovarian follicle induced by local stimulation of its sympathetic nerves. Brain Res. Bull., 9:757-760. 495. Walton, A., and Hammond, J. (1928): Observations on ovulation in the rabbit. J. Exp. Biol., 6:190-204. 496. Wang, C., and Leung, A. (1983): Gonadotropins regulate plasmin¬ ogen activator production by rat granulosa cells. Endocrinology, 112:1201-1207. 497. Weiner, S., Wright, K. H., and Wallach, E. E. (1975): Selective ovarian sympathectomy in the rabbit. Fertil. Steril., 26.353-362. 498. Weiner, S., Wright, K. H., and Wallach, E. E. (1975): Studies on the function of the denervated rabbit ovary: Human chorionic go¬ nadotrophin-induced ovulation. Fertil. Steril., 26:363-368.

/

487

499. Weiner, S., Wright, K. H., and Wallach, E. E. (1975): Lack of effect of ovarian denervation on ovulation and pregnancy in the rabbit. Fertil. Steril., 26:1083-1089. 500. Weiner, S., Wright, K. H., and Wallach, E. E. (1977): The influence of ovarian denervation and nerve stimulation on ovarian contractions. Am. J. Obstet. Gynecol., 128:154-160. 501. Werb, Z., Mainardi, C. L., Vater, C. A., and Harris, E. O., Jr. (1977): Endogenous activation of latent collagenase by rheumatoid synovial cells. N. Engl. J. Med., 296:1017-1023. 502. White, I. C. (1943): Sensory innervation of the viscera. Studies on visceral afferent neurones in man based on neurosurgical procedures for the relief of intractable pain. Res. Publ. Assoc. Res. Nerv. Ment. Dis., 23:373-390. 503. Wilhelmsson, L., Norstrom, A., and Hamberger, L. (1981): Influ¬ ence of 6-keto-PGF|a on collagen synthesis in the human cervix during various phases of the menstrual cycle. Prostaglandins, 22:125130. 504. Williams, J. R. B. (1951): The fibrinolytic activity of urine. Br. J. Exp. Pathol., 32:530-537. 505. von Winiwarter, H. (1910): Contribution a l’etude de l’ovarire humain. Arch. Biol.. 25:683-755. 506. von Winiwarter, H., and Sainmont, G. (1909): Nouvelles recherches sur l’ovogenese et l’organogenese de l’ovaire des Mammiferes Chat. Arch. Biol. (Liege), 24:627-651. 507. Winterhalter, E. H. (1896): Ein sympathisches Ganglion in menschichen Ovarian. Arch. Gynecol., 51:49-55. 508. Wischnitzer, S. (1965): The ultrastructure of the germinal epithelium of the mouse ovary. J. Morphol., 117:387—400. 509. Wislocki, G. B., Bunting, H., and Dempsey, E. W. (1947): Metachromasia in mammalian tissues and its relationship to mucopoly¬ saccharides. Am. J. Anat., 81:1-37. 510. Woessner, J. F., Ir. (1982): Enzymatic mechanisms for the degra¬ dation of connective tissue matrix. In: Symposium on Idiopathic Low Back Pain, edited by A. White and S. L. Gordon, pp. 391-400. Mosby, St. Louis. 511. Woolley, D. E., and Evanson, I. M. (1980): Present status and fu¬ ture prospects in collagenase research. In: Collagenase in Nor¬ mal and Pathobiological Connective Tissues, edited by D. E. Woolley and J. M. Evanson, pp. 241-250. John Wiley & Sons, New York. 512. Wright, K. H., Wallach, E. E., Fromm, E., and Jeutter, D. C. (1976): Studies of rabbit ovarian contractility using chronically im¬ planted transducers. Fertil. Steril. 27:310-318. 513. Wurtman, R. J. (1964): An effect of luteinizing hormone on the fractional perfusion of the rat ovary. Endocrinology, 75:927-933. 514. Yanagishita, M., and Hascal, V. C. (1979): Biosynthesis of proteo¬ glycans by rat granulosa cells cultured in vitro. J. Biol. Chem., 254:12355-12364. 515. Yanagishita, M., Rodbard, D., and Hascal, V. C. (1979): Isolation and characterization of proteoglycans from porcine ovarian follicular fluid. J. Biol. Chem., 254:911-920. 516. Yanagishita, M., Hascal, V. C., and Rodbard, D. (1981): Biosyn¬ thesis of proteoglycans by rat granulosa cells cultured in vitro: Mod¬ ulation by gonadotropins, steroid hormones prostaglandins, steroid hormones, and prostaglandins and a cyclic nucleotide. Endocrinol¬ ogy, 109:1641-1649. 517. Yang, N. S. T., Marsh, J. M., and LeMaire, W. J. (1973): Prostaglandin changes induced by ovulatory stimuli in rabbit Graafian follicles. The effect of indomethacin. Prostaglandins, 4:395-404. 518. Zachariae, F. (1957): Studies on the mechanism of ovulation: Au¬ toradiographic investigations on the uptake of radioactive sulphate (35S) into the ovarian mucopolysaccharides. Acta Endocrinol. (Copenh.), 26:215-223. 519. Zachariae, F. (1958): Studies on the mechanism of ovulation. Per¬ meability of the blood-liquor barrier. Acta Endocrinol., 27:339-342. 520. Zachariae, F., and Jensen, C. E. (1958): Studies on the mechanism of ovulation. Histochemical and physico-chemical investigations on genuine follicular fluids. Acta Endocrinol., 27:343-355. 521. Zoller, L. C., and Weisz, J. (1979): A quantitative cytochemical study of glucose-6-phosphate dehydrogenase and A5-3(3-hydroxysteroid dehydrogenase activity in membrana granulosa of the ovulable type of follicle of the rat. Histochemistry, 62:125-132.

488

/

Chapter 12

522. Zoller, L. C., and Weisz, J. (1980): A demonstration of regional differences in lysosome membrane permeability in the membrana granulosa of Graafian follicles in cryostat sections of the rat ovary: A quantitative cytochemical study. Endocrinology, 106:871-877. 523. Zondek, B., Sulman, F., and Black, R. (1945): The hyperemia ef¬ fect of gonadotropins on the ovary. J. Am. Med. Assoc., 128: 939-944.

524. Zor, U., Strulovici, B., Braw, R., Lindner, H. R., and Tsafriri, A. (1983): FSH induced prostaglandin E formation in isolated rat ovarian theca. J. Endocrinol., 97:43-50. 525. Zsolmar, B.f Varga, B., and Horvath, E. (1982): Increase of ovarian progesterone secretion by B2-adrenergic stimulation in oestrous rats. Acta Endocrinol. (Copenh.), 101:268-272. 526. McCaig, C. D. (1985): A potential difference across mouse ovarian follicle. Experientia 41:609-611.

The Physiology of Reproduction, edited by E. Knobil and J. Neill et al. Raven Press, Ltd., New York © 1988.

CHAPTER 13

The Corpus Luteum and Its Control Gordon D. Niswender and Terry M. Nett lation of Luteal Function, 495 • Regulation of Proges¬ terone Secretion, 497

Biological Functions of Progesterone, 489 Reproductive Tract, 489 • Mammary Gland, 491 • Hypothalamus and Anterior Pituitary Gland, 491 • Other Hormones Produced by the Corpus Luteum, 491 Formation of the Corpus Luteum, 491

Luteolysis, 504 Uterine Involvement in Luteal Regression, 504 • Identi¬ fication of PGF2a as the Uterine Luteolytic Factor, 505

Morphological Changes Associated with Luteinization,

Luteal Function during Pregnancy, 507 Maternal Recognition of Pregnancy, 507

491 • Ontogeny of Luteal Receptors for Hormones ,492 Luteal Phase of the Estrous Cycle, 493 Morphology of Luteal Cells, 493 • Hypophyseal Regu¬

References, 514

would have a dramatic impact on increasing the production of food animals.

The corpus luteum is a transient endocrine organ formed from cells of the follicle following ovulation (Fig. 1). That the corpus luteum is required for a successful pregnancy was first discovered early in the twentieth century by Frankel (1), who found that pregnancy in rabbits was terminated following removal of corpora lutea. Similar findings have since been reported in many different species of mammals. Even though the requirement for the corpus luteum during normal pregnancy was documented in 1903, the nature of the substance that the corpus luteum produced to maintain pregnancy remained unknown for some two decades. In 1929, Allen and Comer (2) showed that a lipoidal extract of the corpus luteum could maintain pregnancy in rabbits ovariectomized a few days after mating. The component in the extract responsible for the maintenance of pregnancy was subsequently called progesterone. Its purification and crystallization was first achieved in the laboratory of Allen and Wintersteiner in 1934 (3). Since that time, there has been a major interest in understanding the factors that reg¬ ulate the life span and function of the corpus luteum. De¬ velopment of methods to limit the function of this gland should have a major effect on limiting reproduction in hu¬ mans, rodents and pet animals. In addition, 25% to 55% of all mammalian embryos are lost during early gestation and much of this loss appears to be due to inadequate luteal function. Development of procedures to prevent this loss

BIOLOGICAL FUNCTIONS OF PROGESTERONE The primary function of the corpus luteum is to secrete progesterone. Progesterone has several biological effects on target tissues in the reproductive system to prepare them for support of pregnancy or to provide nourishment to the conceptus. The following is a brief description of the general effects of progesterone on the reproductive organs of the female during the estrous cycle and in early pregnancy. It is not intended to provide a comparative description of the actions of progesterone in a variety of species.

Reproductive Tract A primary target for progesterone is the mucosal lining of the genital tract. For progesterone to affect the genital tract, the cells must first have been exposed to estradiol, which induces the formation of receptors for progesterone (4). Estrogens act on the oviductal epithelium to promote growth and proliferation of the cells and to induce ciliogenesis. Following ovulation, cilitated cells in the fimbriated end of the oviduct appear to direct the ovum and associated cumulus cells into the infundibulum and then downward toward the ampulla. Together, estrogen and progesterone also regulate contractions of the oviduct that influence the rate of transport of the ovum to the uterus (5). These con-

The authors thank Dr. Heywood Sawyer for the electron micrographs and Ms. Kathy Miller for typing this manuscript.

489

490

/

Chapter 13

S** F04LL.ICI.E

uurrwum FIG. 1. Composite diagram of the mammalian ovary. Progres¬ sive stages in follicular growth and development are depicted. The primary (1°) follicle has a single layer of granulosal cells surrounding the ovum. The secondary (2°) follicle has multiple layers of granulosal cells, and the theca layers begin to dif¬ ferentiate. The tertiary (3°) follicle has fully developed granu¬ losal, theca interna, and theca externa layers, and an antrum tractions are coordinated in a manner that results in the delivery of the conceptus to the uterus at an appropriate time. Progesterone contributes to the secretory activity of the oviductal epithelium to induce the secretion of a fluid that supports the early development of the conceptus. Pro¬ gesterone also appears to induce regression of the oviductal epithelium, which is characterized by a sloughing of the apical portion of the epithelial cells (6). One of the most significant actions of progesterone is to prepare the uterus for pregnancy. The uterine mucosa is composed of a surface layer of columnar epithelial cells overlying a stroma composed of spindle-shaped cells per¬ meated by blood vessels. Numerous glands permeate the surface layer and dip into the stroma. These glands are lined by columnar epithelial cells, some of which are ciliated, whereas others are nonciliated and appear to be secretory. Under the influence of estradiol, the uterine mucosa thickens due to mitotic proliferation of the epithelium and stroma, and the tubular glands lengthen but remain straight. Nuclei in the epithelial cells tend to be located near the center of the cells. Progesterone, secreted by the developing corpus luteum, inhibits cell division, induces a marked coiling of the glands, and increases the vascularity of the stroma sub¬ stantially (7). There is a dramatic reduction in cell prolif¬ eration and an increase in glycogen content of the epithelial

has formed and filled with follicular fluid. The Graafian, or ovu¬ latory, follicle continues to grow until ovulation occurs. The cavity of the ovulated follicle fills with blood (the corpus hemorrhagicum) and the corpus luteum develops after a number of morphological and biochemical changes that are described in the text.

cells. The nuclei of the epithelial cells become more basally oriented and their cytoplasm becomes vacuolar. These changes prepare the endometrial cells to provide nourishment and support to the conceptus until attachment (or implantation) and placentation occurs. In many species, progesterone also acts on the cells of the myometrium to block organized contractions. During nonfertile cycles, the progestational endometrium undergoes deterioration when serum levels of progesterone decrease as the corpus luteum ceases to function. Although the uteri of all species undergo regressive changes as serum levels of progesterone fall, it is most dramatic in primates, in which the lining of the uterus actually is sloughed and results in menstruation. Under the influence of estrogen, the cervix secretes a mucus rich in glycoprotein that appears to align in filaments, facilitating passage of sperm through the cervical canal. Under the influence of progesterone, the consistency of the cervical mucus changes and becomes highly viscous. The glycoprotein filaments form networks that impede passage of materials either into or out of the uterus. Thus, when circulating levels of progesterone are high, the cervix pro¬ vides an effective barrier between the uterus and the external environment (7). Estrogens promote proliferation and comification of the

Corpus Luteum / vaginal epithelium. As the vaginal epithelium thickens, the superficial layers are not close to blood vessels; this leads to keratinization and loss of nuclei. These cells are then sloughed off. This trend is reversed by progesterone. Under the influence of progesterone, the vaginal epithelium thins and is characterized by small nucleated cells. The thinness of the vaginal epithelium allows escape of leukocytes, which may appear in vaginal smears from animals under the in¬ fluence of progesterone. These changes have been most completely characterized in rodents (8) and dogs (9) but occur to a lesser extent in other species as well.

Mammary Gland Besides its effects on the female reproductive tract, pro¬ gesterone also acts to promote lobuloalveolar development of the mammary glands. In order for progesterone to be effective in this regard, the mammary gland must have been exposed to estrogen, and anterior pituitary hormones must also be present, i.e., progesterone alone has little, if any, effect on mammary tissue (10). Progesterone receptors are present in the mammary gland, but little is known con¬ cerning their role in the regulation of mammary gland growth or lactogenesis. During estrous (or menstrual) cycles, pro¬ gesterone appears to have little effect on mammary gland growth. However, during the prolonged secretion of pro¬ gesterone that occurs during pregnancy, there is consider¬ able development of the lobuloalveolar system. The extent to which this development can be credited to progesterone versus that which is due to stimulation by placental lactogen and prolactin has not been resolved.

Hypothalamus and Anterior Pituitary Gland Under the influence of estradiol during the follicular phase of the cycle, gonadotropin secretion is characterized by de¬ creasing serum concentrations of follicle-stimulating hor¬ mone (FSH) and increasing concentrations of luteinizing hormone (LH) in most species. The increasing baseline con¬ centrations of LH result from secretion of low-amplitude, high-frequency pulses of LH during the follicular phase of the cycle (11-14). During the luteal phase of the cycle, when serum levels of progesterone are elevated, the pattern of gonadotropin secretion changes dramatically. Pulses of LH are infrequent and are of higher amplitude than those observed during the follicular phase of the cycle. This ap¬ pears to be due to a direct influence of progesterone on the “gonadotropin-releasing hormone pulse generator” in the hypothalamus. Secretion of FSH is not greatly affected by progesterone. As a result, in those species in which the corpus luteum does not secrete estradiol the secretion of FSH is rather stable at a relatively high baseline during the luteal phase of the cycle. In primates the corpus luteum secretes estradiol, and basal concentrations of FSH remain

491

suppressed during the luteal phase and do not increase until after regression of the corpus luteum (13,15).

Other Hormones Produced by the Corpus Luteum In addition to progesterone, the corpus luteum also se¬ cretes a variety of other hormones. In primates, the corpus luteum is a source of estrogens, but this does not appear to be the case in most other species. In several species, in¬ cluding human (16), pig (17), and rat (18), the corpus lu¬ teum produces relaxin. The function of ovarian relaxin is unclear. It may act synergistically with estrogen and pro¬ gesterone to prepare the endometrium for implantation and development of the blood supply to the conceptus in pri¬ mates. Near term, several changes in the birth canal that are required for parturition are induced by relaxin; however, in most species the fetoplacental unit appears to be the major source of relaxin in late gestation. Corpora lutea from a variety of species also contain oxytocin (19,20). The bio¬ logical function of luteal oxytocin is unknown, but an in¬ volvement in luteolysis has been suggested (21). There is also evidence that the corpus luteum from some species, particularly primates, can secrete the prostaglandin PGF2a (22,23).

FORMATION OF THE CORPUS LUTEUM Morphological Changes Associated with Luteinization Formation of the corpus luteum is initiated by a series of morphological and biochemical changes in the cells of the theca interna and membrana granulosa of the preovulatory follicle. This is termed luteinization. These changes occur as a result of the dramatic increases in serum levels of LH associated with the preovulatory surge of this hormone. The most complete description of the morphological changes associated with luteinization in the rat is that of Anderson and Little (24). Following the ovulatory stimulus, but prior to ovulation, there is hypertrophy of granulosal cells and nuclear activation. After ovulation, the basement membrane breaks down, and blood vessels from the theca interna invade the cavity of the ruptured follicle. The growth of these new vessels appears to be due to an angiogenic factor that must be secreted soon after rupture of the follicle. In rats, the number and size of gap junctions between gran¬ ulosal cells increases as the follicle matures but then de¬ creases just prior to ovulation. During luteinization, many gap junctions reappear between the developing luteal cells in rats. As will be discussed later, the presence of gap junctions between luteal cells during the reproductive cycle is not a universal phenomenon. Gonadotropic hormones appear to amplify and modulate, rather than to induce, the formation of gap junctions. The appearance of gap junctions may be dependent upon intracellular levels of cAMP (25). Cytoplasmic projections, which are characteristic of rat

492

/

Chapter 13

granulosal cells, are also found on luteal cells. However, the rather thin microvilli frequently seen on granulosal cells are not seen on luteal cells. These cytoplasmic projections contain receptors for LH (26,27) and have septate-like junc¬ tions between them (28). All of the cells of the follicle do not differentiate synchronously following ovulation. The amount of smooth endoplasmic reticulum and the number of mitochondria both increase during luteinization. Mito¬ chondria in granulosal cells have lamellar cristae, while those in luteal cells have primarily tubular cristae. Luteal cells during early and midpregnancy in rats also contain lipid droplets and well-developed Golgi complexes. In the ewe, the first signs of luteinization occur prior to ovulation. Dispersion of nuclear chromatin and formation of a nucleolus occurs with a concomitant increase in the number of polyribosomes. This suggests that RNA and pro¬ tein synthesis are important at this stage. There are numerous gap junctions between adjacent granulosal cells in the pre¬ ovulatory follicle, along with desmosome-like structures (29). The formation of smooth endoplasmic reticulum and alter¬ ations in mitochondria occur 30 to 40 hr after the ovulatory surge of LH. The elaboration of smooth endoplasmic retic¬ ulum is necessary for normal biosynthesis of steroids and is the site of localization of 3|3-hydroxysteroid dehydro¬ genase, which is acquired by the granulosal cells within a few hours following ovulation (30). This is the first time that these cells are capable of synthesizing progesterone. In addition, after ovulation the mitochondria become rounded and develop villiform cristae. The development of the smooth endoplasmic reticulum and mitochondria in granulosal-luteal cells is correlated with the initial rise in serum levels of progesterone. Gap junctions are no longer apparent in the ovine corpus luteum by 48 hr after ovulation. Based on detailed morphological studies of the ovulatory follicle and developing corpus luteum, it has been concluded that gran¬ ulosal cells differentiate into the large steroidogenic luteal cells found in the mature corpus luteum in ewes (29,31) and cows (32). In the ewe, the cells of theca interna in the preovulatory follicle possess abundant lipid droplets. Alkaline phospha¬ tase and 3p-hydroxysteroid dehydrogenase are restricted to this cell type prior to ovulation (31). In contrast to granulosal cells, only occasional mitotic figures are seen in the cells of the theca interna. Within 24 hr of ovulation, cells derived from the theca interna begin migrating from their original sites into the deeper, granulosal-derived areas of luteal tissue (33). At later stages, cells derived from the theca interna remain concentrated in the septa derived from follicular infoldings but are also widely distributed throughout the luteal tissue.

Ontogeny of Luteal Receptors for Hormones Since luteinization and formation of the corpus luteum is simply a natural extension of follicular growth and ovula¬

tion, this section will begin with a discussion of the changes that occur in receptors during follicular development. The induction and first appearance of receptors for LH during the course of follicular development has been intensely stud¬ ied, particularly in granulosal cells. Granulosal cells ob¬ tained from immature rat or pig follicles possess few, if any, receptors for LH (34,35). However, as follicles mature and increase in size, the number of receptors for LH in¬ creases dramatically (36-40). On the basis of studies that have employed estradiol-FSH-primed, hypophysectomized rats, it appears that the initial appearance and subsequent increase in the number of receptors for LH in granulosal cells is a result of the synergistic action of estrogen and FSH (39,41). According to Richards et al. (39): (a) Estra¬ diol- 17(3 acts on granulosal cells to increase the concentra¬ tion of its own receptor and induces receptors for FSH; (b) FSH acts on estrogen-primed granulosal cells to increase ^receptors for both FSH and LH; and (c) LH acts on estrogenFSH-primed cells to effect a decrease in receptors for es¬ tradiol, FSH, and LH and at the same time promote an increase in the number of receptors for prolactin. Data from studies that have employed intact, normally cycling rats also indicate that the appearance of an increased number of re¬ ceptors for LH is a result of the combined action of estrogen and FSH (42). Once granulosal cells acquire receptors for LH, they are rendered sensitive to LH and are capable of undergoing luteinization. As demonstrated by Richards et al. (43), the ability of LH to induce luteinization is related to its ability to increase the intracellular concentrations of cAMP. In hypophysectomized rats administration of LH or human chorionic gonadotropin (hCG) to cause ovulation and luteinization of follicular cells is followed by a decrease (“down-regulation”) in the number of LH receptors on gran¬ ulosal cells. However, since Nimrod et al. (42) were unable to detect any significant loss of receptors for LH under physiological conditions, the reason for down-regulation of LH receptors observed during LH-induced luteinization in hypophysectomized animals (39,43,44) is uncertain. The pattern of development of receptors for LH in fol¬ licular granulosal cells in pigs (37) and sheep (45) appears to be similar to that described for rats. There is little in¬ formation regarding receptors for LH in follicular thecal cells. In sheep it appears that receptors for LH appear first in the thecal cells in small follicles and that as the follicle enlarges there is a slight decline in the capacity of thecal cells to bind LH concomitant with a dramatic increase in LH binding to granulosal cells (45). In rats, prolactin is necessary to maintain normal numbers of receptors for LH in the developing corpus luteum (41). This role is consistent with the known biological effects of prolactin on luteal function in this species. However, the situation in other species is not so clear, since prolactin does not appear to influence luteal function in cattle (46-49) or sheep (48,49). Although receptors for prolactin have been reported in porcine luteal tissue (50), the exact role of this hormone in regulating luteal receptors for LH in species

Corpus Luteum / other than rodents will require additional experimentation. In porcine luteal cells, prolactin appears to influence the number of receptors for low-density lipoprotein (51), and it may therefore play a role in regulating substrate avail¬ ability for steroidogenesis.

LUTEAL PHASE OF THE ESTROUS CYCLE Morphology of Luteal Cells In the late 1800s and early 1900s there were two hy¬ potheses concerning the follicular cell type responsible for formation of the corpus luteum. The first held that the corpus luteum was derived exclusively from granulosal cells of the follicle while the cells from the theca interna degenerated shortly after ovulation. The second hypothesis held exactly the opposite view. In 1906, Loeb (52) performed a detailed morphological analysis of the ovary in the guinea pig during the period of luteinization and concluded that cells from both the theca interna and granulosal layers of the ovarian follicle were involved in formation of the corpus luteum. Comer (53) came to the same conclusion concerning the corpus luteum of the sow, as have subsequent investigators examining corpora lutea in cows (32,54-56), ewes (31), rats (57), and women (58). Since the cells of the corpus luteum appear to be derived from at least two different types of follicular steroid-se¬ creting cells (thecal and granulosal cells), it is not surprising that the corpus luteum consists of at least two distinct types of steroidogenic luteal cells in several species (53,59-61). The two types of steroidogenic cells are morphologically distinct. The so-called large luteal cells (32,33,56,62-66)— also referred to as granulosa-lutein (60,67), Type II (68), and D cells (69)—are the most readily distinguished cells in the corpus luteum (Fig. 2). They are the largest, strictly endocrine cells in the body and range from approximately 20 p.m in diameter in rodents to 40 p,m or more in humans (70). Under the light microscope, large luteal cells appear polyhedral, with a lightly staining cytoplasm and a large,

493

centrally located nucleus with a distinct nucleolus. In con¬ trast, small luteal cells have a diameter of 22 |xm or less and appear spindle-shaped with darkly staining cytoplasm, large lipid droplets, and an irregularly shaped nucleus that often contains what has been described as cytoplasmic in¬ clusion. Alternative terms for small luteal cells include theca¬ lutein (60,67), Type I (68), and I cells (69). On a volume basis, large luteal cells comprise about 25% to 35% of the corpus luteum in sheep, whereas small luteal cells represent approximately 12% to 18% of the luteal volume (71,72). The corpus luteum also contains vascular elements and con¬ nective tissue. During the period of maximum secretion of progesterone, vascular elements account for approximately 11% of luteal volume (71). The remainder of the corpus luteum is composed of connective tissue (22-29%) and fi¬ broblasts (7-11%) (71,72). Ultrastructurally, large luteal cells contain all of the ele¬ ments of steroid-secreting cells, that is, numerous mito¬ chondria and an abundance of smooth endoplasmic reticu¬ lum (Fig. 3) (73). Mitochondria in large luteal cells can take on a variety of shapes: spherical, cup-shaped, or elon¬ gated (70); in fact, it is not uncommon to find each of these shapes of mitochondria within a single cell. There is also considerable variation in the size of mitochondria in large luteal cells. In most species, mitochondria with tubular cristae predominate, but some with lamelliform cristae may also be observed. Clustering and regional exclusion of mito¬ chondria are also common features noted in large luteal cells (70). Large luteal cells have an abundance of smooth endo¬ plasmic reticulum, which is characteristically most prevalent in the peripheral region of the cell. In fact, it is often the most abundant cytoplasmic component found at the cell’s periphery. The smooth endoplasmic reticulum is in the form of branched tubules, tubular sheets, and fenestrated cisternae. Often the fenestrated cistemae are centered around a mitochondrion or lipid droplet. The branched tubules may form an anastomotic network that traverses the entire cell. The Golgi complex is quite extensive in large luteal cells and is usually located at one side of the nucleus and occupies

FIG. 2. Large luteal cells (LLC) can be easily distin¬ guished from small luteal cells (SLC) in 1 -p,m-thick sec¬ tions stained with toluidine blue. Small luteal cells are usually spindle-shaped, whereas LLC are typically spherical or polyhedral. In addition, the dark-staining cytoplasm of SLC contains large lipid droplets. The nuclei of SLC sometimes possess cytoplasmic inclu¬ sions (arrow). Both cell types are in close apposition to capillaries (CAP). x800. (From ref. 61.)

494

/

Chapter 13

FIG. 3. Electron micrograph showing a portion of a large luteal cell (LLC). The presence of numerous electron-dense, mem¬ brane-bound secretory granules (SG) in the cytoplasm of LLC distinguishes these cells from small luteal cells (SLC) (see Fig. 5). Note the absence of lipid droplets comparable to those found in SLC. Mitochondria (M), nucleolus (N), rough en¬ doplasmic reticulum (RER). x 7,200. (From ref. 61.)

a comparable volume. There is a paucity of tubular smooth endoplasmic reticulum and mitochondria in the area of the Golgi complex. Unlike other cell types, there is an indication that the smooth endoplasmic reticulum is in direct com¬ munication with the Golgi cistemae (74). There are abundant electron-dense, membrane-bound se¬ cretory granules in the cytoplasm of large luteal cells (Fig. 3). These granules are similar in size to lysosomes and peroxisomes, but they are a distinct form of granule (75) and their contents do not include acid phosphatase or cat¬ alase activity (75-77). Contents of the granules are released at the surface of the cell by exocytosis (Fig. 4) (78-80). The exocytotic release of granular contents is correlated with the secretion of progesterone both in vivo (76) and in vitro (80). Based on these observations, several investigators pro¬ posed that these granules contain progesterone (and a protein that binds progesterone) and provide a mechanism for the "active secretion of progesterone from the luteal cell (7682). However, at the current time this hypothesis seems unlikely since small steroidogenic luteal cells have few, if any, secretory granules. Thus, the mechanisms for secretion of progesterone would have to be different in the two cell types. In contrast, others argue that progesterone exits the cell by simple diffusion due to a concentration gradient between intracellular concentrations and those in the blood (70,83). At least one of the components in secretory granules in large luteal cells in cattle (84) and sheep (20,85) is oxy¬ tocin. Relaxin has also been identified in similar granules in rats (86,87), pigs (88-90), and cows (91). It is possible that the granules may contain more than one secretory prod¬ uct or that there are different types of granules within the same cell. The ultrastructural appearance of small luteal cells is dis¬ tinct from that of large luteal cells (Fig. 5). The nucleus is irregular in shape, and in approximately 10% of the cells it

FIG. 4. Both small (lower left) and large (upper right) luteal cells possess fine structural char¬ acteristics consistent with a steroid-secreting function, i.e., extensive smooth endoplasmic re¬ ticulum (SER), lipid droplets (L), and Golgi ap¬ paratus (G). However, LLC also have character¬ istics typical of protein-secreting cells, including secretory granules (SG), some of which have been exocytosed. Large luteal cells possess a more conspicuous basal lamina (BL) than do small luteal cells, x 14,800. (From ref. 61.)

Corpus Luteum /

495

present in the large luteal cell. Small luteal cells also contain numerous lipid droplets, which are virtually absent in large luteal cells.

Hypophyseal Regulation of Luteal Function In most species three organ systems regulate the function of the corpus luteum. The anterior pituitary gland secretes LH, which is the primary hormone responsible for regulating the secretion of progesterone. In several rodent species, hypophyseal prolactin is also an important regulator of luteal function. In most nonprimate species the uterus has a luteolytic effect during the late luteal phase of the estrous cycle. Finally, the conceptus has either direct or indirect luteotropic effects in most mammalian species.

Rats

FIG. 5. Electron micrograph showing a portion of a small luteal cell. The cytoplasm is characterized by an abundance of smooth endoplasmic reticulum (SER), numerous mito¬ chondria (M), and lipid droplets. Residual bodies (RB) are often observed in these cells. Besides the nucleolus (N), the nucleus contains what appears to be a cytoplasmic inclusion (I). x 7,200. (From ref. 61.)

appears to contain areas of cytoplasm bounded by a com¬ pletely inverted nuclear envelope (Fig. 5) (33). However, to date no one has serially sectioned the nucleus of the small luteal cell to establish whether these structures represent true cytoplasmic inclusions, or rather, if they are simply cytoplasmic invaginations into the nucleus (Fig. 4). The characteristic small luteal cell contains a moderate number of mitochondria of variable size. Their profile may appear as round, elongated, or branching. Both tubular and lamelliform cristae have been observed. Large amounts of endoplasmic reticulum are present in small luteal cells. The endoplasmic reticulum is predominantly of the smooth, tu¬ bular type, but scattered clusters of attached ribosomes are also present. The Golgi complex is less pronounced in the small luteal cell than in large luteal cells, is located perinuclearly, and is associated with numerous small coated or uncoated vesicles (33). A characteristic feature of the small luteal cell is the absence of the secretory granules that are

Rothchild (92) has recently reviewed the factors that reg¬ ulate the function of the corpus luteum in most species, with particular attention to the rat. In rats, the secretion of pro¬ gesterone during the estrous cycle is short-lived and appears to be autonomous the first day of diestrus but requires pro¬ lactin during the second day (93). Postovulatory secretion of progesterone in rats has a pattern similar to the respon¬ siveness of adenylate cyclase to LH (94). Although some minor differences may be present, it appears that the en¬ docrine events controlling the life span and function of the corpus luteum during the estrous cycle in other rodents such as mice, hamsters, and gerbils are similar to those described here for the rat. The one exception appears to be that pro¬ lactin, LH, and FSH are required for normal secretion of progesterone in hamsters (59).

Rabbits Hormonal regulation of luteal function in the rabbit has been reviewed by Hilliard (96) and by Keyes et al. (97). In rabbits, ovulation is induced by cervical stimulation, which causes the preovulatory surge of LH that results in ovulation. Maintenance of the structure and function of the corpus luteum requires estradiol (97). Despite a dependence on estradiol for maintenance of the corpus luteum, rabbit luteal tissue exhibits an acute steroidogenic response to LH (98), contains high-affinity receptors for LH (99), and possesses LH-sensitive adenylate cyclase (94). The corpus luteum of the rabbit contains two cell types: with small luteal cells that are responsive to LH and large cells that do not respond to LH with enhanced secretion of progesterone (100). There is no direct steroidogenic response to estradiol. Thus, the mechanisms whereby estradiol maintains the corpus luteum are not clear.

496

/

Chapter 13

Domestic Ruminants There has been considerable controversy regarding the requirement of the anterior pituitary gland for the regulation of luteal function in ewes and cows. In 1963, Denamur and Mauleon (101) reported that formation and maintenance of the corpus luteum in prepubertal ewes was not influenced by hypophysectomy. However, Kaltenbach et al. (102) found that hypophysectomy on day 1 after a normal or induced ovulation resulted in failure of the corpus luteum to form, while hypophysectomy on day 5 resulted in regression of the partially formed corpus luteum. In subsequent studies, Denamur et al. (103) demonstrated that hypophysectomy of hysterectomized ewes also resulted in regression of the cor¬ pus luteum. Thus, there is agreement that hypophysectomy will prevent further luteal development and/or cause at least partial regression of existing luteal tissue. Even more controversial than the effects of hypophysec¬ tomy on luteal function are the results of studies designed to determine which hypophyseal hormone(s) are responsible for maintenance of luteal function. Injections of prolactin resulted in maintenance of luteal weight in hypophysectomized, hysterectomized ewes (103,104), but this hormone was without effect when infused into hypophysectomized ewes with an intact uterus (105). Injections of LH would not maintain luteal weight in hypophysectomized, hysterectomized ewes (103). On the other hand, infusion of LH, but not prolactin or estradiol was followed by maintenance of the corpus luteum in hy¬ pophysectomized ewes with an intact uterus (105). Based on these studies, it was suggested that LH was the primary luteotropic hormone in ewes. However, data from subse¬ quent studies (106,107) suggest it is likely that prolactin also plays a role in regulating luteal function in hypophy¬ sectomized, nonhysterectomized ewes. Interpretation of the data from these studies is complicated by the fact that the preparations of hormones used for replacement therapy were only partially purified. In addition, it is very difficult to completely remove the anterior pituitary gland from ewes without leaving a few cells adhered to the sella turcica or in the pars tuberalis. Since there would be no hypothalamic control of prolactin secretion in these animals, these cells could begin to hypersecrete prolactin, resulting in normal circulating concentrations of this hormone within a rela¬ tively short time (107). In fact, Schroff et al. (107) reported that hypophysectomized ewes in which LH maintained lu¬ teal weight invariably had normal serum concentrations of prolactin. Thus, there are data that following hypophysec¬ tomy both LH and prolactin may be required for mainte¬ nance of normal luteal function in ewes. The results of a number of additional studies are relevant regarding the roles of LH and prolactin in the regulation of luteal function. Constant infusions of LH prolonged the life span and function of corpora lutea in cyclic ewes (108), LH enhanced secretion of progesterone from the ovary in situ (109) or from luteal tissue in vitro (110,111), and daily injections of antiserum to LH caused luteal regression in

cycling ewes (112). Similar data for cattle led Hansel et al. (46) to conclude that LH was the primary luteotropin in this species. *, Although there is evidence that prolactin is important for maintenance of the corpus luteum in hypophysectomized ewes, confirming data in intact animals are not available. When serum concentrations of prolactin were reduced in ewes by >95% for an entire estrous cycle by injections of 2-bromo-a-ergocryptine there was no effect on serum con¬ centration of progesterone or length of the estrous cycle (49). Similar data are available for cows (47). Infusion of prolactin into intact ewes did not extend the life span of the corpus luteum (48), nor did infusion of prolactin into the ovarian artery result in enhanced secretion of progesterone (113). Finally, P. O’Callaghan and G. Niswender (unpub¬ lished data) have been unable to demonstrate luteal receptors for prolactin in cycling ewes under a variety of conditions. However, others (114) have reported that radioiodinated human growth hormone will bind to luteal membranes to a greater extent than will radioiodinated ovine prolactin and that this binding can be inhibited by ovine prolactin. These authors concluded that luteal receptors for prolactin are pre¬ sent and that their numbers change during pregnancy. Thus, controversy persists regarding the role of prolactin in the regulation of luteal function in domestic ruminants.

Pigs Removal of the pituitary gland during the first 2 days of the estrous cycle did not affect corpora lutea of the estrous cycle in pigs (115). Based on these data it was concluded that corpora lutea in cyclic pigs were capable of normal function without gonadotropic support after the initial stim¬ ulation of ovulation. Duncan et al. (116) were first to study progesterone synthesis by porcine luteal tissue in vitro and Cook et al. (117) demonstrated that LH stimulated the syn¬ thesis of progesterone.

Horses Because of the unique structure of the equine ovary, ovu¬ lation can occur at only one site on the ovary, the ovulation fossa (60). As a result of this anatomical feature, essentially the entire structure of the corpus luteum is contained within the ovarian stroma (Fig. 6). Anatomically, the corpus lu¬ teum of the mare reaches its maximum diameter within approximately 3 days of ovulation (118), but maximum secretion of progesterone does not occur until about 9 days after ovulation (119). Based on histological studies (120,121), it appears that the secretory elements of the equine corpus luteum are de¬ rived primarily, if not exclusively, from granulosal cells. The thecal cells begin degenerating just prior to ovulation, and their degeneration is nearly complete by 24 hr after ovulation. In contrast, granulosal cells, which are approx¬ imately 10 p,m in diameter at ovulation, have enlarged to

Corpus Luteum /

FIG. 6. Ovary from a mare cut midsagittally. The dark area is a developing corpus luteum. Note the constricted portion of the corpus luteum near the center of the ovary. This rep¬ resents the tract in which the follicle grew from the periphery of the ovary to the ovulation fossa (arrow).

15 |xm by 24 hr following ovulation and undergo cytological changes characteristic of luteinization. Luteinization of the granulosal cells appears to be complete by 3 days post¬ ovulation, but they continue to hypertrophy until day 9 (average diameter, 37.5 |xm), when maximal secretory ac¬ tivity is achieved. On day 9, in addition to the large, lightstaining luteal cells, approximately 15% of the luteal cells are small cells. These small cells are eosinophilic and are thought to represent a resting stage that can be converted to the large, light-staining luteal cells. By 12 days posto¬ vulation the large luteal cells begin to decrease in diameter, and by day 16 their diameter averages 20 |xm. The reduction in their size is correlated with a decrease in circulating concentrations of progesterone. Endocrinologically, the corpus luteum of the mare ap¬ pears to be primarily dependent upon luteinizing hormone. Antisera raised against the gondotropin fraction of equine pituitary extracts will induce luteal regression (122). Like¬ wise, administration of hCG or equine pituitary extract can extend the life span of the corpus luteum in mares (118). The concentration of receptors for LH in the corpus luteum of the mare parallels the circulating concentrations of pro¬ gesterone (123). Interestingly, the affinity of these receptors for LH also appears to increase when the secretion of pro¬ gesterone is maximal. This phenomenon appears to be unique to the equine corpus luteum.

497

in secretion of progesterone concomitant with the onset of the LH surge. This increase is important for the display of sexual receptivity by the bitch (124). Luteal growth contin¬ ues for 10 to 20 days after ovulation, at which time secretion of progesterone is maximal. This is followed by decreasing luteal activity, with circulating concentrations of proges¬ terone returning to basal levels by 55 to 90 days after ovulation in the pseudopregnant bitch. Concentrations of progesterone in the pregnant bitch appear to be similar to (126-128) or greater than those in the pseudopregnant bitch (129,130). Cessation of progesterone secretion is much more synchronous at the end of pregnancy, with a decrease to less than 1 ng/ml of blood occurring about 63 to 65 days after the LH surge. Luteal function in the bitch requires the presence of pi¬ tuitary hormones throughout pregnancy or pseudopregnancy since secretion of progesterone ceases following hypophysectomy (131). Two hypophyseal hormones, LH and pro¬ lactin, appear to be necessary for maintenance of the corpus luteum. Apparently, both hormones must be present con¬ tinuously for normal luteal function. Administration of an antiserum to LH results in a dramatic decline in secretion of progesterone (132). Likewise, administration of ergocryptine to reduce circulating levels of prolactin also results in a drastic reduction in circulating concentrations of pro¬ gesterone. Receptors for both LH and prolactin have been quantified in canine corpora lutea throughout the estrous cycle (Fer¬ nandes et al., unpublished observations). The concentration of receptors for LH remains unchanged throughout the cycle; however, concentrations of receptors for prolactin are high through day 40 but appear to decline at days 50 and 60. The physiological significance of these observations remains to be established.

Primates Continued low levels of LH are necessary for normal luteal function in hypophysectomized women (133). Treat¬ ment of normally cycling women during the luteal phase of the estrous cycle with LH or hCG increased the circulating levels of progestin and extended the life of the corpus luteum (134). LH also enhances the secretion of progesterone from human (135) or monkey (136) luteal tissue in vitro. Treat¬ ment of monkeys with antiserum to LH during the luteal phase of the menstrual cycle resulted in menstruation 2 to 4 days later (137). Thus, it appears that in primates LH is the hormone responsible for maintenance of the life span and function of the corpus luteum.

Dogs Regulation of Progesterone Secretion In the bitch, ovulation does not occur until 24 to 72 hr after the preovulatory LH surge. Luteinization of ovarian follicles (and secretion of progesterone) begins prior to ovu¬ lation (124,125), possibly due to the extended interval be¬ tween the LH surge and ovulation. There is a slight increase

The mechanisms involved in the synthesis and secretion of progesterone are complex, although this hormone is the first biologically active compound produced in the steroid biosynthetic pathway (Fig. 7). Cholesterol bound to low-

498

/ Chapter 13

R-COOH I fatty acid I

+ CHOLESTEROL

38 -hydroxysteroid dehydrogenase

A5, A4 - isomerase

FIG. 7. Biosynthetic pathway for synthesis of progesterone from cholesterol ester. The conversion of cholesterol ester to cholesterol is a reversible reaction. Cholesterol side-chain cleavage complex within the mitochondrion converts choles¬

density lipoprotein (LDL) or, in some species, high-density lipoprotein (HDL) produced by the liver is the primary sub¬ strate for progesterone synthesis. However, under some con¬ ditions, luteal cells also synthesize cholesterol from acetate for use in the steroidogenic pathway. The steroidogenic luteal cell contains LDL (or HDL) receptors that are in¬ volved in the transport of lipoprotein from outside to inside the cell. The LDL-LDL receptor complex is internalized by endocytosis. The endocytotic vesicles combine with lysosomes and the cholesterol is liberated. The free cholesterol leaves the lysosome and is either esterified and stored as lipid droplets or is used for steroid biosynthesis or in the membrane constitutents of the cell. For biosynthesis of pro¬ gesterone, cholesterol is transported to the mitochondria, where it is converted to pregnenolone by side-chain cleavage. The pregnenolone is converted to progesterone by 33hydroxysteroid dehydrogenase/A5,A4-isomerase in the smooth endoplasmic reticulum, and the progesterone is secreted. The luteal cell has a limited capacity to store progesterone. The single most important endocrine factor involved in regulating the synthesis and secretion of progesterone in the corpus luteum, irrespective of species, is LH. LH increases the synthesis and secretion of progesterone in vivo (138,139) or when incubated with luteal slices or cells in vitro (110,111,117,140,141).

Receptors for LH The presence of specific receptors for LH in the ovary and testis was first demonstrated by the ability of these tissues to preferentially bind and concentrate radioactively labeled LH or hCG in vivo (34). Subsequent studies designed to determine the subcellular distribution of binding sites, the kinetics of hormone-receptor binding, and the physi-

terol to pregnenolone, which is converted to progesterone in the smooth endoplasmic reticulum by 33-hydroxysteroid dehydrogenase/A5,A4-isomerase.

ochemical properties of the receptor molecule have em¬ ployed tissue slices (142), dispersed cells (143-145), ho¬ mogenates or particulate fractions (146-149), and isolated membranes (150-152) prepared from target tissues. To date it has not been possible to isolate and purify receptors for LH in quantities sufficient for a complete chemical char¬ acterization; however, several lines of evidence suggest that the receptor molecule is a glycolipoprotein (149,150,153). Since it has not been feasible to label receptors for protein hormones directly, these molecules have been labeled in¬ directly using LH or hCG coupled to various markers, such as ferritin (154), fluorescein (155), or radioiodine (156). It has been demonstrated repeatedly that hCG and LH compete for the same specific receptor, and hCG has been labeled most frequently. However, it has recently become clear that the steroidogenic response, the time required for internali¬ zation of the hormone-receptor complex, and the lateral mobility in the membrane of the occupied LH receptor are different for LH and hCG (61). Results from studies that have employed LH covalently linked to ferritin (154) or agarose beads (157), as well as data from autoradiographic (27,158—160), immunocytochemical (160,161), and immunofluorescent studies (155,162) and cell fractionation (151,152,163) have clearly demon¬ strated that receptors for LH are localized in the plasma membrane of target cells. In rat luteal cells the majority of hCG binding sites are localized along regions of the cell surface facing capillaries, which is characterized by mi¬ crovillus folds, whereas the basolateral surfaces of the luteal cells are characterized by junctional complexes and contain very few binding sites (27). Thus, not only are receptors for LH localized in the plasma membrane, they also appear to be concentrated in specific regions of the membrane and not distributed uniformly over the entire cell surface. Once the corpus luteum begins to develop, the secretion

Corpus Luteum / of progesterone by this gland in women (164), rats (165), and cows (166) appears to be highly correlated with the number of receptors for LH. One of the most complete studies to date regarding the relationship between the total number of receptors, the number occupied by endogenous hormone, and secretion of progesterone by the corpus lu¬ teum is that of Diekman et al. (167). The total number of receptors for LH increased 40-fold between days 2 and 14 of the cycle in ewes (Fig. 8). There was a 6-fold increase in both the number of receptors occupied by endogenous hormone and the weight of the corpus luteum and a 10-fold increase in serum concentrations of progesterone during this same period. However, less than 0.5% of the total number of receptors was occupied by endogenous hormone. By day 16 (late luteal phase), both the total number of receptors and the number occupied by endogenous LH had decreased by 75%. During early pregnancy, the numbers of total and occupied receptors were very similar to those observed dur¬ ing the midluteal phase of the cycle (167). There was a high

2

4

6

10

12

14

16

DAY OF CYCLE

FIG. 8. Serum levels of LH (top panel) and progesterone (middle panel), weight of corpora lutea (middle panel), and number of occupied and unoccupied receptors for LH in ovine corpora lutea (bottom panel) collected throughout the estrous cycle from ewes (n = 6). (From ref. 167.)

499

degree of correlation between the total number of LH re¬ ceptors, the number occupied by endogenous LH, and serum concentrations of progesterone. However, the biological significance of this finding has recently been questioned. There is little correlation between episodic peaks of LH in serum and systemic levels of progesterone during the mid¬ luteal phase of the estrous cycle in ewes (168). In addition, when serum levels of LH were increased approximately 1,000-fold during the midluteal phase of the estrous cycle, the increase in serum levels of progesterone was less than 2-fold and lasted less than 6 hr (169). These findings can apparently be explained by the recent observation that in ewes >80% of the progesterone secreted by the corpus lu¬ teum is derived from the large steroidogenic luteal cells, which have few, if any, LH receptors (61). However, it should not be inferred from these data that LH is not im¬ portant for normal secretion of progesterone. Injections of hCG or LH will increase the numbers of large luteal cells; concomitantly the numbers of small luteal cells will decrease (61). Thus, LH may regulate the differentiation of small to large luteal cells, a suggestion originally presented by Don¬ aldson and Hansel (32) for cattle. Therefore, although the secretion of progesterone by large luteal cells is not regulated directly by LH, the number of large luteal cells may depend, at least in part, on this hormone. The observation that < 1 % of the receptors for LH in the ovary (167) are occupied under conditions of maximal ste¬ roid secretion has led to the concept of “spare” receptors for this tropic hormone. However, it seems unlikely that these receptors are really spare, since circulating levels of LH are so low (0.5-1.5 ng/ml) that the large number of receptors is required to insure that sufficient numbers are occupied by endogenous hormone to stimulate steroidogen¬ esis. In addition, intracellular levels of cAMP continue to increase with increased occupancy of LH receptors, even after steroidogenesis is maximal (170,171). This observa¬ tion suggests that the receptors are coupled to adenylate cyclase and biologically functional. It may well be that cAMP is important for aspects of luteal cell function not acutely related to secretion of progesterone. For example, cAMP seems to be necessary for luteinization of granulosal cells (43) and maintenance of the characteristic morphology of luteal cells grown in culture (172). It is also possible that cAMP may be the intracellular agent involved in differen¬ tiation of small luteal cells into large ones if such differ¬ entiation can be unequivocally demonstrated (61). A variety of factors may influence the concentration of LH receptors in the ovary (35), but the primary factor ap¬ pears to be LH itself. Exposure of luteal tissue to high concentrations of LH or hCG invariably results in a dramatic loss (up to 90% in some cases) of LH receptors known as “down-regulation” (144,156,169,173,174). The loss in¬ duced by homologous hormone is time and dose dependent, and it is accompanied by a concomitant loss in hCG-stimulated adenylate cyclase activity and/or steroid production (174,175). However, at present sufficient information is not

500

/ Chapter

13

available for a meaningful evaluation of the physiological significance of down-regulation of LH receptors. Although it is tempting to conclude that the loss of adenylate cyclase activity and tissue responsiveness is related to receptor loss, there may also be a secondary lesion involved, since the administration of dibutyryl cAMP does not stimulate ste¬ roidogenesis in desensitized tissues (173). Two mechanisms have been proposed to explain the loss of LH receptors following administration of a large dose of LH or hCG. Receptors for LH could be inactivated or se¬ questered within the plasma membrane and thus become inaccessible for binding (145,174) or the hormone receptor complex may be internalized (160). The latter possibility seems most likely since there is good evidence that the LHreceptor complex enters the cell via small endocytotic ves¬ icles (27,145,160,162) and the hormone is subsequently degraded by lysosomal enzymes (176-179). That internal¬ ization of the hormone is a degradatory process rather than a mechanism for action by the hormone is suggested by the fact that chloroquine, a lysosomal enzyme inhibitor, blocks degradation of the hormone by the target cell but does not reduce steroid secretion (177). In addition, ovine luteal cells exposed to a 15-min pulse of LH secrete enhanced quantities of progesterone for 3 to 4 hr, whereas exposure to a 15min pulse of hCG results in enhanced secretion of proges¬ terone for more than 6 hr (180). Since the hCG-LH receptor complex is internalized approximately 50 times more slowly than the oLH-LH receptor complex, these data suggest that internalization of the hormone-receptor complex is the mechanism whereby the cell deactivates itself after stimu¬ lation (61). Our current working hypothesis is that the functional life of the receptor for LH in ovine luteal cells is a single binding of hormone followed by internalization and degradation of the hormone. The receptor appears to be recycled to the plasma membrane (181), probably in a manner similar to that proposed for other peptide hormones (182). This sug¬ gests that regulation of the biological effects of LH on luteal cells is a very precise phenomenon and may also explain how the hormone-stimulated cell returns to basal activity. Data obtained in vivo by Suter et al. (1969) provide ad¬

ditional insight into the mechanisms involved in loss and renewal of receptors for LH in ewes administered a phar¬ macological dose of LH during the midluteal phase of the estrous cycle. There was a dramatic increase in receptors occupied by LH within 10 min after the injection, after which the number of receptors occupied by LH decreased rapidly and returned to basal levels within 6 hr (Table 1). In contrast, the number of unoccupied receptors had de¬ creased dramatically within 12 hr but had returned to prein¬ jection levels within 48 hr. There were three interesting observations from this study. First, the total number of receptors (occupied plus unoccupied) for LH in the crude membrane fraction increased significantly within 10 min of the LH injection (“up-regulation”). Second, although the total number of receptors for LH was decreased within 12 to 24 hr after the injection of LH, at no time during the §tudy did serum concentrations of progesterone fall below preinjection levels. Thus, the biological significance of “downregulation” of ovine luteal receptors is unclear. Finally, the number of receptors occupied at 10 min after injection of LH was almost perfectly correlated with the number of receptors lost by 24 hr. This finding certainly suggests that loss of receptors for LH in ovine luteal cells is a function of occupancy by hormone. In rats, PGF2a also appears to be involved in regulating the number of luteal receptors for LH (183). The number of receptors for LH in rats administered a luteolytic dose of PGF2a decreases dramatically. However, PGF2ct does not appear to have the same effect in ewes (184).

Effects of LH on Steroidogenesis

There has been general agreement that the receptor for LH is linked biochemically to the enzyme adenylate cyclase and that binding of LH to this receptor results in activation of adenylate cyclase in corpora lutea (185). The hormonal regulation of adenylate cyclase has recently been reviewed in detail by Bimbaumer et al. (186). The critical steps in the activation of intracellular mechanisms via this process are outlined in Fig. 9. Most cell types have receptors for

TABLE 1. Number of receptors for LH and content of progesterone in the corpus luteum (CL) after injection of 7 mg LH Time after LH (hr) 0 0.17 2 6 12 24 48 72

Unoccupied receptors (mol/CL x 1012) 2.63 2.48 1.45 1.54 0.90 0.95 3.32 3.29

± ± ± ± ± ± ± ±

0.48 0.48 0.48 0.48 0.48a 0.48a 0.54 0.48

Occupied receptors (mol/CL x 1012) 0.41 5.43 1.51 0.52 0.98 0.21 0.41 0.46

Each value represents the least-squares mean ± SEM of 7 determinations. aValues significantly different from control (t = 0, p < 0.05).

± ± ± ± ± ± ± ±

0.66 0.73a 0.73 0.66 0.66 0.72 0.73 0.73

Total receptors (mol/CL x 1012) 3.04 7.91 2.96 2.06 1.88 1.16 3.73 3.75

+ 0.88 + 0.99a

+ + •++ -+±

0.99 0.88 0.88 0 98a 0.99 0.99

Corpus Luteum /

CAMP +

CO

£

Ui
ro a> jE O CO Lr

s.® 13 CO

5 CtjC Sf - ** Ss©-2

^8H| se-s5

O CO ° o

0

-

%

O CJ> CO-n

c o co £ © o > -2 p ra co

3

ex S © o a; © 2 o =6 c CO -C

X (D

▼-

c a

c

©

Asp

0

01

Leu

c

a s' ®

Q-

QC

Gin y-

CM

C «0

C CO

i

I

i

X

t-

Leu

Asp

0

Leu

0

Leu

Asp

c c 0 0 0 0l

Val

©

Val

© _J

CM

S' CC« Xi XI

CL

05 •— c 10 ca

,? 0

S ® x - £ x °>