Sertoli Cell Biology

SERTOLI CELL BIOLOGY

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SERTOLI CELL BIOLOGY

Michael K. Skinner Michael D. Griswold Center for Reproductive Biology School of Molecular Biosciences Washington State University Pullman, WA 99164-4660

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Elsevier Academic Press 525 B Street, Suite 1900, San Diego, California 92101-4495, USA 84 Theobald's Road, London WC1X 8RR, UK This book is printed on acid-free paper. ∞ Copyright © 2005, Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone: (+44) 1865 843830, fax: (+44) 1865 853333, e-mail: [email protected]. You may also complete your request on-line via the Elsevier homepage (http://elsevier.com), by selecting “Customer Support” and then “Obtaining Permissions.” Library of Congress Cataloging-in-Publication Data: Application Submitted

British Library Cataloguing in Publication Data A catalogue record for this book is available from the British Library ISBN: 0-12-647751-5 For all information on all Academic Press publications visit our Web site at www.academicpress.com Printed in the United States of America 04 05 06 07 08 9 8 7 6 5 4 3 2 1

Dedication

This book is dedicated to the memory of Professor Lonnie Russell of Southern Illinois University. Lonnie died in a drowning accident in Brazil in 2001 at the height of a successful research career in reproductive physiology. Lonnie used his expertise in anatomy to help define the intricacies of spermatogenesis with a focus on the function of the Sertoli cells. In 1993 Lonnie Russell and Michael Griswold edited a book titled The Sertoli Cell, which has served as a major resource and inspiration for investigators interested in male reproduction. It is appropriate that now—more than 10 years later—the advances in this field are summarized in this new text. It is also appropriate that this volume is a tribute to the many scientific contributions made by Lonnie Russell. It can only be hoped that this volume generates a small part of the energy and enthusiasm for science that Lonnie was able to stimulate with his wit and enquiring mind. Michael K. Skinner Michael D. Griswold

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Contents

Contributors Preface

xi xv P A R T

I INTRODUCTION CHAPTER 1

History of the Sertoli Cell Discovery

3

Rex A. Hess and Luiz R. França CHAPTER 2

Perspective on the Function of Sertoli Cells

15

Michael D. Griswold CHAPTER 3

Structure of the Sertoli Cell

19

Rex A. Hess and Luiz R. França P A R T

II SERTOLI CELL DEVELOPMENT CHAPTER 4

Embryonic Sertoli Cell Differentiation

43

Andrea S. Cupp and Michael K. Skinner CHAPTER 5

Sertoli Cell Biology in Fishes and Amphibians

71

Jerry Bouma and Joseph G. Cloud CHAPTER 6

Sertoli Cell Biology in Seasonal Breeders

81

Amiya P. Sinha Hikim and Andrzej Bartke

vii

viii

Contents P A R T

III SERTOLI CELL FUNCTION AND GENE EXPRESSION CHAPTER 7

Sertoli Cell Gene Expression and Protein Secretion

95

Michael D. Griswold and Derek McLean

CHAPTER 8

Sertoli Cell Secreted Regulatory Factors

107

Michael K. Skinner

CHAPTER 9

Proteases and Protease Inhibitors

121

Martin Charron and William W. Wright

P A R T

IV SERTOLI CELL ENDOCRINOLOGY AND SIGNAL TRANSDUCTION CHAPTER 10

FSH Regulation at the Molecular and Cellular Levels: Mechanisms of Action and Functional Effects

155

Ilpo Huhtaniemi and Jorma Toppari

CHAPTER 11

In Vivo FSH Actions

171

Charles M. Allan and David J. Handelsman

CHAPTER 12

Sertoli Cell Endocrinology and Signal Transduction: Androgen Regulation

199

Richard M. Sharpe

CHAPTER 13

Thyroid Hormone Regulation of Sertoli Cell Development

217

Paul S. Cooke, Denise R. Holsberger, and Luiz R. França

CHAPTER 14

The Transforming Growth Factor β Superfamily in Sertoli Cell Biology Kate L. Loveland and David M. Robertson

227

Contents

ix

P A R T

V SERTOLI CELL TRANSCRIPTIONAL REGULATION CHAPTER 15

Transcription Factors in Sertoli Cells

251

Jaideep Chaudhary and Michael K. Skinner

CHAPTER 16

Structure and Regulation of the FSH Receptor Gene

281

Leslie L. Heckert

P A R T

VI CELL–CELL INTERACTIONS INVOLVING SERTOLI CELLS CHAPTER 17

The Role of the Sertoli Cell in Spermatogonial Stem Cell Fate

303

Martin Dym and Lixin Feng

CHAPTER 18

Sertoli Cell–Somatic Cell Interactions

317

Michael K. Skinner

CHAPTER 19

Sertoli Cell Lines

329

Kenneth P. Roberts

P A R T

VII SERTOLI CELL PATHOPHYSIOLOGY CHAPTER 20

Sertoli Cell Toxicants

345

Kim Boekelheide, Kamin J. Johnson, and John H. Richburg

CHAPTER 21

Conditions Affecting Sertoli Cells Wael A. Salameh and Ronald S. Swerdloff

383

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Contents P A R T

VIII SPERMATOGONIAL STEM CELLS CHAPTER 22

Gonocyte Development and Differentiation

417

Peter J. Donovan and Maria P. de Miguel

CHAPTER 23

Hormones and Spermatogonial Development

437

Marvin L. Meistrich, Gunapala Shetty, Olga U. Bolden-Tiller, and Karen L. Porter

CHAPTER 24

Long-Term Cultures of Mammalian Spermatogonia

449

Marie-Claude C. Hofmann and Martin Dym

CHAPTER 25

Transplantation

471

Ina Dobrinski

Index

487

Contributors

Charles M. Allan Senior Scientist Andrology Laboratory ANZAC Research Institute Sydney, NSW 2139, Australia Phone: +61-2-9767 9100 Fax: +61-2-9767 9101 E-mail: [email protected] Andrzej Bartke Geriatrics Research Department of Medicine Southern Illinois University School of Medicine P.O. Box 19628 Springfield, IL 62794 Phone: 217-545-7962 E-mail: [email protected] Kim Boekelheide Professor of Medical Sciences Department of Pathology & Laboratory Medicine Division of Biology and Medicine Brown University, Box G-E Providence, RI 02912 Phone: 401-863-1783 E-mail: [email protected] Olga U. Bolden-Tiller Department of Experimental Radiation Oncology University of Texas M. D. Anderson Cancer Center 1515 Holcombe Blvd. Houston, TX 77030 Phone: 713-792-4848 Fax: 713-794-5369 E-mail: [email protected] Jerry Bouma The Jackson Laboratory 600 Main Street Bar Harbor, ME 04609 Phone: 207-288-6344 E-mail: [email protected]

Joseph G. Cloud Professor of Zoology Department of Biological Sciences University of Idaho Moscow, ID 83844-3051 Phone: 208-885-6388 Fax: 208-885-7905 E-mail: [email protected] Martin Charron Division of Reproductive Biology Department of Biochemistry & Molecular Biology Bloomberg School of Public Health Johns Hopkins University Baltimore, MA 21205-2179 Phone: 410-955-7831 Fax: 410-955-2926 E-mail: [email protected] Jaideep Chaudhary Center for Reproductive Biology School of Molecular Biosciences Washington State University Pullman, WA 99164-4231 Phone: 509-335-1945 Fax: 509-335-2176 E-mail: [email protected] Paul S. Cooke Department of Veterinary Biosciences and Division of Nutritional Sciences University of Illinois Urbana, IL 61802 Phone: 217-333-6825 Fax: 217-244-1652 E-mail: [email protected] Andrea S. Cupp Department of Animal Science University of Nebraska Lincoln, NE 68583-0908 Phone: 402-472-6424 Fax: 402-472-6362 E-mail: [email protected]

xi

xii Ina Dobrinski Center for Animal Transgenesis and Germ Cell Research 145 Myrin Bldg., New Bolton Center School of Veterinary Medicine University of Pennsylvania 382 West Street Rd. Kennett Square, PA 19348 Phone: 610-925-6563 Fax: 610-925-8121 E-mail: [email protected] Peter J. Donovan Stem Cell Program Institute for Cell Engineering Johns Hopkins University School of Medicine Broadway Research Building 733 North Broadway Baltimore, MD 21205-2179 Phone: 443-287-5591 Fax: 443-287-5611 E-mail: [email protected] Martin Dym Department of Cell Biology Georgetown University School of Medicine Washington, DC 20007 Phone: 202-687-1157 Fax: 202-687-9864 E-mail: [email protected] Lixin Feng Department of Cell Biology Georgetown University School of Medicine Washington, DC 20057 Phone: 202-687-1194 E-mail: [email protected] Luiz R. França Laboratory of Cellular Biology Department of Morphology Institute of Biological Sciences Federal University of Minas Gerais Belo Horizonte, Brazil 31270-901 Fax: +55-31-34992780 E-mail: [email protected] Michael D. Griswold Dean of Science Professor of Molecular Biosciences Washington State University Pullman, WA 99164-4660 Phone: 509-335-6281 Fax: 509-335-9688 E-mail: [email protected]

Contributors

David J. Handelsman Professor of Reproductive Endocrinology and Andrology University of Sydney Director, ANZAC Research Institute Sydney, NSW 2139, Australia Phone: +61-2-9767 9100 Fax: +61-2-9767 9101 E-mail: [email protected] Leslie L. Heckert Department of Molecular and Integrative Physiology University of Kansas Medical Center 3901 Rainbow Blvd. Kansas City, KS 66160 Phone: 913-588-7488 Fax: 913-588-7430 E-mail: [email protected] Rex A. Hess Professor Reproductive Biology and Toxicology Department of Veterinary Biosciences University of Illinois 2001 S. Lincoln Urbana, IL 61802 Phone: 217-333-8933 Fax: 217-244-1652 E-mail: [email protected] Amiya P. Sinha Hikim Division of Endocrinology Department of Medicine Harbor–UCLA Medical Center David Geffen School of Medicine at UCLA 1000 West Carson Street Torrance, CA 90509 Phone: 310-222-1867 E-mail: [email protected] Marie-Claude C. Hofmann Department of Biology Sciences Center The University of Dayton 300 College Park Dayton, OH 45469 Office/Lab: SC-303C/347 Phone: 937-229-2894/2507 Fax: 937-229-2021 E-mail: [email protected] Denise R. Holsberger Department of Veterinary Biosciences (P.S.C.) Division of Nutritional Sciences University of Illinois Urbana, IL 61802 Phone: 217-244-5782 Fax: 217-244-1652 E-mail: [email protected]

Contributors

Ilpo Huhtaniemi Professor of Reproductive Biology Institute of Reproductive and Developmental Biology (IRDB) Imperial College London Du Cane Road London, W12 0NN, U.K. Phone: +44-20-75942104 Fax: +44-20-75942184 E-mail: [email protected] Kamin J. Johnson Division of Biological Sciences CIIT Centers for Health Research Research Triangle Park, NC 27709 Phone: 919-558-1439 E-mail: [email protected] Kate L. Loveland Monash Institute of Reproduction and Development ARC Centre of Excellence in Biotechnology and Development Monash University 27-31 Wright Street Clayton, Victoria 3168 Australia Phone: +61-3-9594 7125 Fax: +61-3-9594 7111 E-mail: [email protected] Derek McLean Center for Reproductive Biology Department of Animal Sciences Washington State University Pullman, WA 99164-6353 Phone: 509-335-8759 E-mail: [email protected] Marvin L. Meistrich Department of Experimental Radiation Oncology University of Texas M. D. Anderson Cancer Center 1515 Holcombe Blvd. Houston, TX 77030 Phone: 713-792-4866 Fax: 713-794-5369 E-mail: [email protected] Maria P. de Miguel Cell Therapy Laboratory La Paz Hospital Paseo Castellanan 261 Madrid, 28045 Spain Phone: 34-91-7271940 E-mail: [email protected]

xiii

Karen L. Porter Department of Experimental Radiation Oncology University of Texas M. D. Anderson Cancer Center 1515 Holcombe Blvd. Houston, TX 77030 Phone: 713-792-4848 Fax: 713-794-5369 E-mail: [email protected] John H. Richburg Division of Pharmacology and Toxicology College of Pharmacy The University of Texas at Austin 1 University Station, A1915 Austin, TX 78712 Phone: 512-471-4736 E-mail: [email protected] Kenneth P. Roberts Associate Professor Department of Urologic Surgery University of Minnesota 420 Delaware St. SE MMC 394 Minneapolis, MN 55455 Phone: 612-625-9977 E-mail: [email protected] David M. Robertson Prince Henry’s Institute of Medical Research P.O. Box 5152 Clayton, Victoria 3168 Australia Phone: +6-3-95944386 Fax: +6-3-95946125 E-mail: [email protected] Wael A. Salameh David Geffen School of Medicine at UCLA Harbor–UCLA Medical Center Division of Endocrinology Rb-1 1124 West Carson Street Torrance, CA 90502 Phone: 310-222-1867 Fax: 310-533-0627 E-mail: [email protected] Richard M. Sharpe MRC Human Reproductive Sciences Unit Centre for Reproductive Biology University of Edinburgh Chancellor’s Building 49 Little France Crescent Old Dalkeith Road Edinburgh, EH16 4SB Phone: +44-131-242-6387 Fax: +44-131-242-6231 E-mail: [email protected]

xiv

Contributors

Gunapala Shetty Department of Experimental Radiation Oncology University of Texas M. D. Anderson Cancer Center 1515 Holcombe Blvd. Houston, TX 77030 Phone: 713-794-4858 Fax: 713-794-5369 E-mail: [email protected] Michael K. Skinner Director and Professor Center for Reproductive Biology School of Molecular Bioscience Washington State University Pullman, WA 99164-4231 Phone: 509-335-1524 Fax: 509-335-2176 E-mail: [email protected] Ronald S. Swerdloff David Geffen School of Medicine at UCLA Harbor–UCLA Medical Center Division of Endocrinology Rb-1 1124 West Carson Street Torrance, CA 90502 Phone: 310-222-1867 Fax: 310-533-0627 E-mail: [email protected]

Jorma Toppari Departments of Physiology and Pediatrics University of Turku Kiinamyllynkatu 10 20520 Turku, Finland Phone: +358-2-333-7297 Fax: +358-2-2502621 E-mail: [email protected] William W. Wright Department of Biochemistry Reproductive Biology Bloomberg School of Public Health Johns Hopkins University East Baltimore Campus W3508 Wolfe Street Building Baltimore, MD 21205-2179 Fax: 410-614-2356 E-mail: [email protected]

Preface

In 1993 Lonnie Russell and Michael Griswold edited a book titled The Sertoli Cell, which has served as a major resource for investigators interested in male reproduction. It is appropriate that now—more than 10 years later—the advances in this field are summarized in this new text. It is also appropriate that this volume is a tribute to the many scientific contributions made by Professor Lonnie Russell of Southern Illinois University. Lonnie died in a drowning accident in Brazil in 2001 at the height of a successful research career in reproductive physiology. Lonnie used his expertise in anatomy to help define the intricacies of spermatogenesis with a focus on the function of Sertoli cells. He was first an anatomist and physiologist, and he used those tools to produce an enormous amount of information on the testis. His laboratory published the first and only visualization of intact Sertoli cells reconstructed from serial sections in the electron microscope. These papers allowed those of us in the field a fundamental visualization of the beauty and complexity of the Sertoli cells. It is essential for the molecular biologists and biochemists to place their findings in the context of the complex biology of the testis. Part of the goal of this current text is to make that process easier. The editors of this text (the Mikes) have a combined nearly 50 years of experience focused on exposing the secrets of the Sertoli cells. Both of us have spent many years reporting on specific gene products of Sertoli cells and over the combined 50 years we have thoroughly investigated perhaps a dozen gene products. The biggest change in the field has been the use of expression arrays and gene knockout technologies that allow the monitoring of thousands of expressed sequences and the more laborious testing of their physiological functions. In The Sertoli Cell, Russell and Griswold reported that nearly 300 papers were published in 1990 that somehow involved these cells. Since the millennium there have been nearly twice that

number of papers per year, suggesting these new technologies and other factors have stimulated an increased interest in the Sertoli cell. The current text attempts to present a systems biology approach to an understanding of the Sertoli cell. A combination of molecular, cellular, and physiological aspects of Sertoli cells are presented. Due to the essential role Sertoli cells play in the process of spermatogenesis, topics such as spermatogonial stem cells and germ cell transplantation are also presented. An attempt was made to identify areas that have had significant advances over the past decade, as well as suggest important areas for the future analysis of Sertoli cell biology. Information is presented to provide the novice with basic information on the topics as well as to provide experts with new details that have advanced the field. We hope you find Sertoli Cell Biology a useful reference and that it provides insight for an understanding of the Sertoli cell and male reproduction.

ACKNOWLEDGMENTS The chapters provided and hard work of the contributing authors is what made this book possible. The editors thank Ms. Mica Haley, Ms. Jacqueline Garrett and the staff at Elsevier/Academic Press for their assistance and patience. We thank Dr. Rex Hess and acknowledge his interest in the book, particularly in the design of the cover. The cover includes an electron micrograph of a Sertoli cell from Dr. Lonnie Russell and his three-dimensional reconstruction of the cell, as well as a light micrograph of the Sertoli cell provided by Dr. Hess. This book would not have been possible without the editorial and administrative support of Ms. Jill Griffin. Jill’s dedication and efficiency were critical for the book and indispensable to the editors.

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P A R T

I INTRODUCTION

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C H A P T E R

1 History of the Sertoli Cell Discovery

I. II. III. IV. V. VI. VII. VIII. IX.

REX A. HESS

LUIZ R. FRANÇA

Reproductive Biology & Toxicology, Veterinary Biosciences, University of Illinois, Urbana, Illinois

Laboratory of Cellular Biology, Department of Morphology, Institute of Biological Sciences, Federal University of Minas Gerais, Belo Horizonte, Brazil

Brian P. Setchell [10,11], whose foreword in the first book provided a sincere and deserved admirable look at Enrico Sertoli [12].

INTRODUCTION THE STUDENT THE MICROSCOPE THE LABORATORY THE DISCOVERY COMPLETING THE MANUSCRIPT AFTER GRADUATION TRANSITION TO THE MODERN SERTOLI CELL POSTSCRIPT References

II. THE STUDENT Enrico Sertoli was 18 years old when he began his studies in medicine and research at the University of Pavia in Northern Italy in 1860 [3]. He studied general medical subjects at first and then after 2 years began his research studies in the laboratory of the distinguished physiologist and histologist, Professor Eusebio Oehl (1827–1903). It is interesting that Camillo Golgi, for whom the Golgi apparatus is named, was a fellow student at the university and he and Sertoli both studied under Professor Oehl [13] and graduated in the same year, 1865. Sertoli was born on June 6, 1842, to a noble family in the small town of Sondrio, located North of Milano along the Italian–Swiss border [3]. His noble birth in all probability meant that he was expected to attend university and study medicine. Unfortunately, as Sertoli entered his teenage years, the countryside was not at peace and there was talk of war between Prussia and Austria during this period. Accordingly, Sertoli may have had the same urges of young people today to join the local forces and defend his country, but his father surely urged him to complete his medical training before entering the army, which Sertoli did when war broke out after he graduated in 1865 [12].

I. INTRODUCTION The first edition of The Sertoli Cell was an appropriate vision of the late Professor Lonnie D. Russell, because he studied the Sertoli cell in more depth than most other modern-day scientists. He published more than 200 papers, of which nearly half were focused on the Sertoli cell, including the first book devoted to the cell, which he coedited with Michael D. Griswold [1]. Therefore, this chapter is written in honor of Lonnie because he was a fun-loving friend and visionary scientist who always used the microscope and his imagination to find new insights into complex scientific problems of the testis, and in particular the Sertoli cell. Lonnie’s devotion to this cell was exemplified by the license plate that he attached to his automobile, which read “Sertol 1,” and by his cat whose name was also “Sertoli.” Factual events surrounding Sertoli’s life were gathered from reading numerous reviews [2–9], particularly those of the distinguished scholar SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

3

Copyright 2005, Elsevier Science (USA). All rights reserved.

4

Rex A. Hess and Luiz R. França

The University of Pavia was an old, well-established center of higher education located just south of Milano, which according to historical records was established by edict of the Emperor Lotarius in the year 825. The culture surrounding this famous university was surely one that encouraged the highest standards of achievement and bred intellectual inquiry that was capable of producing Nobel Prize–winning scientists, such as Golgi, who also studied under Bizzozero and was later nominated for the first Nobel Prize in Physiology and Medicine in 1901 and then every year until 1906, when he shared the prize with Santiago Ramón y Cajal.

III. THE MICROSCOPE The cellule ramificate or branched cell was discovered using the personal microscope of Enrico Sertoli (Fig. 1.1). He had purchased the microscope in 1862, after he began his research studies under Professor Oehl. The Belthle microscope that he purchased from the Kellner Optical Institute in Wetzlar, Germany, was a state-of-the-art light microscope at that time. Instruments produced by the Kellner Optical Institute were compound microscopes with three or more lenses, and each came with 10× and 20 × magnifier eyepieces. They also had a screw system for lowering the compound lenses to the slide, so that there was less breakage of the glass coverslips.

FIGURE 1.1 The Belthle microscope that Sertoli personally purchased in 1862 and the same microscope that he used to make the famous discovery of the cell that now carries his name. (Photograph kindly provided by Michi Sertoli, the great nephew of Professor Enrico Sertoli, of Milan, Italy.)

In contrast, some of the more common microscopes found in laboratories in 1862, such as the van Deyl and Beck microscopes, were no more than high-powered magnifying lenses. Such microscopes would have posed difficulties for a serious student and may have been reason enough for Sertoli to purchase one of the highest quality microscopes available, knowing that the instrument chosen would become the limiting factor in his histological research accomplishments. We do not know the complete story behind the purchase of this microscope, but it was surely a momentous occasion. It is possible that microscope purchases were required for every medical student on entry into the histology course. Regardless, we can envision that some students were capable of purchasing a compound microscope, whereas others found it necessary to settle for the less expensive brands that provided only a magnifying lens. The quality of the Belthle microscope and its personal importance are evidenced by the care that Sertoli devoted to it, which has permitted its survival for more than 100 years, along with a wooden storage box and the original two-page letter of guarantee that was provided by Belthle from the Kellner Optical Institute (Fig. 1.2).

IV. THE LABORATORY To our knowledge, there are no photographs of Sertoli in his laboratory; therefore, we must assume that the room was similar to others that were photographed in the late 1800s. Professor Oehl’s laboratory was likely a typical large room with a high ceiling and large windows. Because the Edison lamp did not arrive until 1890, few professors, except in the department of physics, would have had electric lights in their offices and laboratories. The furniture would likely have been made from a hardwood and coated with dark stain. Tall wood encasings would have lined the outer walls. A bench would have run the full length of the room with hardwood shelving down the center. It would have encased chemical solutions, flasks, stopcock bottles, and other supplies common to the research lab. Natural gas was used quite extensively in Europe by the late 1860s; however, harnessing the gas was not easy, and Robert Bunsen had not as yet invented the famous “Bunsen burner.” This marvelous invention in 1885 permitted the controlled burning of natural gas through a metal tubule by regulating the amount of gas and air proportions individually, which essentially allowed for increases in the temperature and in the intensity of the flame. Until this device came along, everyone

Chapter 1 History of the Sertoli Cell Discovery

5

FIGURE 1.2 (A) This wooden box is the original storage box for the microscope, which Sertoli maintained so meticulously. (B) The original two-page document that was sent along with the microscope from the German company Belthle in Wetzlar. The paper is a guarantee from the Optics Institute of von C. Kellner. (The date 1862 has been magnified digitally for emphasis. Image kindly provided by Michi Sertoli, the great nephew of Professor Enrico Sertoli, of Milan, Italy.)

simply used a straight tubule with a round base and controlled the flow of gas by a single valve. This led to many accidents and explosions, but the flame was still superior to candle or alcohol lamps. It was common to have a large blackboard and chalk in the laboratory. We can imagine the blackboard being covered with drawings of Sertoli’s observations and maybe even outlines of experiments dealing with Professor Oehl’s own research. On the bench may have been another typical microscope that was often used in physiology, the Cuff simple microscope, which was made by Dollond of London. This monocular-type scope was used for dissections. It was capable of magnifying with fairly good resolution up to 10× to 20×. The instrument had a tube body mounted above a stage, similar to the compound microscope, but it was strictly a magnifying lens on a stand. Thus, this crude magnifier was the precursor of the dissecting microscope. Along that same bench would have been glass jars with specimens preserved in alcohol and acids and other types of solutions that were used in histology. On the opposite side could have been stacks of histology glass slides and possibly an open copy of the first textbook of histology, written by the Swiss scientist, Albert Kölliker [14]. Like many scientists during that period, Sertoli would have worn the typical white smock-like lab coat with five buttons up the front. The coat would protect his white shirt and dark tie, the formal dress of a university student.

V. THE DISCOVERY In anticipation of the microscope’s arrival, Sertoli likely collected several pieces of human testes preserved in a sublimate solution (a precipitating solution formed by adding ammonia to mercuric chloride) that he later reported as the incubation solution of choice at that time [15]. Without being aware, Sertoli was using a very nice model to investigate the seminiferous epithelium, because in humans, unlike in mice and rats, the Sertoli cell occupies about 37% of the epithelium. In contrast, the Sertoli cell occupies about 15–20% in the rodent species. The human testis has a higher ratio of Sertoli to germ cells, due to the reduced efficiency of spermatogenesis. It appears that Sertoli worked only with human testes throughout his career. Thus, working in a medical school was an advantage to Sertoli from the very beginning. Sertoli used several different types of preparations of testes, including microdissections of individual seminiferous tubules, thin sections of the testis after sublimate incubation, pieces of fresh tissue, and frayed sections of tubules. Like all young students in the laboratory, he probably worried that the tissue may have remained in the solution too long. With such concerns, perhaps he had numerous conversations with Golgi and other students regarding the latest methodologies that were being tested in histology to better preserve structures and improve the visualization of cells.

6

Rex A. Hess and Luiz R. França

After the new microscope arrived, Sertoli probably spent the first few weeks working obsessively with the shiny new instrument. He no doubt spent endless hours getting the angle of the sunlight just right so that the cells would appear more clearly. To a scientist today, the methods used by Sertoli were crude and harsh. It is astounding that such important discoveries were made during this early period of development of what we now accept as routine histological procedures. These early observations were made without the benefit of fixatives that could crosslink proteins and bind lipids in the tissue. Alcohols and acids were the primary methods for treatment of fresh samples. Although formaldehyde was discovered in 1859, it was not until after the commercial synthesis of formalin from methanol by Hoffman in 1868 that sufficient quantities became available for testing in various medical and chemical procedures. The actual use of formalin for hardening of tissue did not come about until 1892 when a chemist, Blum, was asked to test formalin as a potential antiseptic agent. Formalin hardened the skin on his fingertips, and the rest is now history for this widely used fixative. At first, Sertoli may have called the cell a tree-like cell or stringy cell or some other description that suggested that this cell had long extensions. On the first page of his publication [15], the term mother cells is used, which suggests that his observations were quite perceptive and even intuitive of the true Sertoli cell function. In 1863, it was necessary to draw observations made through a microscope. Although a photo was actually produced in 1827 on materials hardened after exposure to light for up to 8 hours [16], the word photography was not invented until 1839, by Sir John Herschel. This early type of photography was time consuming and quite expensive; therefore, it was not routine in the scientific laboratory, and a box for holding the film that could be used on a microscope was not invented until 1884, when George Eastman introduced flexible film and the boxed camera [16]. Therefore, Sertoli would have spent many hours drawing what he observed. Sertoli’s first drawings must have been simple (probably similar to those in Figure I of the original plate, reproduced here as Fig. 1.3). Looking through a microscope and drawing what you see is difficult even when the tissue is well preserved and well stained. Unfortunately, the corrosive solutions that Sertoli used extracted the cells and left them rather transparent. Nevertheless, he would have been very excited, as every student would be upon using for the first time a new instrument or observing for the first time a new organelle or cell. In truth, the discoveries by Sertoli were things that had never before been described by others.

FIGURE 1.3 Drawings taken from the original paper (Fig. I a–d) of Sertoli [15].

Reproductive biology literature was scarce in the 1800s. However, Sertoli was reading the work of Kölliker [17] and referred to his work as being the “most authoritative one.” But Kölliker claimed that these cells of interest to Sertoli were polygonal in shape, instead of conical or cylindrical in shape as described by Sertoli, and such statements in the early scientific literature tended to become dogma. To discover something that contradicts the published literature surely provides the ultimate excitement for a scientist, and such exhilaration may have been even greater during that period in time if we take into consideration the culture of science in Europe in 1863. Such discoveries can also be intimidating, especially when one considers that their observation may contradict a famous scientist, such as Kölliker. One cannot help but wonder what Sertoli wrote in his notebook. Today we publish only small portions of the total data that are collected during an experiment. However, in the 1800s, every observation was novel and worthy of discussion. He probably had numerous drawings lying on the bench, and his notebooks were likely filled with drawings and intricate descriptions. We know he described the cell as having branches and blobs at the ends. Maybe the branches surrounded germ cells and maybe the branches were like those of a tree that drops its fruit as the harvest ripens. Anything was possible, because it was new. It was possible that this cell, like the great maestro of a symphony, directed the production of sperm in the testis. Maybe spermatozoa developed directly from the branched cell. Sertoli and the other histology students on the campus, such as Golgi, may have discussed the work of Professor Waldeyer in Berlin. Professor Waldeyer had extracted a substance from logwood (Haematoxylon campechianum) that someone had collected from Central America. Apparently, the substance, now called hematoxylin, was a nonquinonoid that was soluble in water and, when readily oxidized, would stain leather. Waldeyer was the first to propose this solution for staining histological tissues, too. However, it was not

Chapter 1 History of the Sertoli Cell Discovery

until Ehrlich formulated the extract with alum that the method worked efficiently as a counter stain to eosin [18]. Thus, Sertoli had little to work with in 1863 but he must have tried whatever solutions he could find in the laboratory or borrow from other labs. He mentioned in the paper that he tried clearing the tissue with nitric and acetic acids, and even tried staining with iodine. He wrote, as translated by Setchell [10], “Nitric acid shrivels and deforms the cells and turns the nuclei yellow. Tincture of iodine stains these structures yellow, but the nuclei even more intensely than the cells.” Such testing of different methods may have improved the observations, because in some drawings, Sertoli included round germ cells, “seminiferous cells,” embedded within the branched cell limbs (Fig. 1.4). Sertoli drew with intricate detail what he observed. He was the first to recognize and report lipid droplets in this cell, and he mentioned several times that lipid could exert very important functions in the cell, a function that we still know little about today. He also drew the cell as appearing syncytial or as a branched multinucleated cell, which surely raised many questions from students such as Golgi, because such ideas were common among those who studied the brain. Sometimes he simply looked at fresh tissue (Fig. 1.5) and observed what appeared to be an epithelial lining of the branched cells surrounding germ cells. His drawings indicated that the united branched cells were protecting or extending a hand to the germ cells (Fig. 1.4). Even without the privilege of seeing Sertoli’s notebook, we can imagine that it contained numerous pages of drawings and descriptions, many of which he used as the basis for writing the following narrative, as translated by Setchell from Sertoli’s original 1865 paper [10]:

FIGURE 1.4 Drawings taken from the original paper (Figs. II a–d and III a–e) of Sertoli [15].

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FIGURE 1.5 Drawings taken from the original paper (Figs. IV and V a–b) of Sertoli [15].

IV. Finally, some special cells, which I saw in moderate number in the preparations and which, to my knowledge, have not previously been observed and described by anyone. They appear in the form of irregularly cylindrical or conical cells, with indistinct margins, provided with a nucleus, always containing a nucleolus. Their contents comprise fine droplets of fat in a substance that is reasonably transparent because it is homogeneous. These cells almost always have quite transparent extensions, in the interior of which fine droplets of fat can frequently be seen. They have an irregularly shaped body from which often protrude one or more extensions, and two extremities of which the upper is usually large and bounded by a well-marked margin that sometimes appears double (Fig. I a,c,d). Lower down the cell often is contracted somewhat, formed like a sort of collar (Fig. I a,d, Fig. III b). The other, luminal extremity, becomes narrower and forms an extension, which often ends abruptly in a rounded off tip with delicate outlines (Fig. III c,d). Often, the tip is torn and it is not possible to determine how it ends normally. I have observed that other cells bifurcate and send out secondary extensions (Fig. I b,d).

Such detailed descriptions by Sertoli are amazing, considering the crude equipment and conditions that he worked under at the time. To put his work in perspective, it is helpful to consider the time frame of events in microscopy and histology in the 19th century that is shown in Table 1.1. Thus, by the time Sertoli published his first paper in 1865, most of the accepted standard methods of histology were lacking: fixation of tissue, embedding, sectioning with microtomes, and routine histological stains. In the year that Sertoli published his last manuscript as a professor, 1886 [19], major breakthroughs in

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Rex A. Hess and Luiz R. França TABLE 1.1 Significant Scientific Discoveries Surrounding the Period of Sertoli

1839

Theodor Schwann presented his famous “cell theory.”

1839

The 3- × 1-inch glass microscope slide was established as a standard by the Microscopical Society of London.

1840

The first commercial glass coverslips were used.

1841

Kölliker reported that spermatozoa arose from cells within the testis.

1850

Leydig’s article on interstitial cells is published.

1850s Clark’s alcohol-acetic solution and Müller’s solution for fixation were revealed. 1855

Water immersion lens first displayed at the Paris Exposition.

1858

Carmine stain first used by Joseph Gerlach.

1859

Butlerov discovered formalin.

1860s Wilhelm Waldeyer first used aniline dyes, one called Paris blue. 1863

Waldeyer was first to stain histological sections with an extract that became hematoxylin.

1864

Fromman first demonstrated the use of silver for the identification of axons.

1865

Sertoli’s paper on “branched cells” of the testis is published.

1869

Theodor Klebs first used melted paraffin to support a tissue block while sectioning.

1870s First standard microscopical slides used for teaching. 1873

Ernst Abbe published his work on the theory of the microscope, which explained the difference between magnification and resolution. His formula was used to calculate resolution.

1873

Ernst Leitz microscope is introduced with a revolving mount (turret) for five objectives.

1879

Carl Zeiss Jena produced its first oil immersion objective in 1880, designed by Ernst Abbe.

1879

Walther Flemming discovered mitosis.

1880

August Kohler determined the optimum spacing for the light source and the condenser, which would produce sharper images; thus, was born Kohler illumination.

1880s First electrical illumination, rather than sunlight, is reflected off a substage mirror. 1885

The first sliding microtome was developed.

1886

The first rotary microtome was developed.

1886

The substage condenser was developed.

1886

Ernst Abbe introduces the apochromatic objective lens, bringing the red, yellow, and blue colors into one focus, and requiring numerous lens elements.

1886

Ehrlich invented a stable solution of hematoxylin with a long shelf life.

1886

Benda introduced iron-hematoxylin techniques.

1893

Blum tested formalin as a fixative, and Carl Weigert introduced it as routine preservative for tissues.

1894

Zenker’s fixative became available.

1896

“Sudan” stains for lipids introduced by Daddi.

1897

Bouin’s fixative became available.

microscopic technique and instrumentation were just coming on the scene, such as Kohler illumination and Ernst Abbe’s apochromatic lenses, which became available just before Sertoli retired. Despite these handicaps in technology, Sertoli worked with whatever was available and carried the observations to the limits of current microscopic resolution. Thus, we must conclude that Sertoli was an exceptional and determined student, with a keen skill for observation and capable of having original thoughts that others of his day were not giving consideration.

VI. COMPLETING THE MANUSCRIPT Before Sertoli completed the first draft of his manuscript, he probably had numerous discussions with his adviser, Professor Oehl. Professor Eusebio Oehl had just founded the first Institute of Physiology at the University of Pavia in 1861. His research was not focused on the testis and, at the time that he mentored Sertoli and Golgi, he was studying extracted human saliva and salivary ducts. Thus, we must assume that Sertoli’s focus on the testis may not have been top priority for his major adviser. On the other hand, in the late 1800s, every observation under the microscope was something new and it is not inconceivable to think that the adviser would recommend that each new student study a different organ. Regardless of the reason why he studied the testis, Sertoli was making major discoveries that would contradict the published literature and this must surely have made his professor either very excited or very worried. The last figure in Sertoli’s manuscript was a low magnification of a seminiferous tubule cross section showing germ cells and even spermatozoa associated with the branched cells that contained lipid droplets (Fig. 1.6). He had a great desire to observe the borders of the branched cell, and in his final experiment he tried something new [10]: “… in some sections treated with a weak solution of ammonia, I have seen that quite definite dark borders, limit the basal extremity of the branched cells ….” After these remarks, he showed restraint by pointing out the limits of his observations. He ended the paper with his conclusions from numerous observations. One of the most important of which was his conclusion that “it is not likely that these cells produce the spermatozoa …”[10], for which he gave three arguments: (1) Spermatozoa have been observed in the extension of only a very few cells. (2) The spermatozoa are found only in the extensions, where it would be possible for them to have

Chapter 1 History of the Sertoli Cell Discovery

FIGURE 1.6 Drawing taken from the original paper (Fig. VI) of Sertoli [15].

entered accidentally, and I have never seen spermatozoa inside the cells. (3) The formation of the spermatozoa would be consistent neither with the form of the branched cells, which are different from the seminiferous cells, the real progenitors of the spermatozoa, nor with their constant position inside the tubule, nor their tendency to enclose the seminiferous cells among their branches, nor finally their communication with one another through the extensions.

Yet, in the manner of an honorable scientist, he said he could not categorically deny the possibility, but that he did not think the branched cells produced spermatozoa. Nevertheless, with the insight of a keen scientist, Sertoli ended the paper with a suggestion that the “function of the branched cells is linked to the formation of spermatozoa.” This comment along with that on page 1 of the manuscript regarding “mother cells” indicates that Sertoli was indeed the first to suggest that the “branched cell” served as a “cellule madri” or “mother cell” or “sustentacular cell.” It is likely that Sertoli made significant progress in writing the manuscript during 1864. The entire process of manuscript preparation, rewrite, submission, review, resubmission, and printing sometimes takes nearly 1 year nowadays, so it is reasonable to picture the same process taking more than 1 year in the 1800s. However, the time from submission to publication may have been considerably shorter than today, because every observation in the 1800s was publishable and the number of capable reviewers for any subject was limited. Therefore, it is likely that Sertoli completed his major observations and writing in late 1863 or early 1864 and submitted the manuscript for publication prior to graduation in 1865. There has been no mention of why Sertoli was the single author on the paper, even though it is reasonable to assume that his research professor approved of the work in his laboratory. Until about 1950, it was common to see single-author papers or papers with just two authors. This may reflect the fact that much of

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the research performed during the late 1800s and early 1900s did not require collaboration and involved the use of simple tools of investigation. Research efforts back then required a tremendous personal endeavor. Furthermore, until the late 20th century, the evaluation of professorships did not depend on counting the total number of publications. The first manuscript is always very special. A young scientist will dream of the day when his or her own name appears beneath the title of a manuscript. Sertoli may have written the title several times, but the published title had a very specific focus: “Dell’ esistenza di particolari cellule ramificate nei canalicoli seminiferi del testicolo umano,” or as interpreted: “On the existence of special branched cells in the seminiferous tubules of the human testis.” He restricted the study to just one cell type, although he had observed all of the germ cells, had noted what is now called stages of the seminiferous epithelium [20], and even recognized the beginnings of a “wave” of spermatogenesis.

VII. AFTER GRADUATION Sertoli graduated in 1865 and traveled to the University of Vienna to study with Ernst Wilhelm von Brücke, a physiologist [3]. In 1866 he returned home briefly to defend his country in the war between Prussia and Austria. After the war was over, he stayed in the army for a while before returning to science. In 1867 he went to work in the laboratory of Ernst Felix Immanuel Hoppe-Seyler in Tübingen (not yet part of Germany). Throughout his career, Sertoli’s research studies focused on many different organ systems other than male reproduction. At different times, his studies included the following: the lymphatic system, lungs, coccygeal gland, nutrition, kidney, tactile hairs, and smooth muscle [12]. After a series of lectures at the Politecnico in Milan in 1870, he was given a professorship at the Advanced Royal School of Veterinary Medicine in Milan [3]. From 1871 to 1880, Sertoli was professor and chair of anatomy and physiology, and then from 1880 to 1907, he was chair of physiology. It was reported by Negrini [3] that in 1900 at the Anatomy Congress in Pavia, the works of numerous eminent Italian and foreign scientists were presented on a series of microscopes, and one microscope held the label “Cells of Sertoli.” By this date, others were also referring to the branched cells in association with Sertoli’s name [21, 22]. In the fall of 1906, a year before Sertoli retired as professor, we can easily imagine that Golgi’s name

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Rex A. Hess and Luiz R. França

was being mentioned rather frequently in the cafés surrounding the University of Milano. Golgi had been nominated every year since 1901 for the Nobel Prize in Physiology and Medicine. Although there is no documentation that Sertoli and Golgi were long-term friends or anything more than fellow students in the same laboratory, we can imagine that Sertoli may have discussed this important event with his colleagues in Milano. We have no record that Sertoli and Golgi corresponded, but it is easy to imagine that Sertoli congratulated Golgi after he was awarded the Nobel Prize in 1906. Golgi went on to become one of the most famous scientists in neurobiology and cell structure. Every student of histology will remember the name “Golgi” even if they do not always remember the name “Sertoli.” Yet Golgi surely thought about the fact that von Ebner [22] had named the branched cells the “branched cells of Sertoli,” whereas his own name was only associated with a cytoplasmic organelle. Sertoli retired the year after Golgi received the Nobel Prize. He returned to his hometown of Sondrio, due to an illness, and there he lived until his death on January 28, 1910. Sertoli never married and devoted his adult life to teaching and his research.

a primary role in preparing spermatozoa to reach and fertilize the egg [23]. Other misconceptions were prevalent during this transition period. For example, Bardeleben in 1897 thought that the branched cells were derived from Leydig cells [24] and it was not until the 1940s that this idea was replaced [25]. Also, Sertoli implied [15] and von Ebner [26] agreed that the branched cells were syncytial. La Vallette, however, insisted that they were individual [27], and it was several decades before this argument was settled. Lonnie Russell [28] pointed out, in the first edition of this book, that early electron microscopy did not detect the cell boundary, but in 1956, Don Fawcett was able to demonstrate the Sertoli cell membrane with high resolution [29]. In 1878 Sertoli published a statement that the branched cells, which he now called cellule fisse or fixed cells (Table 1.2), no longer divided in the adult testis [30]. It was not until 1963 that this observation was actually accepted [31], and a more detailed understanding of Sertoli cell proliferation and maturation did not come until the 1980s [32]. It is amazing that more than a century was necessary for new and solid information on Sertoli cell function to be published. In this regard, only at the end of the 1960s did scientists in the field show clearly the existence of the blood–testis barrier (now called the Sertoli

VIII. TRANSITION TO THE MODERN SERTOLI CELL Because it would have been difficult to characterize Sertoli and germ cells morphologically, just as it is difficult even today if the tissues are not well fixed and embedded, several decades after the discovery of the Sertoli cell, scientists were still debating the origin and function of the cell [21, 23]. The only common point was that the Sertoli cell had an important role in supporting germ cells, which was based on the strong evidence provided by Sertoli in his 1865 publication. In this regard, although the cyclical changes in the cell and spermatogenesis were being noted at the end of the 19th century, it was strongly claimed that Sertoli cells originated from germ cells or that part of the Sertoli cell population gave rise to germ cells and formed other Sertoli cells, in an amitotic process. In this way, it appears that Sertoli cells were originally defined as what we now characterize functionally as stem cells. Among other functions that they were ascribing for Sertoli cells at that time, we can cite nutrition, secretions, dehydration of germ cells, shaping of the sperm nucleus, formation of the spermatid bundles, and spermiation of germ cells. During this period, many researchers strongly suggested that the Sertoli cell had

TABLE 1.2 Significant Discoveries by Enrico Sertoli that Are Associated with Spermatogenesis Cellule ramificate (branched cells) of the seminiferous tubules, later called cellule fisse or fixed cells [30, 42]; later called branched cells of Sertoli by von Ebner [22], and finally Sertoli cells by Hanes in 1910 [43]. Cellule mobili (germ cells) that form spermatozoa; the first to note “mobility” of these germ cells within the seminiferous epithelium. Noted maturation of germ cells was not uniform in the seminiferous epithelium [2], thus hinting of stages; however, Sertoli always gave credit to von Ebner for discovery of the cycle of the seminiferous epithelium [2]. Cellule germinative or spermatogonia. Spermatozoa were derived from “nematoblasti” or spermatids. Noted “branched cells” were highly resistant to chemical digestion compared to germ cells. Recognized spermatid differentiation, tail formation, nuclear elongation, formation of head, and the cytoplasmic droplet. Three stages of spermatocyte (cellule seminiferi) development [30]. Two types of spermatogonia. A spermatogenic wave [30]. Cytoplasmic bridges between germ cells [30]. Sertoli cells do not divide in the adult testis, in contrast to germ cells [19].

Chapter 1 History of the Sertoli Cell Discovery

11

cell barrier) [33–35]. This barrier appears only after the formation of tight junctions between adjacent Sertoli cells and is responsible for the formation of the basal and luminal compartments of the seminiferous epithelium, the former being essential for the development of young spermatocytes and spermatids. Sertoli’s publication in 1865 was so convincing that scientists rather quickly gave the cell his name. This acceptance probably allowed others to continue their focus on spermatozoa and their origin in the testis, the real interest of those studying male reproduction at that time. As such, other topics became more important to establish (Table 1.2), such as the development of germ cells and their association in stages, as well as the duration of the cycle of the seminiferous epithelium [20, 36–38]. During this post-Sertoli period, endocrine aspects of testis function and diseases involving the Sertoli cell became topics of serious inquiry [10]. From 1865 until 1955, the average number of papers published with Sertoli as a key word was approximately one per year, with most of the growth starting in 1952 with the classical publication by Leblond and Clermont defining the stages of the cycle of the seminiferous epithelium [20] (Fig. 1.7). This transition period included two world wars, which of course reduced the ability to perform research, but at the same time allowed the development of significantly improved technology, for example, the transmission electron microscope, which promoted a burst in new research involving the Sertoli cell. For that reason, from 1966 to 1970 the number of papers per year more than doubled to 31 per year. It is interesting to note that the rate doubled again from 1971 to 1975, the period during which Lonnie Russell began publishing his research. From the time of his first publication in 1973, the number of Sertoli cell publications per year has grown by more than threefold. The number of Sertoli cell papers published per year has continued to increase and as of 2002 had reached more than 400 papers per year.

IX. POSTSCRIPT FIGURE 1.7 (A) Graph of the total number of publications retrieved from the National Library of Medicine via PubMed that use the term Sertoli cell in the title, abstract, or as a mesh term. (B) Graph showing the mean number of Sertoli cell papers per year for each period shown.

It was reported by Brian Setchell [12] that Sertoli was a quiet man, “… speaking little in the laboratory, sometimes passing whole days in silence.” Sertoli was a stout man [12], of “medium height, stocky, robust, and with a large head.” Such a personal description provides a striking contrast to the modern-day person we now call “Mr. Sertoli Cell.” Lonnie Russell was never quiet. He was a conversationalist and without doubt would have died much younger if he had to spend even one day without speaking a word.

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Rex A. Hess and Luiz R. França

Lonnie was neither short nor stocky, but we might label him robust with a large head. Lonnie detested bureaucracy and departmental business, whereas Sertoli became department chair early in his career and served until shortly before his death. Furthermore, whereas Sertoli never married, Lonnie crossed that bridge numerous times. Lonnie was tall and a very large man, whose stature was a command performance at every scientific meeting. He loved to debate and never shirked from raising a touchy question following the presentation of a paper at a meeting. So, it is interesting that the modern-day scientist who truly immortalized the name Sertoli was personally a contrast in many ways to the real Professor Sertoli. However, the two men did share many wonderful traits, not the least of which was that they studied the same cell. Both scientists loved microscopy, which is noted in the detailed drawings by Sertoli [15, 30] and the exceptional transmission electron microscopy by Russell [1, 39–41]. If there is an afterworld, surely the two “Sertologists” will still be searching for the holy grail, to prove the omnipotent theory [1] that Sertoli is the “mother cell.” The following is a poem written by Luiz Renato de França and dedicated to the memory of Lonnie D. Russell: From Sertoli to Lonnie: “Let my children go” Since the beginning of your life I took care of you Knowing your needs and my limits I grew up and developed as much as I could At some critical moments I took you in my arms And you deeply touched my soul Now, my dear friend It is time for you to go Good luck on your Journey And I will always be here for you

References 1. Russell, L. D., and Griswold, M. D., eds. (1993). “The Sertoli Cell.” Cache River Press: Clearwater, FL. 2. Ober, W. B., and Sciagura, C., (1981). Leydig, Sertoli and Reinke: Three anatomists who were on the ball. Pathol. Ann. 16, 1–13. 3. Negrini, F. (1910). Enrico Sertoli. Commemorazione del Prof. Francesco Negrini. Clinica Verterinaria 33, 146–161. 4. Negrini, F., et al. (1908). Onoranze al Prof. Enrico Sertoli. Clinica Veterinaria 31, 49–62. 5. Usuelli, F. (1934/1935). Enrico Sertoli (1842–1910). Ann. Veterinario Italiano 13, 455–461. 6. Taddia, C. (1985/1986). Istofisiologica dell cellule del Sertoli nei mamiferi. In “Facolta di Medicina Veterinaria.” Universita degli Studi di Milano, Italy.

7. Belloni, L. (1915). Enrico Sertoli in la medicina a Milano dal settecento al 1915. Storia Milano. Fondazione Treccani degli Alfieri, 1862 16, 1028. 8. Zanobio, B. (1970). Sertoli, Enrico. In “Dictionary of Scientific Biographies. American Council of Learned Societies” (C. C. Gillispie, ed.), pp. 319–320. Charles Scribner’s Sons, New York. 9. Internet websites used in the search for information on Sertoli: http://www.ibmsscience.org/history_zone/histology.htm http://www.geocities.com/hotsprings/2615/medhist/micro.html http://www.cas.muohio.edu/%7Embi-ws/microscopes/ index.html http://www.whonamedit.com/doctor.cfm/556.html http://www.sertoli.com/pages/568089/index.htm http://lsvl.la.asu.edu/paperproject/Microsopehistory/ http://www.az-microscope.on.ca/history.htm http://www.omni-optical.com/micro/sm101.htm http://www.nobel.se/medicine/articles/golgi/ http://www.arsmachina.com/micromenu.htm 2003. 10. Setchell, B. P. (1984). Male reproduction. In “Benchmark Papers in Human Physiology Series” (L. L. Langley, ed.), pp. 10–20. Van Nostrand Reinhold Company, New York. 11. Setchell, B. P. (1986). Some important contributors to our understanding of the male reproductive system: Monesi, Sertoli, Spallanzani and Aubry. Exerpta Medica Int. Cong. 716, 1–12. 12. Setchell, B. P. (1993). Foreword. In “The Sertoli Cell” (L. D. Russell and M. D. Griswold, eds.), pp. v–vi. Cache River Press, Clearwater, FL. 13. Andraos, J. (2002). “Nobel Prizes in Physiology & Medicine,” p. 12. Department of Chemistry, York University, Toronto. 14. Kölliker, A. (1863). “Handbuch der Gewebelehre des Menschen,” 4th ed. W. Logier, Berlin. 15. Sertoli, E. (1865). Dell’esistenza di particolari cellule ramificate nei canalicoli seminiferi del testicolo umano. Morgagni 7, 31–40. 16. Gernsheim, H. (1986). “The Concise History of Photography.” Thames & Hudson, London. 17. Kölliker, A. (1841). “Die samenfaden entwickeln sich aus oder in zellen, die sich zur zeit der geschlechtsreife oder der brunst in den hoden bilden, durch vorgänge, die den bei der entwicklung der thierischen elementartheile statt fidenden analog sind, von der gewöhnlichen entwicklung der thiere aus eiern dagegen bedeutend abweichen, in Beitrage zure Kenntnis der Geschlechtverhältnisse und der Samenflussigkeit wirbelloser Thiere.” W. Logier, Berlin. 18. Ehrlich, P. (1886). Fragekasten. Zeit Mikroskopie 3, 150. 19. Sertoli, E. (1886). Sur la caryokinése dans la spermatogénese. Arch. Italiennes Biologie 7, 369–375. 20. Leblond, C., and Clermont, Y. (1952). Definition of the stages of the cycle of the seminiferous epithelium in the rat. Ann. NY Acad. Sci. 55, 548–573. 21. Regaud, C. (1897). Sur la morphologie de la cellule de Sertoli et sur son role dans la spermatogénese chez les mammiféres. Compt. Rend. Anat. Paris 1, 21–31. 22. Ebner, V. V. (1888). Zur spermatogenese bei den säugethieren. Arch. Mikrosk. Anat. 31, 236–292. 23. Loisel, G. (1901). Origine et role de la cellule de Sertoli dans la spermatogenese. Compt. Rend. Soc. Biol. Paris IIs(iii), 974–977. 24. Bardleben, K. V. (1897). Die zwischenzellen des säugtierhodens. Anat. Anz. 13, 529–536. 25. 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. Contrib. Embryol. 210, 11–131. 26. Ebner, V. V. (1902). Mannliche geschlechtsorgne. In “Handbunch der Gewebelchre des Menschen” (A. Kolliker, ed.), p. 402. W. Engelman, Leipzig, Germany.

Chapter 1 History of the Sertoli Cell Discovery 27. La Vallette, S. V. (1878). Über die genese der samenkörper. Arch. Mikrosk. Anat. 15, 261–314. 28. Russell, L. D. (1993). Form, dimensions, and cytology of mammalian Sertoli cells. In “The Sertoli Cell” (L. D. Russell and M. D. Griswold, eds.), pp. 1–37. Cache River Press, Clearwater, FL. 29. Fawcett, D. W., and Burgos, M. (1956). The fine structure of the Sertoli cells in the human testis. Anat. Rec. 124, 401–402. 30. Sertoli, E. (1878). Sulla sturttura dei canalicoli seminiferi dei testicolo. Arch. Sci. Med. 2, 107–146, 267–295. 31. Attal, J., and Courot, M. (1963). Développement testiculaire et établissement de la spermatogenése chez le taureau. Ann. Biol. Animale Biochim. Biophys. 3, 219–241. 32. Orth, J. (1982). Proliferation of Sertoli cells in fetal and postnatal rats: A quantitative autoradiographic study. Anat. Rec. 203, 485–492. 33. Kormano, M. (1967). Dye permeability and alkaline phosphatase activity of testicular capillaries in the postnatal rat. Histochemie 9, 327–338. 34. Setchell, B. P., Voglmayr, J. K., and Waites, G. M. (1969). A blood–testis barrier restricting passage from blood into rete testis fluid but not into lymph. J. Physiol. (Lond.) 200(1), 73–85.

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35. Dym, M., and Fawcett, D. W. (1970). The blood–testis barrier in the rat and the physiological compartmentation of the seminiferous epithelium. Biol. Reprod. 3(3), 308–326. 36. Brown, H. H. (1885). On spermatogenesis in the rat. J. Microsc. Sci. 25, 343–370. 37. Roosen-Runge, E. C., and Giesel, Jr., L. O. (1950). Quantitative studies on spermatogenesis in the albino rat. Am. J. Anat. 87, 1–23. 38. Ortavant, R. (1954). Contribution à l’étude de la durée du processus spermatogénétique du Bélier à l’aide du 32P. Soc. Biol. Paris CR 148, 37–55. 39. Russell, L., and Clermont, Y. (1976). Anchoring device between Sertoli cells and late spermatids in rat seminiferous tubules. Anat. Rec. 185(3), 259–278. 40. Russell, L. D. (1979). l Further observations on tubulobulbar complexes formed by late spermatids and Sertoli cells in the rat testis. Anat. Rec. 194(2), 213–232. 41. Russell, L. D. (1980). Sertoli–germ cell interactions: A review. Gamete Res. 3, 179–202. 42. Walker, C. E., and Embleton, A. L. (1906). On the origin of the Sertoli or foot-cells of the tesis. Proc. Royal Soc. Lond. 1(xxviii), 50–52. 43. Hanes, F. M. (1910). The biological significance of the Sertoli cells. Proc. Soc. Exp. Biol. Med. 7, 136–137.

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C H A P T E R

2 Perspective on the Function of Sertoli Cells MICHAEL D. GRISWOLD Center for Reproductive Biology, School of Molecular Biosciences, Washington State University, Pullman, Washington

I. INTRODUCTION II. THE VIEW IN 1993 III. CHANGES TO OUR VIEW IN THE LAST TEN YEARS IV. IMPORTANT QUESTIONS REMAIN V. THE “PERMISSIVE” VIEW VI. THE NEXT TEN YEARS References

II. THE VIEW IN 1993 We started our discussion in 1993 with the assumption originally stated by Enrico Sertoli that Sertoli cells are “linked to the production of spermatozoa.” The extent to which that link is mandatory was unknown. Clearly, both testosterone and follicle-stimulating hormone (FSH) acted on the Sertoli cells and the result of those actions had measurable impacts on spermatogenesis. At the time, some investigators believed that Sertoli cells regulated virtually all aspects of germ cell development (the dogmatic Sertologists), whereas others believed that germ cell development was affected by Sertoli cell functions but that most germ cell development appeared to be autonomous (germ cell worshiping cult).

I. INTRODUCTION More than 10 years have passed since Lonnie Russell and I sat down with a few beers at my home in Idaho and wrote the Preface to The Sertoli Cell [1]. This book was the first major work dedicated to our favorite cell. We pointed out that there are “dogmatic, religious Sertologists” who believed in the so-called “omnipotent” Sertoli cell theory and that there was a “germ cell worshiping cult” who believed in only minimal involvement by Sertoli cells. The reaction of my colleagues to that irreverent and impertinent preface was interestingly mixed. Some thought that it was outside of the bounds of serious science, whereas others thought it was the most useful part of the book. Despite the light tone of our writing, we did raise serious questions relating to the function of Sertoli cells as we understood them in 1993. Have any or all of those questions been answered in the intervening decade? Have new questions surfaced with the new research? SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

III. CHANGES TO OUR VIEW IN THE LAST TEN YEARS The last decade has not been kind to the league of dogmatic Sertologists of which both Lonnie and I were charter members. Two general categories of experiments including germ cell culture and germ cell transplantation have established that most germ cell development is quite autonomous. In 2002 the Dym laboratory published convincing evidence that they had established a cell line that underwent meiosis and spermatogenesis in vitro in the absence of Sertoli cells [2].

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Michael D. Griswold

They reported the in vitro generation of spermatocytes and spermatids from telomerase-immortalized mouse type A spermatogonial cells in the presence of stem cell factor. Meiosis was apparent but the round spermatids never developed tails. Further studies will be necessary to determine if these cells can function as “normal” haploid germ cells, but the work clearly illustrates the potential for germ cell development in the absence of somatic cell input. Several other studies on invertebrates and lower vertebrates such as teleosts have also shown successful spermatogenesis in vitro in the absence of somatic cells [3]. Germ cell transplantation has been used to demonstrate that spermatogonial stem cells can be maintained in culture in the absence of Sertoli cells for several months [4]. Transplantation of the cultured germ cells into recipients demonstrated that functional stem cells were present in the cultures. Germ cell transplantation has also provided evidence that the germ cells in the testis are in control of their own fate. The transplantation of rat germ cells into the mouse testis showed that rat stem cells can develop and go through meiosis and form relatively normal elongated spermatids even though they interacted with mouse Sertoli cells [5]. In addition, the time required for the completion of this process in each mammalian species is unique and fixed. Franca et al. [5] used the technique of spermatogonial transplantation to ask the question as to which cell type(s) determined the rate at which germ cells proceeded through spermatogenesis. Rat germ cells were transplanted into a mouse testis, and the mouse was euthanized 12.9–13 days after administration of a single dose of [3H]thymidine. The investigators found that two separate timing programs existed for germ cell development in the recipient mouse testis: one of rat and one of mouse duration. Rat germ cells that were supported by mouse Sertoli cells always differentiated with the timing characteristic of the rat and generated the spermatogenic structural pattern of the rat, demonstrating that the timing of the cell differentiation process of spermatogenesis was regulated by germ cells alone.

IV. IMPORTANT QUESTIONS REMAIN Taken together the experiments described above show that much of the development of germ cells can occur relatively independent of Sertoli cells and that the timing of the process is intrinsic to the germ cells. So where does that leave the views of the “dogmatic Sertologist”? Successful and complete spermatogenesis resulting in sperm capable of fertilization in the absence of Sertoli cells still has not been demonstrated

in mammals. So, although the germ cells apparently have a great deal of developmental autonomy, there still appears to be a Sertoli cell requirement for complete and successful spermatogenesis. In the models for germ cell–Sertoli cell interactions that Lonnie Russell and I proposed in 1993 we suggested that important molecular communications occurred between germ cells and Sertoli cells but we pointed out that not one signal from either of these cell types had been identified with certainty. Although a number of growth factors have been shown to affect germ cell–Sertoli cell interactions in vitro, and growth factors that affect stem cells in vivo have been found, the identification of a distinct signaling system between Sertoli cells and germ cells undergoing spermatogenesis remains elusive.

V. THE “PERMISSIVE” VIEW We also suggested in 1993 that the action of hormones on spermatogenesis may be primarily permissive in nature. This concept of the Sertoli cell acting in a permissive way fits well with the “support cell” or “nurse cell” view and it may be useful to view the major function(s) of Sertoli cells as “permissive.” In his 1994 review article, Sharpe [6] suggested that we view the function of Sertoli cells and their response to hormones as affecting the efficiency of spermatogenesis. Adapting these views to current understanding would be to suggest that Sertoli cells provide the environment, structural organization, and biochemical milieu that allow the efficient but autonomous development of germ cells into spermatozoa. The knockout of several genes that are unique to Sertoli cells in the testis has supported the view of the “germ cell worshiping cult” or the modified “permissive” view. In the past few years, the knockout of the FSH receptor and the FSHβ gene gave emphasis to the permissive nature of FSH actions. The FSH receptor and the FSHβ knockout males were fertile but produced reduced numbers of sperm in a smaller than normal testis [7–9]. In Table 2.1 a number of known Sertoli cell products whose genes have been knocked out by naturally occurring mutations or by homologous recombination are listed. Note that there is consistency in the description of the spermatogenic phenotypes. The elimination of these genes associated with the function of Sertoli cells, in general, leads to lowered sperm counts but does not totally block spermatogenesis. The spermatogonial phenotype of the androgen insensitivity or insufficiency models still provides the “dogmatic Sertologists” some solace. The action of testosterone on spermatogenesis is more difficult to visualize as “permissive” because, in the absence of

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Chapter 2 Perspective on the Function of Sertoli Cells TABLE 2.1 Examples of Naturally Occurring and Engineered Knockout of Genes Expressed in Sertoli Cells and the Resulting Spermatogenic Phenotype Gene

Phenotype

Reference

FSH receptor

Fertile, underdeveloped testis with 50% reduction in Sertoli cells

[16]

Inhibin α

Initially develops normally, but ultimately develops gonadal stromal tumor

[16]

Transferrin

Fertile but with abnormalities in spermatogenesis

[17]

Clusterin or SGP-2

Fertile with minor abnormalities in spermatogenesis

[18]

Cyclic protein 2 or cathepsin L

Fertile with increased numbers of atrophic tubules

[19]

Pem- androgen regulated homeobox

Fertile and normal

[20]

Phosphatidylserine synthase 2

Lowered fertility

[21]

Fyn tyrosine kinase

Fertile, a significant reduction in testis weight and degenerated germ cells were observed at 3 and 4 wk of age

[22]

Desert hedgehog

Anastomotic seminiferous tubules, pertitubular cell abnormalities, and absence of adult-type Leydig cells

[23]

Bclw

Initially fertile with progressive degeneration

[24]

Tyro 3, Axl, and Mer tyrosine kinases

Mice lacking any single receptor, or any combination of two receptors, are viable and fertile, but male animals that lack all three receptors progressively produce no mature sperm

[25]

Gata 1

Tissue specific knockout, no altered phenotype, normal spermatogenesis

[26]

response to testosterone, a condition such as is seen in the testicular feminized mutant (tfm) mice and humans, spermatogenesis is clearly blocked at meiosis [10]. Note that the interpretation of the action of androgens in the tfm condition is complicated by cryptorchidism. The phenotype of the leuteinizing hormone (LH) receptor knockout differs in that spermatogenesis is blocked at the round spermatid stage [11]. In other models blocking androgen action such as EDS-treated rats, or hypophysectomy, some spermatids are found [12]. The abundance of evidence can be interpreted to indicate that a testosterone-dependent signal or product from Sertoli cells is mandatory for successful spermatogenesis. Sertoli cells may be essential for several processes but the clearest effect is on the elongation of spermatids and the formation of sperm tails. A recent publication using gene chip technology looked at the action of testosterone on the GnRH-deficient mutant mouse (hypogonadal; hpg). Hpg mice age 35–45 days were injected subcutaneously with 25 mg testosterone propionate, and the animals were sacrificed 4, 8, 12, and 24 hr after treatment. Although the expression of many genes was changed, at early time points the expression of more genes was depressed rather than stimulated, suggesting that a primary effect of androgens may be to inhibit the expression of genes that maintain the prepubertal condition [13]. The role of the Sertoli cells in testis formation is much easier to reconcile with the “dogmatic Sertologists” view. The initial expression of Sry and other genes involved

in testis formation is clearly an important function of Sertoli cells and a prerequisite for the formation of a testis [14]. Other potential functions of Sertoli cells in early testis development could include the inhibition of meiosis and the regulation of the mitosis of gonocytes. The fetal testis of the mouse (presumably the Sertoli cells) produces a putative inhibitor of meiosis, and germ cells that are exposed to it develop as prospermatogonia [15]. None of the agents that allow Sertoli cells to regulate meiosis and mitosis have been identified as yet.

VI. THE NEXT TEN YEARS Predictions about the contents of a third book dedicated to the Sertoli cells are almost entirely speculation, but some areas of progress can be anticipated based on current technology. If there is a third book dedicated to Sertoli cells and if it is published about 10 years from now, we will have a nearly complete knowledge of the genes expressed in Sertoli cells throughout their development and we will know what many of these genes do (at least at the physiological level). We will probably have candidates for a complete picture of genes that influence testis determination, suppression of meiosis in the embryonic and prepubertal mammals, stimulation of meiosis during puberty, and for influencing the morphological development of gonocyte to spermatozoa. This knowledge will provide for much

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Michael D. Griswold

greater insight into genetic diseases that affect fertility. We will probably have the capability to maintain most germ cells in culture in the absences of Sertoli cells and then to transition them to haploid cells in vitro. We will have in vitro systems that will produce gametes capable of successful fertilization. These systems will enable the efficient and routine genetic manipulation of gametes. What will we still lack in 10 years? It is unlikely that we will have a contraceptive that is targeted to the Sertoli cells. Several factors have led to this prediction including the evidence presented in Table 2.1 that interfering with Sertoli cell functions may impede the efficiency of spermatogenesis but may not cause complete aspermatogenesis. It is likely that the predictions made here will be about as accurate as my annual picks in the NCAA basketball pool, where I usually end up last. However, one prediction can be made with relative assurance of accuracy. Investigators will continue to be fascinated by the beauty and complexity of Sertoli cells, and there will be further pursuit of the original premise of Enrico Sertoli that, the cells are somehow linked to the production of spermatozoa.

References 1. Russell, L. D., and Griswold, M. D., eds. (1993). Preface. In “The Sertoli Cell,” pp. 365–390. Cache River Press, Clearwater, FL. 2. Feng, L. X., et al. (2002). Generation and in vitro differentiation of a spermatogonial cell line Science 297, 392–395. 3. Saiki, A., et al. (1997). Establishment of in vitro spermatogenesis from spermatocytes in the medaka, Oryzias latipes. Dev. Growth Differ. 39(3), 337–344. 4. Nagano, M., et al. (1998). Culture of mouse spermatogonial stem cells. Tissue Cell 30(4), 389–397. 5. Franca, L. R., et al. (1998). Germ cell genotype controls cell cycle during spermatogenesis in the rat. Biol. Reprod. 59(6), 1371–1377. 6. Sharpe, R. M. (1994). Regulation of spermatogenesis. In “The Physiology of Reproduction,” 2nd ed. (E. Knobil and J. D. Neill, eds.), pp. 1363–1433. Raven Press, New York. 7. Kumar, T. R., et al. (1997). Follicle stimulating hormone is required for ovarian follicle maturation but not male fertility. Nat. Genet. 15(2), 201–204. 8. Dierich, A., et al. (1998). Impairing follicle-stimulating hormone (FSH) signaling in vivo: Targeted disruption of the FSH receptor

9.

10.

11.

12.

13. 14. 15. 16. 17.

18.

19.

20.

21.

22. 23.

24. 25. 26.

leads to aberrant gametogenesis and hormonal imbalance. Proc. Natl. Acad. Sci. USA 95(23), 13612–13617. Wreford, N. G., et al. (2001). Analysis of the testicular phenotype of the follicle-stimulating hormone beta-subunit knockout and the activin type II receptor knockout mice by stereological analysis. Endocrinology 142(7), 2916–2920. Fritz, I. (1978). Sites of actions of androgens and follicle stimulating hormone on cells of the seminiferous tubule. In “Biochemical Actions of Hormones” (G. Litwack, ed.), pp. 249–278. Academic Press, New York. Zhang, F. P., et al. (2001). Normal prenatal but arrested postnatal sexual development of luteinizing hormone receptor knockout (LuRKO) mice. Mol. Endocrinol. 15(1), 172–183. Kerr, J. B., Maddocks, S., and Sharpe, R. M. (1992). Testosterone and FSH have independent, synergistic and stage-dependent effects upon spermatogenesis in the rat testis. Cell Tissue Res. 268(1), 179–189. Sadat-Ngatchou, P., McLean, D. J., and Griswold, M. D. (in press). Biology of Reproduction. Lovell-Badge, R. (1992). The role of Sry in mammalian sex determination. Ciba Found. Symp. 165, 162–182. McLaren, A. (1995). Germ cells and germ cell sex. Philos. Trans. R. Soc. Lond. B Biol. Sci. 350(1333), 229–233. Matzuk, M. M., et al. (1992). Alpha-inhibin is a tumour-suppressor gene with gonadal specificity in mice. Nature 360(6402), 313–319. Sylvester, S. R., and Griswold, M. D. (1994). The testicular iron shuttle: A “nurse” function of the Sertoli cells. J. Androl. 15(5), 381–385. Bailey, R. W., et al. (2002). Heat shock-initiated apoptosis is accelerated and removal of damaged cells is delayed in the testis of clusterin/ApoJ knock-out mice. Biol. Reprod. 66(4), 1042–1053. Wright, W. W., et al. (2003). Mice that express enzymatically inactive cathepsin L exhibit abnormal spermatogenesis. Biol. Reprod. 68(2), 680–687. Pitman, J. L., et al. (1998). Normal reproductive and macrophage function in Pem homeobox gene-deficient mice. Dev. Biol. 202(2), 196–214. Bergo, M. O., et al. (2002). Defining the importance of phosphatidylserine synthase 2 in mice. J. Biol. Chem. 277(49), 47701–47708. Maekawa, M., et al. (2002). Fyn tyrosine kinase in Sertoli cells is involved in mouse spermatogenesis. Biol. Reprod. 66(1), 211–221. Pierucci-Alves, F., Clark, A. M., and Russell, L. D. (2001). A developmental study of the Desert hedgehog-null mouse testis. Biol. Reprod. 65(5), 1392–1402. Russell, L. D., et al. (2001). Spermatogenesis in Bclw-deficient mice. Biol. Reprod. 65(1), 318–332. Lu, Q., et al. (1999). Tyro-3 family receptors are essential regulators of mammalian spermatogenesis. Nature 398(6729), 723–728. Lindeboom, F., et al. (2003). A tissue-specific knockout reveals that Gata1 is not essential for Sertoli cell function in the mouse. Nucleic Acids Res. 31(18), 5405–5412.

C H A P T E R

3 Structure of the Sertoli Cell

I. II. III. IV. V. VI.

REX A. HESS

LUIZ R. FRANÇA

Reproductive Biology & Toxicology, Veterinary Biosciences, University of Illinois, Urbana, Illinois

Laboratory of Cellular Biology, Department of Morphology, Institute of Biological Sciences, Federal University of Minas Gerais, Belo Horizonte, Brazil

likely the work of Don Fawcett at Harvard in the 1950s [3, 4] that firmly established the importance of studying Sertoli cell morphology, because from 1951 until 1973, the year that Lonnie Russell published his first manuscript on testicular morphology [5], the number of publications dealing with the Sertoli cell jumped to approximately 440. From 1973 to 1990, about 4000 Sertoli cell papers were published, which is an indication of the recognition of the importance of Sertoli cells in testicular function. Such growth in Sertoli cell interest is a direct result of the tremendous impact that morphological studies, especially those of Dr. Russell, have had on the reproductive sciences.

BRIEF HISTORY FORM AND FUNCTION NUCLEUS AND NUCLEOLUS CYTOPLASM THE SERTOLI (BLOOD–TESTIS) BARRIER MISCELLANEOUS OBSERVATIONS References

I. BRIEF HISTORY The Sertoli cell received its family name in a paper published by von Ebner [1] in which he described the cells as “the cells of Sertoli.” It is amazing that the early scientists were able to deduce the cell’s basic structure so well, especially when consideration is given to the poor resolution of microscopes in the 1800s (see Chapter 1) and the lack of proper fixation, embedding, sectioning, and staining. In fact, Sertoli’s original observations were so intuitive that few scientists at that time bothered to study the cell in great depth, because most everyone accepted his descriptions and went on to other more important topics of the day. Most scientists waited for the improved resolution provided by the electron microscope before returning to the study of the Sertoli cell. From 1865, when Sertoli published his famous observations, until 1950, approximately 85 manuscripts were published with Sertoli cell or other descriptive names for this cell in the title or as key words. Then in 1953 the first paper to observe the testis with the transmission electron microscope was published [2]. However, it was most SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

II. FORM AND FUNCTION The total number of Sertoli cells establishes the upper limit of sperm production by the testis [6–11], and spermatogenic efficiency is highly correlated with Sertoli cell support capacity, which is the best indication of Sertoli cell function [12]. Sertoli cell individual volume shows high variation in mammals, varying from approximately 2000–3000 μm3 to 6000–7000 μm3 in mammals already investigated [11–13]. However, paradoxically, it is generally recognized that species in which Sertoli cell volume is high are those with the lowest spermatogenic efficiency. The volume density of Sertoli cells in the seminiferous epithelium also changes considerably in mammals, and the mean figures range from approximately 15% in mice and rabbit

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Copyright 2005, Elsevier Science (USA). All rights reserved.

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Rex A. Hess and Luiz R. França

FIGURE 3.1 (A) Drawing of a reconstructed rat Sertoli cell surface (stage V). Numerous indentations and sheet-like cytoplasmic processes are noted to provide crypts for developing spermatids and recesses for spermatocytes and spermatogonia. (Modified and reprinted from Wong and Russell [139], Copyright 1983 Am. J. Anat., by permission of Wiley Liss, Inc., a subsidiary of John Wiley & Sons, Inc.). (B) Drawing of a Sertoli cell cytoplasm showing the inclusion of elongate spermatid nuclei within deep crypts. (Modified and reprinted from Fawcett [140] by permission of the American Physiological Society.)

to 40% in the woodchuck and humans [11, 13]. As stated for the cell volume, it is also generally noted that species with lower Sertoli cell occupancy in the seminiferous tubule epithelium, such as mice, rabbits, rats, and hamsters, are those with higher Sertoli cell and spermatogenic efficiencies [11]. The Sertoli cell, which is columnar in height (Fig. 3.1) and assumed to always extend from the basement membrane of the seminiferous epithelium to the lumen [13], performs its nurse-like cell functions by extending its cytoplasm in thin arm-like processes in two dimensions and sheet-like or cylindrical processes in three dimensions around the developing germ cells (Fig. 3.2) and by forming specialized junctional complexes that consist of gap and tight junctions, actin filaments, and smooth endoplasmic reticulum [14–20]. Approximately 40% of the Sertoli cell surface (Fig. 3.2) contacts the surface of elongated spermatids [21, 22], which illustrates the extent to which the Sertoli cell stretches its cytoplasm to communicate directly with the developing germ cells. Enrico Sertoli used the term mother cells on the first page of his publication [23], suggesting that this cell type served a unique function in its relationship to the developing germ cells. Indeed, the Sertoli cell cytoplasm is indented by the germ cells (Figs. 3.1–3.4) in every stage of the cycle of the seminiferous epithelium, with certain stages showing tremendous indentation

FIGURE 3.2 Ultrastructural section through a single rat Sertoli cell at a level where the cell surrounds nine step 9 elongating spermatids (GC; 1–9). The Sertoli cell edge is enhanced with black ink to demonstrate the tremendous volume density of its plasma membrane.

Chapter 3 Structure of the Sertoli Cell

21

FIGURE 3.3 Sertoli cell variation within the cycle of the seminiferous epithelium is illustrated with sections from the rat stage V (Type A cell) and late stage VI (Type B cell). In stage V, the elongate spermatid (ES) heads are found deep within crypts of the Sertoli cell (SC), adjacent to round spermatids (RS) and pachytene spermatocytes (PS), with the movement being toward the basement membrane in the Type A cell (arrow). In late stage VI, the elongate spermatids (ES) are found near the tubule lumen, with round spermatids and pachytene spermatocytes. The drawing of a Type B cell shows the Sertoli cell membrane encapsulating spermatid heads near the lumen, with movement of the spermatids in a proluminal direction (arrow).

or deep crypts, as seen in stage V in the rat, and often referred to as Type A Sertoli cells [13]. Type B Sertoli cells are those that support the movement of elongate spermatids toward the lumen, as seen starting in stages VI to VIII in the rat (Fig. 3.3). Thus, spermiation appears to separate these two basic structural features of the Sertoli cell. The cycle of the seminiferous epithelium is also associated with other Sertoli cell changes, the most notable being its relative volume, with volume density being smallest in stages VII and VIII in the rat and greatest in stages XII through XIV [11, 24, 25], which reflects the unique ability of this cell to alter its form and function, without noticeable alteration in its volume [26], throughout the cycle.

III. NUCLEUS AND NUCLEOLUS One of the first features of the Sertoli cell that is taught to new students is the unique appearance of the nucleus and its tripartite nucleolus (Fig. 3.5). In most species, the nucleus is found very near to the base of the cell, if not sometimes appearing to rest on the basement membrane. Fixation and embedding techniques make huge differences in the appearance of the nucleus, with neutral buffered formalin fixation and

paraffin embedding resulting in the most distorted appearance, even making it difficult to recognize some Sertoli cells (Fig. 3.5). Glutaraldehyde provides the optimum fixation, and epoxy resin embedment provides the highest resolution of nuclear detail, even compared to glycol methacrylate. The Sertoli cell nucleus is large (250–850 μm3) and can take on several different shapes depending on the stage of the seminiferous cycle [11] and age of development (Fig. 3.6). From birth to adulthood, the nucleus is often elongated, extending toward the lumen, but there are definite changes in appearance as the testis matures. The early support cells of the seminiferous tubule are often called pre-Sertoli cells because the nuclear appearance differs from the adult form, yet in some species the pre-Sertoli cells still have the typical nuclear features of irregular shape and a prominent nucleolus [27]. The large tripartite nucleolus and the numerous satellite karyosomes or chromocenters [28], which are prominent in the adult Sertoli cell nucleus, are smaller or less abundant in the developing testis (Fig. 3.6). For instance, on day 5 postbirth it is easy to recognize rat Sertoli cell nuclei, because they are the only nuclei located along the basement membrane. Between day 5 and 10 postbirth, however, germ cells migrate to the basement membrane and some Sertoli cell nuclei

FIGURE 3.4 A montage of the typical Sertoli cell. The organelles depicted are for illustrative purposes and not necessarily photographed from the cell shown at lower magnification. In the base of the Sertoli cell, the nucleus (Nu) contains a large nucleolus (Nuc) and satellite karyosomes (Sk). In this region, lipid droplets (Li) are often noted and the Sertoli cell–Sertoli cell tight junctional complexes are found (Jct). The body of the Sertoli cell cytoplasm contains the typical epithelial cell organelles and inclusion bodies: a Golgi apparatus (Go) that is small compared to most secretory cells; an abundance of rough endoplasmic reticulum (RER); smooth endoplasmic reticulum (SER) that is often found adjacent to mitochondria (Mit); Mit are either elongate parallel to microtubules (Mt) or circular with a donut-like shape; glycogen granules can be seen near the nucleus but other inclusions such as multivesicular bodies (MVB) are seen scattered throughout the cytoplasm; lysosomes (Ly) are usually seen near the MVB and typically near the apical border in stages where spermiation has occurred; unique structures called ectoplasmic specializations (Eps) are found adjacent to the spermatid heads; and tubulobulbar complexes (Tub) are seen with higher magnification within the curvature of the late spermatid head and represent a Sertoli cell invagination or penetration by spermatid cytoplasm, which participates in attachment of elongating spermatids to Sertoli cells and in the elimination of excess cytoplasm prior to spermiation.

Chapter 3 Structure of the Sertoli Cell

23

FIGURE 3.5 Recognition of Sertoli cells in histological sections depends on several factors including quality of fixation, the embedding material, and thickness of the section. Shown here are different Sertoli cell nuclei taken from different stages of spermatogenesis in adult mouse testes embedded in different media. (A–E) Mouse testis fixed with Bouin’s, embedded in paraffin, and stained with PAS/hematoxylin. Most nuclei are smaller in size than in the plastic resins. The euchromatin appear washed out, and satellite nucleoli (S) are difficult to recognize next to the nucleolus (Nuc). (F–J) Mouse testis fixed by vascular perfusion with 4% glutaraldehyde, embedded in glycol methacrylate (GMA), and stained with PAS/hematoxylin. The deep nuclear indentations (arrows) are visible, and prominent nucleoli (Nuc) are easily recognized along with the satellite karyosomes (S). (K–O) Mouse testis fixed by vascular perfusion with 4% glutaraldehyde, embedded in epoxy resin for electron microscopy, sectioned 1 μm for light microscopy and stained with toluidine blue. Nuclear indentations are prominent (arrow), and the euchromatin (E) show greater detail than with the GMA embedding.

become oval in shape (Fig. 3.6), which makes their recognition without special markers rather difficult. By day 25, nearly all Sertoli cell nuclei are displaced from the basement membrane by germ cells, and it is not until the mature testis is formed that the nuclei begin to appear along the base again in association with specific stages of spermatogenesis. Pituitary gonadotrophins have been shown to increase nuclear diameter [29].

In the adult testis, nearly all Sertoli nuclei contain deep indentations of their nuclear envelope (Fig. 3.7), which gives the nucleus irregular shapes that have been called pyramidal, triangular and even “planoconvex” [13]. Such invaginations of the nuclear envelop are invested with vimentin intermediate filaments, but it is not known if the filaments help to maintain the structure or if the filaments are better anchored in such

FIGURE 3.6 Sertoli cell nuclei change in appearance and density from early development to adulthood. Shown here are rat testes from day 5 to day 100. SC, Sertoli cell; GC, germ cells. (A–B) Day 5 postnatal SC nuclei (box and arrows) are packed tightly along the basement membrane, with considerable variation in the shape, ranging from triangular to oval. A single prominent nucleolus is present as are small satellite karyosomes that are usually found along the nuclear envelope. Gonocytes GC are located in the center of the seminiferous cords. (C) Day 15 SC nuclei are larger in size and appear more angular than on day 5. Some are displaced from the basement membrane by the migration of GC. Satellite chromocenters (arrow) are more prominent in the SC nuclei. (D) Day 25 SC nuclei are completely displaced from the basement membrane by GC. The nuclei now show deep invaginations and several satellite karyosomes (arrow). (E) Day 35 SC nuclei still contain an abundance of satellite karyosomes and most remain displaced away from the base. (F) Day 100 SC nuclei show the typical adult form lying along the basement membrane, with euchromatic appearance and a prominent tripartite nucleolus.

Chapter 3 Structure of the Sertoli Cell

25

FIGURE 3.7 Sertoli cell nucleus showing typical indentations (arrows) and prominent nucleolus, lying near the basal lamina (Bl). Lateral Sertoli cell–Sertoli cell junctions (Jct) are seen on both sides of the cell. Peritubular myoid and Leydig cells are noted beneath the seminiferous tubule.

structures. Information regarding the function of these nuclear indentations is limited; however, one study has shown that some proteins can be found in higher concentrations at nuclear invagination areas [30]. The nucleoplasm is typically euchromatic and stains very light blue with hematoxylin due to the relatively small amount of heterochromatin (Fig. 3.7) that is scattered finely in the nucleus and along the membrane (Fig. 3.8), particularly in the developing testis. Unusual formations, such as crystalloids, have been reported in the Sertoli cell nuclei of large mammals, such as the bull [31], but in most species the Sertoli cell nucleus has evenly distributed chromatin with only the nucleolus standing out as a distinct intranuclear structure. Using cytochemistry, however, it has been possible to observe other components, such as calcium, which were described as “large round-shaped” precipitates [32]. The nucleolus is large and easily recognized (Figs. 3.5–3.7), and the entire complex is often found in three distinct parts (tripartite; Figs. 3.9 and 3.10) [33, 34] in many species. With hematoxylin staining, the nucleolus and satellite structures are strongly basophilic. Only the main nucleolar body is seen in most histological and ultrastructural sections (Fig. 3.5), and although this could be due to the thickness of sections, it has also been found that the number of large chromocenters varies between 1 and 10 or more [28]. The large satellite chromocenters are usually considered part of the tripartite nucleolus structure (Figs. 3.5, 3.7, 3.9, 3.10), but have limited participation in RNA synthesis [35];

nevertheless, the chromocenters contain nearly all of the heterochromatin found in the Sertoli cell nucleus [28]. Others have localized decondensed DNA in the fibrillar components and other regions of the nucleolus [33, 36],

FIGURE 3.8 Sertoli cell nucleus showing a deep indentation containing intermediate filaments (Fi). Nuclear pores (Np) are seen along the nuclear envelope (Ne). A thin proteinaceous band crosses between the nuclear envelope at the opening of the pores. Heterochromatin (Ht) forms a very thin line attached to the nuclear envelope.

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Rex A. Hess and Luiz R. França

FIGURE 3.9 Sertoli cell nucleus sectioned through the two prominent satellite chromocenters, with the nucleolus out of the plane of section. Nuclear pores are barely seen because the chromatin is so fine with few clumps along the nuclear membrane. The Golgi complex is seen adjacent to the nucleus in three small sections.

FIGURE 3.10 The tripartite nucleolus often seen in the Sertoli cell nucleus. The vesicular nucleolus body is flanked by more dense satellite chromocenters that are attached to the nucleolus by thin filaments (arrowheads).

suggesting that transcriptional activity may be active in this region, but it remains to be determined whether this activity has overall significance. The chromocenters are smaller and more numerous in the developing testis and there is a continuous decrease in numbers with age, but the number was not shown to be correlated with androgen receptor presence, although there were distinct mouse strain differences [28]. It is interesting that the X chromosome was found near the center of one of the chromocenters, whereas the Y chromosome was found at the nuclear periphery [33]. Telomeric sequences were shown by in situ hybridization to give a strong signal in the chromocenters [33]. The Sertoli cell nucleus has a high density of nuclear pores (Fig. 3.8), the number of which has been shown to vary depending on the stage of the spermatogenic cycle, with the highest density appearing to occur in stages XIII–I of the rat [37]. It was suggested that this increase in density corresponded with the apparent increased metabolic needs of the Sertoli cell. Species differences are found in Sertoli cell morphology, particularly the nucleus. To illustrate this aspect, note that the nucleolus in ruminant Sertoli cells is heavily vacuolated (Fig. 3.11) or multivesiculated [38, 39]. In the monkey testis, Sertoli cell nuclei are typically located approximately in the middle of the seminiferous epithelium, in contrast to most other species, including rodents, where the nuclei are typically near the basement membrane (Fig. 3.12), except during spermiation [40].

FIGURE 3.11 Nucleus of the bovine Sertoli cell showing indentations of the nuclear envelope and prominent vesiculation of the nucleolus. (Used with the permission of Dr. Karl-Heinz Wrobel, University of Regensburg, Germany.)

Chapter 3 Structure of the Sertoli Cell

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FIGURE 3.12 Seminiferous tubules from monkey and mouse testes at similar stages of spermatogenesis. Sertoli cell nuclei are encircled to illustrate species differences in numerical density of Sertoli cells per tubule cross section. Also, note that the nuclei are located midway between basement membrane and lumen in the monkey tubule, but along the basement membrane in the mouse.

IV. CYTOPLASM A. Mitochondria Sertoli cell cytoplasm contains an abundance of mitochondria, which is indicative of its high metabolic activity [13]. In the body of the cell, mitochondria are more numerous (Fig. 3.13) and scattered among the other organelles. Mitochondrial shape seems to vary more in the Sertoli cell than in most other epithelial cell types, and these morphological appearances are species specific and can be used to differentiate between Sertoli and germ cells [41, 42]. The mitochondria can be rather long (Fig. 3.14), even reaching 2–3 μm [13] or more, but they can also appear to be “cup shaped” or even shaped like “donuts” (Figs. 3.4, 3.13, 3.15). Mitochondrial cristae of the Sertoli cell are predominantly tubular, but the orthodox foliate forms are also present [13]. The number of cristae, however, is less than that typically found in steroid-producing cells such as the Leydig cell [13]. Cytoplasmic organelles of the Sertoli cell are usually polarized within the cell body by the extensive tracts of microtubules that travel from base to apex (Fig. 3.16). When the testis is exposed to a microtubule inhibitor, such as the fungicide carbendazim [43, 44], the organelles collapse into a perinuclear region, particularly the mitochondria (Fig. 3.16).

B. Endoplasmic Reticulum Rough endoplasmic reticulum is located in the basal region and is sparse [13], but smooth endoplasmic reticulum is a predominant organelle in adult Sertoli cells (Figs. 3.4 and 3.16), suggesting that the cell has a predominant function related to the metabolism of lipids or steroids. It is often found adjacent to mitochondria (Figs. 3.4 and 3.13–3.15) and in some species is found evenly dispersed between the elongate spermatid head near the tubule lumen or surrounding lipid droplets [13]. In the water buffalo, aggregates and whorls of smooth endoplasmic reticulum can be quite large in the base of the cell, as well as near the heads of elongate spermatids [38]. In the body of Sertoli cells from all species studied, microtubules help to maintain the linear appearance of the smooth endoplasmic reticulum and the alignment of mitochondria (Fig. 3.16). Apparently this reticulum is a continuous structure, which includes thin branches that surround the heads of elongate spermatids at the tubulobulbar complex, a spermatid projection into the Sertoli cell cytoplasm (Fig. 3.17). The smooth endoplasmic reticulum is also a component of the ectoplasmic specialization, which composes the unique Sertoli/germ cell junctional complex (Figs. 3.18 and 3.19). It has been suggested that the amount of smooth endoplasmic reticulum in Sertoli cells might be

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FIGURE 3.13 Sertoli cell cytoplasm in the body of the cell. Germ cells (GC) are outlined by the black lines. Mitochondria (M1) are aligned in the microtubule direction but some mitochondria have the unique donut shape (M2), whereas others have a C-shape (M3) and are closely aligned to smooth endoplasmic reticular structures (S). L, lysosome.

FIGURE 3.14 Sertoli cell mitochondria (Mi) near the basal lamina (Bl) are long, thin, vesiculated structures that are easily distinguished from the germ cell mitochondria that exhibit wide spaces in their lamellae. ER, endoplasmic reticulum; Ly, lysosome; Li, lipid droplet.

FIGURE 3.15 Sertoli cell base lying on the basement membrane (BM), which contains an amorphous layer next to the Sertoli cell plasmalemma and small collagen fibrils. Rare, small endocytotic or exocytotic vesicles (En) are found along the basal membrane. The thin peritubular myoid cell (PM) is found adjacent to the basement membrane. Mi, Sertoli cell mitochondrion; ER, endoplasmic reticulum.

Chapter 3 Structure of the Sertoli Cell

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FIGURE 3.16 (A) Sertoli cell cytoplasm in a control testis showing thin microtubules (arrows) oriented in the long axis from base to apical regions (double arrow). Organelles such as mitochondria (M) and endoplasmic reticulum (ER) are oriented in the direction of the microtubules. (B) Sertoli cell cytoplasm in a testis exposed to the fungicide carbendazim, a microtubule poison. The mitochondria are no longer oriented longitudinally along microtubule tracts, but are collapsed (circle) near the Sertoli cell nucleus (SC).

FIGURE 3.17 Tubulobulbar complexes are seen penetrating into the apical cytoplasm of the Sertoli cell from the adjacent germ cell. The complexes end with bulbous heads that contain a fuzzy outer layer on the Sertoli cell side. SER, smooth endoplasmic reticulum.

FIGURE 3.18 Sertoli cell cytoplasm adjacent to the developing spermatid nucleus is a highly specialized attachment site that is given the name ectoplasmic specialization. The ectoplasmic specialization consists of the spermatid plasmalemma over the acrosome, the Sertoli cell membrane, and a zone of actin filament bundles sandwiched between the Sertoli cell membrane and a thin layer of smooth endoplasmic reticulum cisternae.

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FIGURE 3.19 Early formation of the ectoplasmic specialization. The Sertoli cell cytoplasm adjacent to an early step elongating spermatid nucleus is highly specialized to begin the formation of the ectoplasmic specialization. The closeness of Sertoli and germ cell (GC) membranes is noted.

related to paracrine interactions with germ cells [45]. Morphometric studies comparing Sertoli cell structure in W/W (which lack virtually all germ cells) and control mice showed that although the testes of mutant animals were about eight times smaller than controls, the numbers of Sertoli cells in the two groups did not differ; nevertheless, Sertoli cell volume and surface area in the mutant animals were significantly smaller. Smooth endoplasmic reticulum was also significantly reduced in the mutant animals, whereas other organelle volumes and surface areas, expressed per cell, did not differ significantly in W/W and control animals.

C. Lysosomes and Multivesicular Bodies The Sertoli cell is phagocytotic and helps to remove spermatid cytoplasm through the tubulobulbar complexes and by phagocytosis of the residual bodies and degenerating germ cells. Such activities lead to the formation of lysosomes and multivesicular bodies that are located throughout the cytoplasm (Fig. 3.4), but particularly near ectoplasmic specializations and residual bodies (Fig. 3.20). As such, their presence will change depending on the stage of spermatogenesis. As a curiosity that is related to residual bodies, Sertoli cells phagocytose daily about 0.3% of the testis mass, meaning that in 1 year the total volume phagocytosed by this cell would correspond to the weight of both testes [46].

FIGURE 3.20 The residual body (RB) contains leftover cytoplasm from the developing germ cell (GC), which is phagocytosed by the Sertoli cell. Outlines of Sertoli cell cytoplasm are noted by the black lines, indicating the extensive ramification of its cellular arms or processes as it engulfs the residual bodies. Small collections of germ cell mitochondria can be seen within the residual bodies (arrows).

Chapter 3 Structure of the Sertoli Cell

D. Golgi The Golgi apparatus is relatively small in the Sertoli cell (Fig. 3.4), and the multiple components are typically dispersed in the supranuclear region (Figs. 3.9 and 3.13). Studies have shown that this is a single network of stacked saccules [47–49]. It is interesting that condensing vesicles that are associated with secretory pathways are not reported in the Sertoli cell [13]. However, small exocytotic or endocytotic vesicles are found on the basement membrane side of this cell (Fig. 3.15).

E. Cytoskeleton Primary elements of the cytoskeletal system include microtubules, actin filaments, and intermediate filaments (vimentin). The general roles of these elements in cellular structure and function are well known and include maintaining cell shape and polarity, moving or positioning of intracellular organelles, forming of pseudopodia and other cytoplasmic extensions such as microvilli, sorting and targeting of proteins, and anchoring of organelles to the plasmalemma. The Sertoli cell appears to carry these important cellular functions even further, because these elements are involved in a number of unique seminiferous epithelial activities associated with anchoring of germ cells, translocation of germ cells through the different stages of development, and nurturing and preparing the release of mature spermatids from the epithelium into the seminiferous tubule lumen [50–57]. For a complete overview of this unique set of Sertoli cell cytoplasmic components, the reader is encouraged to study the outstanding chapter by A. Wayne Vogl et al. presented in The Sertoli Cell [58]. Two of the major Sertoli cell components depend on the cytoskeletal elements for structure and function: the ectoplasmic specialization and the tubulobulbar complex. The ectoplasmic specialization is a cell junctional complex that forms between two Sertoli cells [59] and between Sertoli and germ cells [52], although the junctions are different between the two cell types [15]. It consists of the plasma membrane of the two adjacent cells, bundles of actin filaments on the Sertoli cell cytoplasmic side sandwiched between the Sertoli cell plasma membrane and a thin cistern of smooth endoplasmic reticulum (Figs. 3.18 and 3.19). Since 1993, considerable data have been published to support the original hypothesis that the ectoplasmic specialization functions as an anchor for the developing elongate spermatids and with this anchor the Sertoli cell is able to translocate the germ cell into deep crevices and then move the cell back out toward the lumen in preparation for spermiation

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[50–53, 55, 60–76]. The in vitro work of Vogl, in particular, has clearly shown that Sertoli cell microtubules are capable of moving isolated ectoplasmic specializations in both directions, suggesting that both a kinesin-type and dynein motor proteins are associated with the junctional complex [50, 53, 55, 77]. Tubulobulbar complexes are unique plasma membrane specializations that form as thin tubular indentations of Sertoli cell cytoplasm or protrusions of the adjacent germ cell cytoplasm. The complexes have bulbous regions at the end of a distal tubule that terminates as a bristle-coated pit. These structures form between adjacent Sertoli cells and between Sertoli cells and the head of late spermatids (Fig. 3.17), just prior to spermiation [17, 19, 59, 78–82]. They are found in numerous species, but only the Sertoli cell–germ cell complexes are reported in some species. Although these structures can reach up to 4 μm in length, they are recognizable only by transmission electron microscopy [59]. Their function appears to be at least threefold: (1) to help anchor germ cells to the Sertoli cell, (2) to help eliminate spermatid excess cytoplasm, and (3) to aid in the recycling or elimination of the ectoplasmic specialization, a function that is necessary for remodeling of the Sertoli cell barrier and required before the spermatid can be released in the act of spermiation. Disruption of tubulobulbar complex formation results in abnormal development of the spermatid head and larger than normal cytoplasmic droplets at the time of sperm release [59]. Cytoskeletal elements have also been visualized as participating in other important functions of the Sertoli cell. Microtubules are a major component of the long axis of the cell body, as seen by immunostaining for αtubulin (Fig. 3.21) and easily recognized by electron microscopy [58, 83, 84]. Several of the cytoplasmic organelles are aligned and transported by the microtubule tracts, as seen by their linear appearance in the cell body (Fig. 3.16). Vimentin intermediate filaments also appear along the long axis but more in the base, and a strong band is continuous around the Sertoli cell nucleus (Fig. 3.21). The importance of microtubules and intermediate filaments in helping to maintain Sertoli cell structure is best illustrated in the studies of chemical toxicant effects on the seminiferous epithelium [44, 84–96]. For example, the fungicide carbendazim, a mild microtubule poison that binds to αtubulin [44], disrupts Sertoli cell microtubule formation and induces sloughing of the elongate spermatids in a stage-specific manner (Fig. 3.22). The elongate spermatids are sloughed by breaking off from the epithelium with attached Sertoli cell cytoplasm, yet the microtubules still stain positive in the sloughed cytoplasm, whereas in the body of the

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FIGURE 3.21 Seminiferous epithelium stained immunohistochemically for αTubulin (A, C) and vimentin

intermediate filaments (B, D). αTubulin (arrows) extends from the base of the Sertoli cell to the apex in (A) rat and (C) ram testes. Vimentin staining (arrows) is reserved for the perinuclear region of the Sertoli cell in (B) rat and (D) ram testes, with the most intense staining located along the base, but wisps of vimentin filaments reaching out to the tips of elongating spermatids. (Ram testes photographs used with permission of Dr. Karl-Heinz Wrobel, University of Regensburg, Germany.)

Sertoli cell the microtubule staining is lost. This study supports the general hypothesis that is now well accepted that microtubule nucleation in the Sertoli cell is unique in that the nucleation centers are not found in the perinuclear centrosome, but rather at the apical cell periphery [57, 95]. Microtubules grow in the apical to basal direction and appear to anchor within the perinuclear intermediate filaments [57]. Loss of the microtubules within the Sertoli cell body, as shown after carbendazim treatment, is associated with a simultaneous collapse of the intermediate filaments (Fig. 3.22), which apparently weakens the apical cytoplasm and thus allows the spermatids, whose tails are dangling into the tubule lumen, to break away from the Sertoli cell. These data provide further proof that the microtubules extend into the nuclear crevasse and interact with the intermediate filaments for maintenance of the Sertoli cell structural scaffolding.

V. THE SERTOLI (BLOOD–TESTIS) BARRIER Sertoli cell–Sertoli cell tight junctions form the blood–testis barrier, which is now referred to as the Sertoli cell barrier. Several proteins, including actin, zonula occludens 1 (ZO-1), occludin, claudin, espin, and gelsolin, have been described as components of the Sertoli cell barrier or associated with the ectoplasmic specialization, and most of them belong to three classes of tight junction integral membrane proteins known as occludins, claudins, and junctional adhesion molecules [97]. However, species variations in tight junction organization are observed [72, 98], and a precise and rigidly organized actin–tight junction relationship is not absolutely mandatory for the presence or maintenance of tight junctions [72]. Although the mechanisms are not yet known, several molecules

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Chapter 3 Structure of the Sertoli Cell

FIGURE 3.22 Effects of disrupting αtubulin in Sertoli cells. (A) αTubulin staining of a control stage VII rat seminiferous tubule shows striations extending from the basement membrane to the lumen, where step 19 spermatids are lining up for spermiation. (B) The fungicide carbendazim is a microtubule poison that causes rapid sloughing of the late elongate spermatids [44]. Note the small strands of Sertoli cell cytoplasm clinging to the sloughing heads of sperm. (C) Immunostaining for αTubulin following carbendazim treatment for 2 hr. All staining is lost in the body of Sertoli cells, but staining is found in the sloughed cells in the lumen. (D) Control tubule stained for vimentin intermediate filaments. Most staining is found perinuclear but small filament strands extend into the body of Sertoli cells. (E) Carbendazim treatment causes sloughing of late spermatids into the lumen, denuding the apical cytoplasm of Sertoli cells. (F) Immunostaining for vimentin following carbendazim treatment for 2 hr. All staining is lost in the body of Sertoli cells, but staining is found perinuclear where vimentin has collapsed following treatment.

such as transforming growth factor 3 (TGF3), occludin, protein kinase A, protein kinase C, and signaling pathway (TGF3/p38 mitogen-activated protein kinase) have been shown to regulate Sertoli cell tight junctions [74, 99–102]. Also, recent in vitro studies in rats have shown that nitric oxide synthase (NOS) is an important physiological regulator of Sertoli cells tight junction dynamics, exerting its effects through the NO/soluble guanylate cyclase/cGMP/protein kinase G signaling pathway [103, 104]. The extracellular matrix is also able to regulate the Sertoli cell tight junction functions through mechanisms involving the participation of tumor necrosis factor α (TNFα) and its regulatory role on collagen α3 (IV) and other proteins that maintain the homeostasis of the extracellular matrix [75].

This investigation also showed that TNFα inhibited occludin, which is known to associate with Sertoli cell tight junction barrier assembly, and induced the expression of Sertoli cell collagen type IV, gelatinase B [matrix metaloprotease 9 (MMP9)] and tissue inhibitor of metaloproteases 1, and promoted the activation of pro-MMP9 [105].

VI. MISCELLANEOUS OBSERVATIONS A. Sertoli Cell Cycle Germ cell associations in the seminiferous tubules in mammals are organized in cyclical patterns of

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renewal and development; therefore, several studies were developed that were aimed at correlating these cyclic changes with morphological alterations in Sertoli cells during spermatogenesis in rats. In this regard, Ye et al. [26] investigated the presence of cyclic differences in volumes and surface areas of several Sertoli cell parameters using a sampling technique at the electron microscope level that sampled Sertoli cells proportionally within the seminiferous tubule. Significant differences were found among different stages of the cycle for the surface area of the cell, the volume of lipid, and the volume and surface area of the rough endoplasmic reticulum (RER). The volume and surface area of RER peaked at midcycle, and lower values were observed near the end of the cycle, showing an approximately 15-fold difference between the minimum and maximum values. Apparently, this variation found for RER parameters correlated with known patterns of protein secretion within the epithelium and with the secretion of specific proteins and important factors that control protein secretion. Another morphometric study performed at different stages of the cycle correlated Sertoli cell surface relationships and the changing volumes of developing germ cells in adult rats [22]. As a reflection of germ cell development, cyclic variation was noted for the Sertoli cell surface area that faces the basal compartment germ cells, but not the basal lamina. No cyclic variation was observed in the amount of Sertoli cell contact with each other at the level of the Sertoli cell barrier. However, when the adluminal compartment area was studied, significantly less Sertoli cell–Sertoli cell contact was seen in some stages of the cycle. Surface contact of germ cells with Sertoli cells (Figs. 3.2 and 3.20) increased progressively as germ cells entered the intermediate compartment and progressed in spermatogenesis. Also, the area in which Sertoli ectoplasmic specializations faced germ cells changed dramatically during spermatogenesis, reaching its maximum in elongating spermatids (Figs. 3.1 and 3.18).

B. Sertoli Cell Aging Compared to other testicular cells, the Sertoli cell is relatively resistant to chemical insults. However, the cell is sensitive to certain chemical exposures (see Chapter 20) and to abnormal conditions such as the formation of tumors and specific mutations, which can result in morphological changes in the nucleus and cytoplasmic organelles [106–110]. Aging is another factor that appears to be detrimental to the Sertoli cell [28, 110–114]. In addition to aging changes in the shape

of the Sertoli cell nucleus, alterations occur in the endoplasmic reticulum, lysosomes, specific Sertoli cell molecular markers [113], and in the Sertoli cell barrier [114]. A study by the Robaire laboratory [114] clearly demonstrated that with aging in the Brown Norway rat there is a complete breakdown in the Sertoli cell barrier (the blood–testis barrier), because lanthanum nitrate was seen penetrating the basal and adluminal compartments and entering the seminiferous tubule lumen.

C. Other Cytoplasmic Inclusions Other less obvious Sertoli cell components are observed, such as lipid droplets (Fig. 3.4). It is interesting that lipid droplets were first reported in this cell by the original discover of the cell [23], and the droplets are typically found in the basal compartment of all Sertoli cells, but the amount varies considerably between stages of spermatogenesis, as well as between species [13]. Lipids were originally the focus of considerable speculation in the 1800s [1], as well as during more modern times [115, 116]. More recent studies showing a significant increase in lipids in the postspermiation stages, following cimetidine treatment [117], have supported the hypothesis that Sertoli cell lipids are evidence of an ability to recycle lipids from the breakdown of germ cell degeneration and from residual body phagocytosis [13]. However, others have shown that the Sertoli cell’s abilities of uptake and conversion of fatty acids in vitro are nearly equal to those of the Leydig cell, but the effects of diabetes are less severe in the Sertoli cell [118]. Thus, the question of lipid function and metabolism in this cell type remains open. Glycogen particles are also found in the Sertoli cell cytoplasm (Fig. 3.4) and glycogen metabolism is reported for this cell [119, 120]. Glycogen recognition requires histochemistry for light microscopy [121, 122] and special fixation, such as ferrocyanide-osmium [123], is required for optimal recognition by electron microscopy. The amount of glycogen present is stage and species dependent [124, 125], with the dog Sertoli cell containing an abundance [13]. Cryptorchidism causes a decrease in the amount of glycogen in Sertoli cells [126]. In men having the Sertoli cell–only syndrome, an increased concentration of glycogen and intermediate filaments in certain Sertoli cells results in these cells staining darker [127]. An accumulation of glycogen within the cell cytoplasm is associated with the formation of Sertoli cell tumors and other types of malignancies [128–131].

Chapter 3 Structure of the Sertoli Cell

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D. Other Sertoli Cell Membrane Modifications In addition to the unusual membrane adaptations seen between Sertoli cells and the heads of elongated spermatids, there are small protrusions of the Sertoli cell cytoplasm into the cytoplasm of the developing germ cells (Fig. 3.23). In some species, such as the water buffalo, adjacent plasmalemmas of two Sertoli cells can be rolled like the lamina of a myelin sheath (Fig. 3.24).

E. Seminiferous Tubule/Rete Testis Transitional Region

FIGURE 3.23 The Sertoli cell (SC) cytoplasm becomes very thin as it expands as a sheet between two germ cells (GC), as outlined by the black lines. In a small region of the spermatid, the Sertoli cell’s cytoplasm protrudes into the germ cell.

The region that connects rete testis to the seminiferous tubule is known by several names, including tubulus rectus, intermediate region, straight tubules, terminal segment, and transitional zone [13, 132–137]. The region is lined by Sertoli cells with few spermatogonia. The transition from Sertoli cells to rete testis epithelium is abrupt (Fig. 3.25). In all species studied to date, Sertoli cells lining this junction have an appearance that differs from that of the typical adult differentiated cell. There are differences in the concentration and organization of cytoplasmic organelles, height of the cells, and appearances of the Sertoli cell nuclei. The nuclei appear more similar to those seen in the developing testis (compare Figs. 3.6 and 3.25). The Sertoli cell cytoplasm forms stringy processes that

FIGURE 3.24 Sertoli cells from water buffalo testis showing the rolling of adjacent Sertoli cell cytoplasm (arrows) into a myelin-like formation. The space between the Sertoli cells is a thin tight junction (arrows), compared to the wider intercellular space between Sertoli and germ cells (GC; arrowheads). (From Pawar et al. [38], Fig. 3.8, p. 47.)

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FIGURE 3.25 (A) The Sertoli cell–rete testis junction in the rat. The transition is rapid and the cuboidal rete testis epithelial cells cannot be mistaken for the taller Sertoli cells that extend into the lumen and appear to be stacked on one another and usually do not surround germ cells. Sertoli cell nuclei at this junction (b) appear similar to those found in the developing testis (Fig. 3.6) and stain darker than those found within seminiferous tubules that are active in spermatogenesis (a). (B) The Sertoli cell–rete testis junction in the bull. The Sertoli cell cytoplasm extends into the lumen forming a plug-like valve. (Used with permission of Dr. Karl-Heinz Wrobel, University of Regensburg, Germany.)

appear to form a “plug” or “valve” in the tubule lumen and it is generally accepted that this junction plays a role in regulating the flow of fluid from the seminiferous tubule into the rete testis, a concept suggested in 1902 [138].

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rat as a model for man. Novartis Found. Symp. 242, 82–95; discussion 95–97. Syed, V., and Hecht, N. B. (2001). Selective loss of Sertoli cell and germ cell function leads to a disruption in Sertoli cell–germ cell communication during aging in the Brown Norway rat. Biol. Reprod. 64, 107–112. Levy, S., Serre, V., Hermo, L., and Robaire, B. (1999). The effects of aging on the seminiferous epithelium and the blood–testis barrier of the Brown Norway rat. J. Androl. 20, 356–365. Lacy, D. (1962). Certain aspects of testis structure. Br. Med. Bull. 18, 205–208. Lacy, D. (1960). Light and electron microscopy and its use in the study of factors influincing spermatogenesis in the rat. J. Roy. Microsc. Soc. 79, 209–225. Sasso-Cerri, E., Giovanoni, M., Hayashi, H., and Miraglia, S. M. (2001). Morphological alterations and intratubular lipid inclusions as indicative of spermatogenic damage in cimetidinetreated rats. Arch. Androl. 46, 5–13. Hurtado de Catalfo, G. E., and De Gomez Dumm, I. N. (1998). Lipid dismetabolism in Leydig and Sertoli cells isolated from streptozotocin-diabetic rats. Int. J. Biochem. Cell Biol. 30, 1001–1010. Slaughter, G. R., and Means, A. R. (1983). Follicle-stimulating hormone activation of glycogen phosphorylase in the Sertoli cell-enriched rat testis. Endocrinology 113, 1476–1485. Guo, T. B., Chan, K. C., Hakovirta, H., Xiao, Y., Toppari, J., Mitchell, A. P., and Salameh, W. A. (2003). Evidence for a role of glycogen synthase kinase-3beta in rodent spermatogenesis. J. Androl. 24, 332–342. Montagna, W., and Hamilton, J. B. (1952). Histological studies of the human testis. II. The distribution of glycogen and other HIO4-Xchiff reactive substances. Anat. Rec. 112, 237–249. Nicander, L. A. (1957). Histochemical study on glycogen in the testes of domestic and laboratory animals, with special reference to variations during the spermatogenic cycle. Acta. Neerl. Morph. 1, 233–240. Russell, L. D., and Burguet, S. (1977). Ultrastructure of Leydig cells as revealed by secondary tissue treatment with a ferrocyanide-osmium mixture. Tissue Cell 9, 751–766. Fouquet, J. P. (1968). Infrastructural study of the glycogen cycle in the Sertoli cells of the hamster. C. R. Acad. Sci. Hebd. Seances Acad. Sci. D. 267, 545–548. Erkan, M., and Sousa, M. (2002). Fine structural study of the spermatogenic cycle in Pitar rudis and Chamelea gallina (Mollusca, Bivalvia, Veneridae). Tissue Cell. 34, 262–272. Gotoh, M., Miyake, K., and Mitsuya, H. (1987). A study of cryptorchidism. III. The histochemistry of complex carbohydrates in the testes of cryptorchid patients. Hinyokika Kiyo. 33, 905–914. Tedde, G., Montella, A., Fiocca, D., and Delrio, A. N. (1993). The sertolian epithelium in the testis of men affected by “Sertoli-cell-only syndrome.” Ital. J. Anat. Embryol. 98, 105–117. Henley, J. D., Young, R. H., and Ulbright, T. M. (2002). Malignant Sertoli cell tumors of the testis: A study of 13 examples of a neoplasm frequently misinterpreted as seminoma. Am. J. Surg. Pathol. 26, 541–550. Lee, P. J. (2002). Glycogen storage disease type I: Pathophysiology of liver adenomas. Eur. J. Pediatr. 161, S46–S49. Gurbuz, Y., and Ozkara, S. K. (2003). Clear cell carcinoma of the breast with solid papillary pattern: A case report with immunohistochemical profile. J. Clin. Pathol. 56, 552–554. Kolligs, F. T., Bommer, G., and Goke, B. (2002). Wnt/betacatenin/tcf signaling: A critical pathway in gastrointestinal tumorigenesis. Digestion. 66, 131–144.

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132. Roosen-Runge, E. C. (1961). The rete testis in the albino rat: Its structure, development and morphological significance. Acta. Anat. (Basel) 45, 1–30. 133. Nykanen, M. (1979). Fine structure of the transitional zone of the rat seminiferous tubule. Cell Tissue Res. 198, 441–454. 134. Lindner, S. G. (1982). On the morphology of the transitional zone of the seminiferous tubule and the rete testis in man. Andrologia 14, 352–362. 135. Ezeasor, D. N. (1986). Ultrastructural observations on the terminal segment epithelium of the seminiferous tubule of West African dwarf goats. J. Anat. 144, 167–179. 136. Wrobel, K. H., Schilling, E., and Zwack, M. (1986). Postnatal development of the connexion between tubulus seminiferous and tubulus rectus in the bovine testis. Cell Tissue Res. 246, 387–400.

137. Wrobel, K. H., Sinowatz, F., and Mademann, R. (1982). The fine structure of the terminal segment of the bovine seminiferous tubule. Cell Tissue Res. 225, 29–44. 138. Spangaro, S. (1902). Uber die Histologischen Veränderungen des Hodens, Nebenhodens, Und Samenleiters von Geburt an biz zum Griesalter, mit besonderer Berucksichtgung der Hodenatrophie, des elastischen Gewebes und des Vorkommens von Krystallen im Hoden. Anat. Record 18, 593–771. 139. Wong, V., and Russell, L. D. (1983). Three-dimensional reconstruction of a rat stage V Sertoli cell. I. Methods, basic configuration, and dimensions. Am. J. Anat. 167, 143–161. 140. Fawcett, D. W. (1975). Ultrastructure and function of Sertoli cells. In “Handbook of Physiology: The Male Reproductive System” (D. W. Hamilton and R. O. Greep, eds.), pp. 21–55. American Physiology Society, Washington, DC.

P A R T

II SERTOLI CELL DEVELOPMENT

This Page Intentionally Left Blank

C H A P T E R

4 Embryonic Sertoli Cell Differentiation

I. II. III. IV.

ANDREA S. CUPP

MICHAEL K. SKINNER

Department of Animal Science, University of Nebraska, Lincoln, Nebraska

Center for Reproductive Biology, School of Molecular Biosciences, Washington State University, Pullman, Washington

INTRODUCTION OVERVIEW OF EMBRYONIC TESTIS DEVELOPMENT STAGES AND TIMING SUMMARY References

II. OVERVIEW OF EMBRYONIC TESTIS DEVELOPMENT The genital ridge is primarily composed of a single layer of coelomic epithelium from 9 to 9.5 days postcoitus (dpc) in the mouse (Fig. 4.1) [1]. At 9–10 dpc the primordial germ cells (PGCs) migrate from extragonadal sites within the yolk sac to colonize the urogenital ridge [2, 3]. The gonad is bipotential after germ cell migration and morphologically can be distinguished from the adjoining mesonephric tissue but cannot be identified as an ovary or a testis [4]. Two morphological events occur to alter the bipotential gonad at 11 dpc of development. First, Sertoli cells differentiate in part from the coelomic epithelium and start to proliferate, aggregating with the PGCs. Second, mesonephric cells (endothelial or preperitubular in origin) migrate from the mesonephros and enclose the pre–Sertoli-PGC aggregates to form seminiferous cords (Fig. 4.1) [5]. Both of these events rely on expression of Sry by the developing Sertoli cell [6]. As the Sertoli cell differentiates (11–13 dpc) it acquires the ability to produce Müllerian inhibiting substance (MIS), which inhibits Müllerian duct growth. The Müllerian duct is the precursor female reproductive tract that differentiates into the cervix, uterus, oviduct, and portions of the anterior vagina [7, 8]. After seminiferous cord formation, cells within the interstitium differentiate to form immature Leydig cells, while the preperitubular cells further differentiate to form a single layer of cells enclosing each seminiferous cord. In the rat, steroidogenesis occurs within

I. INTRODUCTION In the past 10 years, we have experienced a veritable explosion in the amount of information that has been uncovered on genes important in the regulation of testis differentiation. With the identification of the testis determining gene, Sry, and localization of its expression in the Sertoli cell, new information has been elucidated on factors regulating the male sex differentiation pathway. The Sertoli cell is the critical cell type that initiates development of testis-specific gene expression, induces testis morphology, and establishes crucial parameters for spermatogenic function and capacity. Therefore, proper differentiation of the Sertoli cell during embryonic testis differentiation is mandatory for normal adult testis development and function. In this chapter, we examine embryonic testis development and the crucial role of the Sertoli cell in both sex determination and morphological events that result in the formation of a testis. The events regulating testis development are discussed using primarily the mouse as a model; however, comparisons will be made to rats, domestic livestock, and humans where information is available. SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

43

Copyright 2005, Elsevier Science (USA). All rights reserved.

44

Andrea S. Cupp and Michael K. Skinner Developing Testis Stage I: Urogenital Ridge Mouse

9 dpc

Stage II: Indifferent / Bipotential Gonad 10-10.5 dpc

Stage III: Sex Determination 10.5-11 dpc

Stage IV: Cord Formation 11-12 dpc

Stage V: Development of Functional Testis 12-13 dpc 12

Developing Gonad Mesonephros Genital Ridge

Germ cell Colonization Sertoli cell Differentiation

Sry Expression Sertoli Cell Migration Sertoli/Germ Cell Aggregation

Migration of Pre-peritubular And Endothelial cells to form seminiferous cords

Seminiferous cord formation complete Initiation of Leydig cell migration and differentiation

FIGURE 4.1 Morphological stages that occur during testis differentiation from genital ridge formation to cord formation and cell proliferation in the mouse with staging in number of tail somites (ts) and days postcoitus (dpc).

the testis at 14.5 dpc when 3β-hydroxysteroid dehydrogenase (3β-HSD) is first expressed in pre-Leydig cells [9, 10]. Testosterone produced by the Leydig cells maintains and stabilizes the Wolffian duct, which is the precursor of the male reproductive tract structures: the epididymis, vas deferens, and portions of the secondary sex glands [11]. The seminiferous cords develop lumen around puberty to become the seminiferous tubules. A major function of the Sertoli cell within the testis is to provide the appropriate environment (e.g., production of proteins and growth factors) and cytoarchitectural support for the developing germ cells [12]. The comparisons between events that occur in the mouse, rat, and pig during testis development and correlated time points in embryonic testis development are presented in Table 4.1.

III. STAGES AND TIMING There are at least five different morphological stages of testis development, which are depicted in Fig. 4.1: (1) development of a genital ridge, (2) formation of an indifferent or bipotential gonad, (3) sex determination, (4) induction of testicular cords in the testis, and (5) development of a functional testis. These stages are represented in Fig. 4.1 by timelines in the mouse and are compared to other species (human, pig, cattle, sheep, and rats) in Table 4.1.

The first two stages, genital ridge formation and formation of an indifferent gonad, occur whether the individual has XX or XY chromosomes and, thus, is independent of testis or ovarian development. The last three stages of gonadal development are dependent on genes that are expressed within the indifferent gonad to result in either a testis or ovary. Expression of the gene Sry (sex-determining region of the Y chromosome) by the Sertoli cell directs the indifferent gonad to become a testis [13–15], whereas the absence of Sry and expression of Wnt4 appear to regulate formation of an ovary [16]. Thus, formation of testis-specific morphogenic structures such as seminiferous cords is dependent on differentiation of the Sertoli cell, expression of Sertoli cell–specific genes, and proliferation of the Sertoli cell population to result in normal testis development (reviews [1, 17]).

A. Stage I: Genital Ridge Development The initial step in the development of gonads is the formation of the genital ridge (Fig. 4.1) and urogenital system from the intermediate mesoderm; this step begins at 9–9.5 dpc in the mouse. The Wolffian duct, which is the precursor of the male reproductive tract system, is derived from lateral mesoderm and runs the length of the urogenital system and develops from the mesonephric duct [18]. The Müllerian duct, which is the precursor of the female reproductive system, appears between 11.5 and 12.5 dpc from invaginations

45

Chapter 4 Embryonic Sertoli Cell Differentiation TABLE 4.1 Gestational Age at Each Stage of Testis Development in Humans, Pig, Cattle, Mice, Rats, and Sheep Species

Genital ridge

Bipotential gonad

Testis cord formation

Reference

Human Gestational age

5 weeks

6 weeks

7–8 weeks

Sinisi et al., 2003

Gestational age Crown rump length

18–20 days 8–10 mm

21 days 10–12 mm

26 days 15 mm

McCoard et al., 2001

Cattle Gestational age Crown rump length

27–31 days <11 mm

32–39 days 11–18 mm

40 days >19 mm

Wrobel and Sub, 1998

Mouse Gestational age Total no. of somites No. of tail somites

9–10 dpc 13–28 s <9 s

10–11.5 dpc 29–48 s 9–18 s

12.5 dpc 49–52 s 24–30 s

Karl and Capel, 1998 Rugh, 1968

Rat Gestational age

10–11 dpc

11.5–12.5 dpc

13.5–14 dpc

Magre and Jost, 1984; Magre et al., 1980; Magre et al., 1981

Sheep Gestational age

20

23 dpc

30–35 dpc

Pellinieme et al., 1981; Payen et al., 1996; Quirke et al., 2001; Mauleon, 1961; McNatty et al., 1995

Pig

of surface epithelium from the mesonephros running parallel to the Wolffian duct [1, 17]. Only one of the two ductal systems, Müllerian or Wolffian, will develop further in mammals. The duct that develops is totally dependent on gonadal differentiation (whether an ovary or testis develops) and expression of factors that will support or regress the female or male ductal structures [11, 19]. The Sertoli cell of the testis will produce MIS, which will cause regression of the precursor female reproductive structures, whereas Leydig cells, which differentiate much later in testis development, will produce testosterone, allowing for maintenance of the male reproductive tract structures (Wolffian duct [11]). Histologically, the formation of the genital ridge is the formation of a pair of mounds on either side of the dorsal mesentery of the hind gut at 9 dpc in the mouse. These “mounds” are formed by a single layer of coelomic epithelial cells that almost immediately begin to proliferate and thicken (Fig. 4.1) [4]. In the pig, the genital ridge forms around embryonic day 20–21 of the approximate 114-day gestation and, like the rodent, consists of a single layer of coelomic epithelium that presumably migrates from the dorsal mesentery to line the mesenchymal layer forming the genital ridge [20, 21] (Table 4.1).

indifferent gonads. Lhx1 (Lim 1), Wilms’ tumor 1 (Wt1), steroidogenic factor 1 (Sf1), Emx2, and Lhx9 are all expressed in the urogenital ridge by 9.5 dpc (Fig. 4.2). Mice with disruptions in any of these genes have arrested gonadal development, and the mice die shortly after birth from developmental defects in the adrenal cortex (Sf1), kidneys (Wt1, Emx2), or kidneys and brain (Lhx1) [22–27]. Of these genes, the null mutants of Sf1, Wt1, and Emx2 all have similar phenotypes because the gonad starts to form and then it regresses. Thus, these genes are thought to be critical in expansion of the initial genital ridge. Lim1 or Lhx1 is the gene that has the least defined function of all the genes expressed at this time in

1. Gene Expression Resulting in Proliferation of the Genital Ridge and Indifferent Gonad Formation

9 dpc

FIGURE 4.2 Expression of Sertoli cell genes during mouse testis

Several genes encoding transcription factors mediate the early events in the development of the

development. References for each gene are as follows: Lhx1: [22], Sf1: [1, 141], Emx2: [25], Lhx9: [27], Wt1: [1, 30, 35], Sox9: [1, 141], DAX-1: [1, 94, 95, 141], Dmrt1: [89], Sry: [1, 141], GATA-4: [141], and MIS [141].

? ?

10 dpc

11 dpc

12 dpc

13 dpc

Lhx1 Sf-1 Emx2 Lhx9 W-t Sox9 DAX-1 Dmrt1 Sry GATA-4 MIS 14 dpc

46

Andrea S. Cupp and Michael K. Skinner

gonadal development. Lim1 is a homeodomain protein that is expressed throughout the mesonephric ductal system and its expression is detected by 9 dpc (Fig. 4.2). Lim1 mutants that survive do not have kidneys or gonads; however, the exact stage of gonadal disruption has not yet been determined [22]. The Wilms’ tumor suppressor gene Wt1 was initially identified as a gene inactivated in a subset of Wilms’ tumors, a form of pediatric kidney cancer [28, 29]. Inactivation of both Wt1 alleles results in embryonic lethality and a failure of kidney and gonadal development [23]. Wt1 expression occurs in the genital ridge of the indifferent gonad around 9.5 dpc (Fig. 4.2) and then becomes localized to the Sertoli cells of the testis and granulosa cells of the ovary [30, 31]. An initial function of the Wt1 gene is in the formation of a subset of mesonephric tubules, but its role in early gonadal development is not known [32]. In Wt1 mutant embryos, a thickening of the coelomic epithelium occurs, but there is no further development of the gonad and by 14.5 dpc the gonad is completely regressed [23]. Thus, a potential role for Wt1 in the gonad is in proliferation and thickening of the coelomic epithelium within the genital ridge to become the indifferent gonad. In humans, a number of additional diseases were shown to be associated with Wt1 mutations that primarily resulted in urogenital abnormalities [33]: WAGR (Wilms’ tumor, aniridia, genitourinary abnormalities, mental retardation), Denys-Drash (DDS), and the Frasier syndrome. The major abnormalities associated with these diseases are varied and can range from cryptorchidism and hypospadia to streak gonads and sex reversal of internal and external genitalia [34–38]. The absence of expression of Wt1 does not affect Sox9 or Lhx9 expression; however, both DAX-1 and Sf1 were determined to be absent in Wt1 mutant mice when compared to wildtype controls [39]. Furthermore, Wt1 also has been determined to activate the endogenous promoter of DAX-1 [40]. Therefore, both Sf1 and DAX-1 expression may be regulated directly or indirectly by Wt1. The Sf1 gene encodes an orphan nuclear receptor that regulates the expression of several genes involved in gonadal development and steroidogenesis [41]. In the mouse, targeted deletions of both Sf1 alleles result in the absence of both the adrenal gland and the gonad, supporting a role for Sf1 as an essential regulator of the first endocrine cells in the gonad [24]. In the early gonad, Sf1 mRNA can be detected at 9–9.5 dpc (Fig. 4.2) and is localized to the coelomic epithelium and the mesenchymal cells of the mesonephros between the tubules. The gonads of Sf1 mutant embryos do not develop beyond the early indifferent stage and XY Sf1 mutant embryos display sex reversal [24]: The Müllerian ducts develop into uteri, oviducts, and upper vagina.

Sf1 disrupted mice lack both adrenal glands and gonads supporting the suggestion that Sf1 is an essential regulator of the first endocrine cells within the gonad. Ftz-fl gene disrupted mice (encodes both Sf1 and the alternative transcript ELP) are normal at 10.5 dpc; however, at 11.5 to 12 dpc a number of the cells on the gonads degenerate via apoptosis or programmed cell death. Wt1 expression is normal in both Sf1 and Ftz-fl mutant embryos, which supports previous research from Wt1 mutants with no Sf1 expression that Wt1 is critical in inducing expression of Sf1. In Lhx9 mutants only minimal expression of Sf1 was detected [27], but normal expression of Wt1 occurred. These results suggest that Wt1 is necessary but not sufficient to activate Sf1 expression. Recent research has also determined that Lhx9 can bind to the Sf1 promoter and induce expression of Sf1 [39]. It appears that Wt1 and Lhx9 bind to Sf1 and have an additive effect on induction of expression of Sf1 in the early gonad [39]. Expression of Lhx9 occurs in the genital ridges of mice around 9.5 dpc (Fig. 4.2) and by 11.5 it is expressed at high levels in the cells of the coelomic epithelium, whereas more moderate levels are expressed in the adjacent mesenchyme (future tunica albuginea [27]). At E13.5 (after morphological testis development), expression of Lhx9 is in the interstitial mesenchyme and less within the testicular cords. Thus, as morphological testis differentiation is completed, the expression of Lhx9 is downregulated in the cells of the coelomic epithelium. At 11.5 dpc there is no difference in the gonadal structures between Lhx9 gene–deficient mice and wild-type mice. However, by 12 dpc no proliferation was evident in the Lhx9 mutants and by 13.5 dpc no discrete gonads had formed in the Lhx9deficient embryos as seen in the wild-type mice. Lhx9deficient embryos did not have the level of cellular proliferation or apoptosis within cells of the genital ridge as was present in the wild-type embryos, suggesting that Lhx9 may be necessary for the proliferation of gonadal cells, especially cells that are proliferating within the coelomic epithelium during transition of the genital ridge into the indifferent gonad [27]. In mice lacking Lhx9, germ cells migrate normally, but somatic cells of the genital ridge fail to proliferate so a discrete gonad does not form. Also, in Lhx9deficient mice (as stated previously) there is reduced expression of Sf1 that is essential to the production of both adrenal and gonads within mice [27]. This means that Lhx9 may lie upstream of Sf1 in the developmental cascade of events that occur during testis development. Emx2 is a mouse homologue of the Drosophila head gap gene ems [42, 43]. Emx2 mutant mice die due to failure of the urogenital system to develop.

Chapter 4 Embryonic Sertoli Cell Differentiation

Emx2 mutants do not have kidneys, ureters, gonads, and genital tracts; however, the adrenal glands and bladder develop normally. Expression of Emx2 was found in the genital ridge next to the Wolffian duct at 9.5 dpc (Fig. 4.2). Wolffian duct development in 10.5dpc mutant embryos was normal and the mesonephric tubules adjacent to the Wolffian duct were also similar to wild-type. At 11.5 dpc degeneration of the Wolffian duct was observed in several sites and the normal thickening of the coelomic epithelium did not occur in mutants [25]. There was no change in expression of Wt1 in Emx2 mutants, demonstrating that Emx2 expression was not critical for Wt1 expression. However, like Wt1 deficient mutants, the coelomic epithelium of Emx2 mutants did not proliferate and many cells throughout the gonad went through a rapid apoptosis [25]. Therefore, Emx2 may be important in proliferation of the coelomic epithelium, differentiation of cells from the coelomic epithelium, or survival of gonadal cells that are derived from this cell origin.

B. Stage II: Development of Bipotential Gonad The second stage of gonad formation is development of the bipotential or indifferent gonad (Fig. 4.1). As discussed previously the indifferent or bipotential gonad arises through a thickening or proliferation of cells on the ventromedial surface of the mesonephros directed presumably by genes that are upregulated around 9 to 9.5 dpc. The indifferent gonad becomes visible around 10 dpc in the mouse (other species time points are given in Table 4.1) and can be distinguished from the mesonephros but cannot be identified as a testis or ovary [4]. The indifferent gonad is unique because it has the capacity to differentiate into either the testis or the ovary, depending on transcription factors expressed within the differentiating somatic cells of the coelomic epithelium. Commitment of the indifferent gonad to either the testis or the ovary occurs after migration of primordial germ cells from extragonadal origins to the gonadal ridge. Therefore, the indifferent gonad is composed of primordial germ cells and a thickened layer of coelomic epithelium prior to sex determination. 1. Primordial Germ Cell Migration Primordial germ cells are the progenitors of male and female gametes. In mammals, PGCs are of extragonadal origin [2, 44]. In the mouse embryo, PGCs are thought to be derived from a small population of epiblast cells (possibly 40–50 cells) set aside in the extraembryonic mesoderm at the mid-primitive-streak

47

stage (7–7.5 dpc) [45]. PGCs migrate from the epiblast through the posterior primitive streak to the allantoic stalk and then to the dorsal mesentery via the hindgut to colonize the urogenital ridges [46–50] in the mouse around 9–11 dpc (Fig. 4.1 and Table 4.1). The migration of PGCs is composed of four distinct phases. In the first phase, PGCs are passively carried from the yolk sac into the hindgut as a result of embryonic folding. During the second phase, the PGCs leave the hindgut and enter the dorsal mesentery. From the dorsal mesentery, the PGCs migrate by amoeboid movement between the mesenchymal cells of the dorsal mesentery toward the genital ridges to complete the third phase. The fourth phase is the colonization phase, in which the PGCs arrive at and populate the genital ridges [51]. As PGCs migrate they proliferate rapidly and their number increases to around 25,000 at 13.5 dpc [47] in the gonads. 2. Contribution of Extracellular Matrix Proteins to PGC Migration The migration of the PGCs was initially thought to be primarily through amoeboid movement [52] in many different species [51, 53–62] from the rat and mouse to humans. However, current evidence suggests that extracellular matrix components may be responsible for defining the pathway and pattern of PGC migration. The migrating PGCs can be morphologically characterized by their large, heterochromatic nuclei, the presence of filiform and lobate pseudopodia, microfilaments, and polarized cytoplasm and by expression of alkaline phosphatase [63]. As they migrate PGCs are frequently connected to each other by cytoplasmic threads [64], which may aid in transporting them along the migration pathway. Furthermore, transient interactions between fibronectin molecules on the extracellular matrix and corresponding receptors on the PGCs may also induce germ cell movement [65–67]. Laminin, type IV collagen, chondroitin sulfate [68], and hyaluronan [69] also line the migratory pathway of PGCs and are expressed in the basal laminae surrounding the migration route. Therefore, many extracellular matrix proteins and specifically proteoglycans and glycosaminoglycans may interact to contribute to the migration of PGCs along their route to the genital ridges. 3. Contribution of Growth Factor Secreted Chemoattractants to PGC Migration A major method of PGC movement has been determined to be through chemoattractants secreted from the urogenital ridge or secreted by cells along the PGC

48

Andrea S. Cupp and Michael K. Skinner

migratory pathway. Kit Ligand [KL; also called stem cell factor (SCF), mast cell growth factor (MGF), and steel factor (SF)] has been demonstrated to be an important growth factor in PGC migration. Kit Ligand is the product of the murine Steel (SL) gene, and its receptor, c-kit, is the product of the murine White spotting (W) gene. By in situ hybridization analysis and histochemical studies of mouse embryos, it has been demonstrated that c-kit transcripts and protein are expressed in migrating PGCs. Also Kit Ligand mRNA expression is associated with cells present in the migratory pathways of PGCs [70–72]. Mouse embryos with mutations at either W or Sl loci show a greater than 99% decrease in the number of PGCs. The deficiency of PGC numbers in Sl/sl embryos was correlated to either a failure in proliferation or an excessive rate of cell death rather than defects in migration. In addition to stem cell factor, transforming growth factor beta 1 has also been demonstrated to induce PGC migration to the genital ridge [73].

makeup, germ cells within the gonad will enter meiosis by 13.5 dpc if there is no interference by Sertoli cell secreted factors. Thus, in the female, germ cells will enter meiosis and the gonad will follow normal ovarian differentiation. The entrance into meiosis by the germ cells may alter the ability of the gonad to form a testis. There is recent evidence that meiotic germ cells may produce factors that inhibit the mesonephric cell migration necessary for formation of seminiferous cords. At 14.5 dpc mesonephric cell migration could not be induced into normal XX gonads, but cells did migrate into germ cell–depleted XX gonads [79]. Even though germ cells are not necessary for induction of testicular cords, they can inhibit migration of cells into the gonad after they have entered meiosis. Therefore, Sertoli cell–germ cell interactions are important during testis morphogenesis, and further elucidation of their interactions is necessary to understand mechanisms regulating abnormal gonadal morphogenesis (e.g., ovotestis formation).

4. Germ Cell–Sertoli Cell Interactions Affecting Germ Cell Mitosis

5. Transcription Factors Expressed in the Indifferent/Bipotential Gonad That Potentially Direct Testis Differentiation

Germ cells behave very differently during male and female gonadogenesis. In both sexes, the germ cells migrate to the coelomic epithelium of the gonad between 9.5 and 11 dpc, and they proliferate there until 12.5 dpc (Table 4.1). At this stage in the male, the germ cells are enclosed within the testis cords and enter mitotic arrest. The germ cells in females continue to undergo mitosis until 13.5 dpc when they enter meiotic arrest [74]. The interaction between germ cells and Sertoli cells is thought to play a critical part in sex determination. The Sertoli cell is hypothesized to produce factors that prevent male germ cells from entering meiosis. Initially, it was proposed that enclosure of germ cells within the XY gonads during cord formation inhibited meiosis because PGCs enter meiosis I when they develop in the adrenal gland [75] or when cultured in vivo [74]. When seminiferous cord formation was disrupted by cyclopamine in testis organ cultures [76] or in Desert hedgehog (Dhh) null mice XY gonads on a mixed genetic background [77], it was determined that enclosure of the PGCs in cords was not essential for inhibition of germ cell meiosis. Instead it was concluded that inhibition of meiosis was dependent on a diffusible factor originating from the Sertoli cell. One such factor, prostaglandin D2, has been reported to be produced by Sertoli and germ cells to function in Sertoli cell differentiation and potentially in inhibiting germ cells’ entrance into meiosis. This factor, prostaglandin D2, has also been demonstrated to masculinize female urogenital ridges [78]. Regardless of their chromosomal

Several genes are expressed around 10.5 dpc in the indifferent gonad that appear to be important in subsequent sex determination and gonadal morphogenesis. The four most critical genes to testis development that are expressed at this time are Sry, DAX-1, Dmrt1, and Wnt4 (Fig. 4.2). Other genes that are potentially downstream of Sry, DAX-1, Dmrt1, and Wnt4 are discussed later. Sry was identified as the potential testis determining factor when its translocation from the Y chromosome caused sex reversal [80]. Sry is a member of a transcription factor superfamily that contains a DNAbinding domain call the high-mobility group (HMG) box. HMG box-containing proteins bind and bend DNA acting as architectural transcription factors. The HMG box is the only region within Sry that is conserved among species, and in humans sex-reversing mutations cluster around the HMG box region of Sry [81]. Therefore, it is thought that the HMG region of Sry is critical for testis determination [80, 82, 83]. In rodents, Sry expression is tightly regulated during sex determination [84, 85], whereas in other species Sry may be expressed in many tissues throughout adult life [86, 87]. (Initial Sry expression for different species is presented in Table 4.2.) Sry expression is potentially regulated by Wt1. Wt1, as discussed previously, is a transcription factor containing four contiguous C2H2–type zinc-finger motifs that act as DNA–RNA binding or protein–protein interaction domains. Recent research has determined

Chapter 4 Embryonic Sertoli Cell Differentiation

49

TABLE 4.2 Initial Embryonic Testis SRY Expression Species

SRY/Sry

Cord formation

Human

7 weeks gestation

7–8 weeks

Hanley et al., 2000; Sinisi et al., 2003

Mouse

10.5 dpc

11.5 dpc

Koopman et al., 1990; Hacker et al., 1995

Rat

12 dpc

13.5 dpc

Berta et al., 1990; Gubbay et al., 1990; Jager et al., 1990; Jost et al., 1981

Pig

E21–20 dpc

26 dpc

Daneau et al., 1984; Parma et al., 1999; Pelliniemi et al., 1981

Sheep

23 dpc

30 dpc

Payen et al., 1996

that Wt1 binds to and acts synergistically with SRY to activate transcription. It is believed that the Sry–Wt1 interaction is mediated by the Wt1 zinc-finger domain and the Sry HMG box [88]. Dmrt1 is expressed around 10.5 dpc in the bipotential gonads of both sexes (Fig. 4.2). After 12.5 dpc transcripts of Dmrt1 are localized to Sertoli and germ cells and its expression is sexually dimorphic at this time, having very weak expression in the ovary after sex determination [89]. Dmrt1 is the best-conserved sexdetermining gene between phyla because c-elegans (mab-3) and Drosophila (dsx) have homologous genes that are involved in sex determination [90]. Although the sex-specific expression of Dmrt1 has been established, the role in sex determination has not been determined. Humans carrying deletions in the region of the chromosome containing the Dmrt1 gene are XY sex-reversed [91]; however, mice carrying homozygous deletions of Dmrt1 are not XY sexreversed but are infertile due to defects in germ cell proliferation and testis development. Dmrt1 does not appear to be important in ovarian development because Dmrt1 mutant females are fertile [92]. The different phenotypes in the human and mouse models may be due to additional gene deletions occurring on chromosome 9 [93]. Currently there are seven different DM genes, termed Dmrt1–7 and several other DM genes; Dmrt3 and Dmrt2 are located adjacent to Dmrt1 on chromosome 9. Deletion of several of these genes in the human model may explain the sex-reversal phenotype, which does not appear to occur within the Dmrt1 mutant mice. Recent expression data have demonstrated that of the seven DM genes three are expressed in the mouse gonad, with Dmrt4 being expressed at similar levels in the testis and ovary. Dmrt3 is more highly expressed in the testis, and Dmrt7 is expressed at higher levels in the ovary [93]. Several Dmrt genes may be involved in gonadal sex differentiation and may be able to partially compensate for other DM gene functions. DAX-1 (Dosage-sensitive sex-reversal-adrenal hypoplasia congenital-critical region of the X chromosome, gene 1) is expressed around 10.5 dpc in the mouse

Reference

indifferent gonad (Fig. 4.2). Its expression is upregulated in both sexes from 11.5 dpc and is downregulated in males at 12.5 dpc, but remains in females throughout ovarian development [94, 95]. It was originally proposed that DAX-1 was not necessary for normal testis differentiation. However, recent research with Nr0b1 (DAX-1)-deleted mice crossed to Mus domesticus poschiavinus mice (a strain known to be susceptible to XY sex reversal because of an altered Sry allele) demonstrated that DAX-1 is necessary (with normal Sry expression) for testis differentiation [96]. DAX-1 appears to be a dosage-sensitive gene because XY individuals with duplications in DAX-1 show male-to-female sex reversal [97]. In the indifferent gonad, DAX-1 is expressed in the somatic cells, but after sex determination its expression is restricted to the Leydig cells of the testis and thecal cells of the ovary during late embryonic development and adulthood [94, 98]. DAX-1 is proposed to be an antagonist of Sry function in males because both genes are expressed in the same tissues at a similar time period [95]. Coexpression of DAX-1 and Sry in XX mice results in females, whereas XX mice carrying only the Sry transgene develop as males [95]. An early increase in DAX-1 expression is also thought to cause female development in individuals who have Sry expression. The timing of expression of these two genes appears to have dramatic effects on gonadal morphogenesis. Another function of DAX-1 appears to be in regulation of Sf1. Most in vitro studies support a role for DAX-1 in the inhibition of Sf1-mediated gene transcription [99, 100], but the exact nature of their interaction is still unclear. Both Sf1 and DAX-1 are expressed in similar tissues—adrenal gland, gonad, hypothalamus, and pituitary—so it is possible that they may interact to regulate the development of all of these organs [1]. Wnt4 is a member of the Wnt signal transduction family and was originally identified as a mammalian homologue of the Drosophila wingless gene. The wingless family regulates paracrine interactions leading to the development of both the kidney and angiogenesis of many different organs [101–104]. Wnt4 is proposed

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to antagonize male development in a dosage-dependent manner and to be necessary for ovarian development. Overexpression of Wnt4 leads to upregulation of DAX-1. So Wnt4 in concert with DAX-1 may regulate female gonadal development [105]. Similar to the DAX-1 mutants, Wnt4 knockout mice exhibit similar defects in both kidney and adrenal gland development. Gonadal development and steroidogenic function are affected exclusively in the Wnt4 null females [16]. Wnt4 null females are masculinized and have Wolffian duct structures in the absence of a Müllerian duct system. They also express steroidogenic enzymes required for production of testosterone such as 3β-HSD and 17αhydroxylase. In normal XX females, Wnt4 may be expressed to suppress androgen biosynthesis so the Wolffian duct structures would not be maintained [16]. In the absence of Wnt4, androgens would be expressed and the Wolffian duct would remain. The absence of the Müllerian duct in the Wnt4 null females is harder to explain. An antagonistic interaction may exist between Wnt4 and MIS in that if Wnt4 is not present MIS can be secreted to initiate regression of the Müllerian duct. Overexpression of Wnt4 also causes disruptions in gonadal morphogenesis, but in this case it appears to be in the XY gonad. In transgenic mice overexpressing Wnt4, abnormal development of male gonadal vasculature and testosterone production occurred [106], but there was no XY sex reversal. During testis development the formation of the coelomic vessel and organization of blood vessels around the seminiferous cords is one of the earliest morphological sexspecific events. It appears that Wnt4 may affect the ability of blood vessels to develop side branches, resulting in abnormal vascular development [106] in the developing testis. In summary, Wnt4 expression and overexpression appear to antagonize testis development and testis morphogenesis, resulting in either no testis formation or abnormal testis development, respectively.

C. Stage III: Sex Determination Sex determination in the testis occurs when Sry in conjunction with other transcription factors is expressed within the pre-Sertoli cells to initiate a cascade of events resulting in the formation of a testis. Several elegant experiments using XX mice overexpressing an Sry transgene have demonstrated that Sry is required for normal testis development [107]. Sry appears to be important in at least two critical stages of testis development. The first involves differentiation of the Sertoli cell lineage [108–110] and the second induction of testicular morphogenesis, resulting in formation of testicular cords [107, 111] (Fig. 4.1).

Sry expression within the bipotential gonad regulates expression of genes that differentiate the Sertoli cell lineage from precursor somatic cells in the coelomic epithelium [10.75–11 dpc; 10–15 tail somites (ts)] [112] (Fig. 4.1). The initial differentiation of the Sertoli cell lineage from the coelomic epithelium is a critical step in testis differentiation. The Sertoli cell is the first cell of the testis to differentiate [113], and the Sertoli cells are the only cells in the testis that demonstrate a bias for the presence of a Y chromosome [114]. After differentiation, the Sertoli cells begin to proliferate (11.25 dpc; 16–18 ts), doubling in number at Sry peak expression (11.5 dpc; 18 ts) [107]. Simultaneously, the Sertoli cells move into the gonad proper, forming aggregates with primordial germ cells. Proliferation of the Sertoli cells increases the size of the testis (with respect to the ovary) and appears to be solely dependent on the expression of Sry [107, 115] or Sry-regulated genes. Differentiation and proliferation of the Sertoli cells in the indifferent gonad is required for testis development. (See Table 4.2 for initial Sry expression in other species.) 1. Sertoli Cell Origin The origin of PGCs has been established as cells that migrate from the base of the allantois via the gut mesentery and colonize in the gonad between 10.5 and 11.5 dpc [45, 64]. Peritubular myoid cells and endothelial cells have been demonstrated to be derived from the migrating mesonephric cells [5]. However, the origin of the Sertoli and Leydig cells has been controversial for some time. Initially, Sertoli cells were thought to be derived from mesonephric cells that migrated into the gonad during testis differentiation and/or from the coelomic epithelium [116]. Differentiated Sertoli cells have been determined to be present within the gonad by 11.5 dpc since MIS, the first known Sertoli cell–secreted hormone, was detected in media from testis cultures at 11.5 dpc [117]. The mesonephros is a mesenchymal tissue derived from the intermediate mesoderm containing the epithelial mesonephric duct and mesonephric tubules. Ultrastructural and staining similarities have been revealed through electron microscopy studies between Sertoli cells and mesonephric tubule cells at the mesonephric–gonadal junction [118–121]. Based on these similarities, it was suggested that the mesonephric tubule cells dedifferentiate from the epithelial structure at the mesonephric–gonadal junction, migrate into the gonad, and contribute to the Sertoli cell population [118–121]. However, no direct experiments have been conducted that support this theory.

Chapter 4 Embryonic Sertoli Cell Differentiation

2. Arguments Against the Sertoli Cell Being Derived from the Mesonephros Migration experiments using a lacZ transgenic mesonephros (ROSA26) [122] placed in apposition to a wild-type gonad demonstrated that mesonephric cell migration occurred into the male gonad between 11.5 and 16.5 dpc. However, in these experiments Sertoli cells never migrated during this time frame. Instead the cell types that migrated were peritubular myoid, endothelial, or endothelial-associated cells [6]. Therefore, if Sertoli cells do arise from the mesonephros, they must migrate prior to 11.5 dpc. Evidence from Pax2 null mutants would suggest that Sertoli cells do not originate from the mesonephros. In these mutants mesonephric tubules never develop into mature tubules or make contact with the cells of the precursor gonad; however, testis organogenesis proceeds normally with all representative cells present, including Sertoli cells [123].

3. Evidence That Sertoli Cells Originate from the Coelomic Epithelium The coelomic epithelium has also been implicated as a source of Sertoli cells due to its ultrastructural similarities from EM studies [60, 123, 124]. The coelomic epithelium is a single layer of cells that covers the entire coelomic cavity, including the morphological structures that will become the gonad. In the chick, experiments using India ink demonstrated that coelomic epithelial cells migrated into the gonad comprising portions of the developing testis [125]. In the rat, the basement membrane components collagen I and III were fragmented beneath the coelomic epithelium at the onset of gonadogenesis [126]. Both of these observations would suggest that cells from the coelomic epithelium move into the testis and are precursors of cell populations within the testis. In more recent experiments, the cells of the coelomic epithelium were directly labeled using a fluorescent lipophilic dye, DiI. These cells were imaged and their fate was determined between 15 and 30 ts corresponding to 11.2–12.5 dpc. The coelomic epithelial cells moved to the interior of the gonad and became Sertoli cells as well as other cell lineages. Prior to 18 ts, cells of the coelomic epithelium become pre-Sertoli cells within the seminiferous cords. After 18 ts, labeled coelomic epithelial cells no longer became Sertoli cells but were always found outside testis cords, suggesting that they become a portion of the interstitial cell population. Movement of cells from the coelomic epithelium to the interior portion of the gonad decreased around 12.5 dpc and ended as the tunica albuginea began to form in the XY gonad. Therefore, at

51

least a portion of the Sertoli cell population originates from the coelomic epithelium during early testis differentiation, and later, the cells of the coelomic epithelium become populations of cells within the interstitium [112]. 4. Sex Determination Genes and Sertoli Cells The Sertoli cell plays a prominent role in sex differentiation. Through experiments conducted with XX–XY chimeras it was determined that Sertoli cells are the only cell type within the testis required to have a Y chromosome for normal testis development [114]. Sertoli cells express Sry, the male sex-determining gene, and are thought to control male development by influencing the differentiation of other cell types in the XY gonad [95, 127, 128]. Other genes in the sex determination pathway have been studied to determine their function during testis differentiation. Of these genes, Sox9 (Sry-related HMG box-9) has received the most attention [129]. Sox9 is basally expressed prior to Sry expression [130], but is upregulated just prior to testis morphogenesis [131]. Most evidence would suggest that Sry upregulates Sox9 expression; however, there have been no experiments that demonstrate direct regulation of Sox9 by Sry. Overexpression of Sry or Sox9 causes sex-reversal in XX mice [13, 132]. Individuals with mutations in either Sox9 or Sry will develop as XY females [80, 133]. Sox9 has also been demonstrated to directly regulate MIS in the presence of SF-1 [134]. MIS causes regression of female reproductive tract structures and is one of the first Sertoli cell markers within the testis. Therefore, Sox9 may be as critical to testis development as Sry. The Sox9 gene may not be without its compensatory molecules. Another Sox gene, Sox8, has a similar overlapping expression pattern to Sox9 during mouse testis development [135]. Sox8 also has been determined to regulate the MIS gene, in a manner similar to Sox9, acting synergistically with Sf1. Both Sox8 and Sox9 appear to have arisen from a common ancestral gene and may have retained common functions during testis determination [135]. Sry appears to upregulate several genes such as Sox9 and Sox8, which may compensate for each other during testis development. The absence of estrogen and estrogen’s actions may be necessary for testis development. Estrogen receptor double knockout XX mice (without either an α or β receptor) experience female-to-male sex reversals [136]. The gonads of these sex-reversed mice display an overall increased expression of Sox9 [137, 138]. On further characterization of the somatic cell type within these ovaries, it was determined they express Sertoli

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cell–specific genes Sox9, TIF1β [139] and TIF2 [140] in both prepubertal and adult ovaries [138]. Therefore, estrogen may be responsible for suppression of testisspecific genes and maintenance of granulosa cell differentiation within the ovary. Further investigation is necessary to determine if the somatic cells within the estrogen receptor double knockout XX gonads are truly Sertoli cells or a cell type that is somewhere in the differentiation pathway between granulosa and Sertoli cells. The GATA family of transcription factors has been shown to be important in the regulation of genes directing differentiation in multiple organs. In the testis and ovary of the mouse, three members of the GATA family are expressed: GATA-1, 4, and 6. Only GATA-4 is expressed during gonadal differentiation and is present in the somatic cells of the bipotential gonad of the mouse at 11.5 dpc (Fig. 4.2). GATA-4 appears to be expressed in a sexually dimorphic manner with high levels of expression in Sertoli cells of the differentiating testis and reduced expression in the developing ovary [141]. In the pig, expression of GATA-4 is in the coelomic epithelium as it migrates and proliferates to form the indifferent gonad, making this gene a good marker for early gonadal development [21]. The function of GATA-4 during gonadal development remains to be determined. However, GATA-4 was found to activate the MIS promoter through a Gata element that appears to be highly conserved in several species [141]. Expression profiles of both GATA-4 and MIS also appear to be coordinately regulated with expression of GATA-4 preceding that of MIS in both rodents [141] and pigs [142]. Thus, GATA-4, along with several other transcription factors Sf1, Wt1, and DAX-1 [100, 143–145], may be important in regulation of MIS gene expression and Müllerian duct regression within the developing male. Normal function of the GATA transcription factors requires interaction with zinc-finger proteins of the FOG (Friend of GATA) family [146, 147]. One member, FOG-2, is expressed in the developing mouse gonads by 11.5 dpc [148–150]. FOG-2 null mutants die at 14.5 dpc due to cardiac defects. Partial rescue of cardiac function using a cardiac alpha myosin heavy chain driven FOG-2 (αMHC-FOG-2) transgene expressed in the myocardium extends their life span to 17.5 dpc [151]. GATA-4 null mutants die at 7–9 dpc, whereas the use of a GATA-4 knockin allele (GATA-4ki) allows for interaction with FOG-2 and extends the life span of these mice until 13.5 dpc, at which time altered gonadal morphogenesis can be determined [152]. Testis morphogenesis in FOG-2 null αMHC-FOG-2 transgenic fetuses was abnormal and resembled XX ovaries. Examination of the gonadal histology just

after cord formation confirmed that no cords formed in FOG-2 mutant XY gonads. Expression of Sf1 and Wt1 appeared to be normal but expression of Sry was significantly reduced (only 25% of controls). Expression of other Sertoli cell–specific genes, Sox9, MIS, and Dhh, was absent in both FOG-2 and GATA-4ki mutant XY gonads, indicating that Sertoli cell differentiation did not take place. The expression of P450scc, 3β-HSD, and P450c17 were undetectable in both FOG-2 and GATA-4ki mutant XY gonads [153], suggesting that Leydig cell differentiation did not occur either. Therefore, both FOG-2 and GATA-4 are important in Sertoli cell differentiation, seminiferous cord formation, and Leydig cell differentiation. It is possible that both of these transcription factors interact physically with Sry or other genes to initiate enhanced expression of Sry. Without normal expression of Sry, Sertoli cell differentiation and testis morphogenesis cannot occur.

D. Stage IV: Induction of Testicular Morphology and Seminiferous Cord Formation The formation of testicular cords is a key process in establishing the adult testis morphology. The formation of the seminiferous cords, and potentially the formation of sex-specific vasculature, occurs around 11.5–12 dpc (23–25 ts) and is complete by 12.5 dpc (30 ts) [6, 107, 154, 155] (Fig. 4.1) in the mouse. At this time, the Sertoli cells have ceased proliferating [107] and induction of cord formation has been initiated by migration of cells from the adjacent mesonephros into the developing testis to surround the PGC–Sertoli cell aggregates [5, 117]. Mesonephric cell migration is hypothesized to be the result of Sry-regulated expression of paracrine growth factors secreted by Sertoli cells. Paracrine growth factors secreted by Sertoli cells [6, 154, 155] are speculated to act as chemoattractants to induce mesonephric cells to migrate and surround Sertoli cell–germ cell aggregates within the developing testis [156–161]. As the peritubular and mesonephric cells migrate, they may also secrete factors that enable cells within the testis to morphologically differentiate. The ovary of the mouse is basically quiescent at this time and does not form follicular structures until several days postnatally in the rodent. Therefore, formation of seminiferous cords is the first morphological indication of testis differentiation and is a result of Sry gene expression. The process of seminiferous cord formation is reliant on factors produced entirely by the testis. The gonadotropin receptors for follicle-stimulating hormone (FSH) and luteinizing hormone (LH) are not present in the rat testis until approximately 16–17 dpc

Chapter 4 Embryonic Sertoli Cell Differentiation

of testis development [162–164]. Furthermore, the beta subunits for both FSH and LH are not produced by the developing pituitary until approximately 16–17 dpc of embryo development in the mouse [165] with similar later time periods of gonadotropin subunit expression in pigs (after 50 dpc) [166] and sheep (around 70 dpc) [167]. Thus, a paracrine growth factor produced within the testis and triggered by transcriptional regulators such as Sry must be the initiator of testis morphogenesis. 1. Morphological Changes in Sertoli Cells during Sertoli Cell Differentiation and Aggregation An initial step in cord formation is aggregation of Sertoli cells with PGCs. For this aggregation to occur, the Sertoli cells must differentiate and undergo a mesenchymal-to-epithelial cell transformation and polarization [168, 169]. Polarization of the Sertoli cells occurs through accumulation of extracellular matrix proteins such as collagen type IV, laminin [170], fibronectin [171], and heparin sulfate proteoglycan [172] along the basal surface. This correlates with localization of cytokeratin, vimentin, and actin in the basal cytoplasm [170, 173, 174]. As the Sertoli cells differentiate, a number of morphological changes occur including a change in expression of mesenchymal to epithelial cell markers from vimentin to cytokeratin [171] and a change in expression of cytokeratin 19 to cytokeratin 18 (cytokeratin 19 is expressed in the ovary) [175]. In the rat, early gonad expression of laminin α5 and β1 chains are present, but after formation of the testicular cords the laminin α5 chain rapidly disappears. Therefore, the laminin α5 chain can be used as a sexspecific marker for Sertoli cells prior to differentiation into an epithelial cell type. The absence of a laminin α5 chain in the testis can be used to determine that Sertoli cells have undergone differentiation [176, 177]. In addition to laminin, desmin can be used as a marker for Sertoli cell differentiation. In the indifferent gonad, desmin is expressed, whereas after cord formation desmin is no longer expressed within the Sertoli cell [176, 177]. Aggregation of Sertoli cells prior to formation of cords in the testis also requires changes in adhesion of cells to each other and to the extracellular matrix. Both integrin subunits [178] and lectins [179] have been demonstrated to be involved in the early steps of cell aggregation leading to cord formation. Disruption of cell polarization or basal lamina formation by inhibition of protein glycosylation or disruption of actin filaments also prevents cord formation [168, 169]. In addition, treatment of testis organ cultures with retinoic acid [180, 181], or increasing cAMP [182],

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disrupts formation of the basement membrane and perturbs formation of seminiferous cords. 2. Genes Regulating Initial Sertoli Cell Transition and Aggregation The genes that regulate the process of Sertoli cell mesenchymal-to-epithelial cell transition and Sertoli cell aggregation are yet unknown but are thought to be induced by Sry expression. Lhx9 is downregulated as the Sertoli cells form seminiferous cords and initiate secretion of MIS [27]. Recent experiments have localized an increase in Sox9 expression [183, 184] to Sertoli cells after they have aggregated with the PGCs. Therefore, transcription factors that initiate Sertoli cell transformation may also allow for enhanced expression of Sox9 and reduced expression of Lhx9. As the Sertoli cell mesenchymal-to-epithelial transition takes place, a decline in laminin α5 chain gene expression and a significant increase in MIS production occur. Later after birth, when MIS expression diminishes and disappears, expression of the laminin α5 chain increases. Thus, it is possible that transcription factors that positively regulate the laminin α5 chain also negatively regulate MIS [177]. Overall, several extracellular matrix proteins can be used as markers to determine if Sertoli cells have made the transition from mesenchymal to epithelial cell types in the developing testis. Of those extracellular matrix proteins, the most promising appear to be the laminin α5 chain and cytokeratin 19. 3. Mesonephric Cell Migration A second step in seminiferous cord formation is the migration of mesonephric cells from the adjacent mesonephros into the differentiating testis (Fig. 4.1). Sry expression is necessary in order to have normal mesonephric cell migration [6, 111]. Removal of the mesonephros prior to mesonephric cell migration results in no seminiferous cord formation. Therefore, the mesonephric cell migration appears to be crucial to testis development. If migration of mesonephric cells is blocked, or if the mesonephros is removed, no seminiferous cords will form [5, 117]. Also, delayed cell migration leads to cord formation occurring at a later time period as seen in B6 XYAKR mice. Alternatively, in B6 XYPOS mice, which have reduced Sry expression, cell migration is nearly absent and these mice form ovaries or ovotestes [113]. Mesonephric cell migration is directed by cells within the testis. This was elegantly demonstrated in experiments where mesonephric cells migrated through ovaries toward testis explants [6]. Therefore, these

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experiments support the concept that a cell type in the testis induces mesonephric cell migration. The Sertoli cell is the logical candidate to produce paracrine growth factors, which induce mesonephric cell migration [6, 155]. This was concluded since the only other cell type present is the germ cell and germ cell–deficient mice have normal cord formation [185]. Several factors have been proposed to be the Sertoli-derived chemokine, due to alterations in seminiferous cord formation in vitro [156–158] and altered testis morphogenesis in vivo [159, 161]. These factors are discussed in detail later. Several different cell types from the mesonephros have been proposed to migrate into the differentiating testis including endothelial cells, preperitubular cells, and cells associated with vasculature [5]. It is difficult to distinguish cell types at this point in gonadal development because some of the cell markers have not been established. However, most literature would agree that both endothelial and preperitubular cell types migrate from the mesonephros and are involved in completion of seminiferous cord formation [5, 186, 187]. Whether these cells migrate coordinately in response to a single cell migration factor or independently due to separate cell migration factors is unknown. It is also unclear as to whether preperitubular cell migration is dependent on endothelial cell migration or vice versa. In addition to preperitubular and endothelial cells, several laboratories have independently reported migration of cells that appear to be Leydig cell precursors [186, 187]. In both experiments, identification of mesonephric cell types during migration was tracked through the use of cocultures with mesonephroi from either ROSA26 (expressing β-galactosidase) or EGFPtransgenic mice in combination with nontransgenic indifferent testes. Migrating cells with either β-gal or GFP were identified through morphology, [187] positive staining for 3β-HSD, or in vitro culture characteristics after isolation [186]. The results of these experiments suggested that a portion of the Leydig population originated from mesonephric cells that migrate into the testis prior to cord formation. Other laboratories have demonstrated that a portion of Leydig or interstitial cells may be derived from the coelomic epithelium. In experiments where cells of the coelomic epithelia were injected with a DiI fluorescent dye, cells migrating after 12.5 dpc were localized to areas outside of the seminiferous cords, indicating that they were precursors of the interstitial or Leydig cell populations [112]. More research is necessary to determine the percentage of cells that originate from either the coelomic epithelium or from the mesonephros and whether those cells become fetal Leydig cells.

The window of opportunity for proper mesonephric cell migration and induction of testis-specific genes also appears to be critical for normal cord formation and testis development. Induction of migrating cells from the mesonephros can occur as late at 16.5 dpc; however, the best time for proper migration within the male mesonephros is before 12.5 dpc. It is postulated that after 12.5 dpc all critical cells from the male mesonephros have migrated and been donated to the adjacent differentiating testis, suggesting that after 12.5 dpc there are no available cells to migrate into the donor gonad within the organ cocultures [154]. In contrast to this theory, events may also that occur with the developing mesonephros that do not allow for cell migration after a certain development period. Differentiation of the male and female reproductive tracts from the Wolffian and Müllerian ducts may prohibit further cell migration from the mesonephros. Or paracrine factors that induce cell migration may not have viable receptors within the mesonephros after a certain period during gonadal development. The establishment of a critical time for mesonephric cell migration may also explain how a delay in the signal for mesonephric cell migration, as observed in lines of mice that form ovotestis, may result in aberrant gonad formation [154]. 4. Growth Factors Involved in Mesonephric Cell Migration and Testis Cord Formation Many experiments have been conducted to identify Sertoli cell–derived growth factors that induce seminiferous cord formation. Seminiferous cord formation is regulated by factors expressed within the testis since neither gonadotropin receptors nor subunits for gonadotropin hormones are present within the testis or produced within the pituitary gland at this time [10, 162, 163, 165]. Logically the Sertoli cell is the presumed cell type that induces seminiferous cord formation because it expresses Sry and is the first cell type to differentiate in the testis. In addition to this information, germ cell deficient mice still have normal cord formation. Therefore, in contrast [185] to the ovary, which is dependent on primordial germ cell migration for follicle formation [188, 189], the testis does not need germ cells to migrate into the developing gonad for cord formation. These observations support the hypothesis that Sertoli cells secrete a diffusible factor that induces mesonephric cells to migrate toward and enclose the Sertoli cell–PGC aggregates within the developing testis. The mesonephric cells as they migrate may also secrete factors that allow for seminiferous cord formation to be completed. The role of specific growth factors is described later.

Chapter 4 Embryonic Sertoli Cell Differentiation

5. Fibroblast Growth Factor 9 The family of fibroblast growth factors encompasses approximately 22 members, which are widely involved in many developmental processes. Expression of fibroblast growth factor 9 (FGF-9) occurs in at 11.5 to 12.5 dpc in the indifferent and differentiating testis of XY mice and is not present at any time in XX mice or in mesonephros from either XY or XX individuals. In FGF-9 knockout mice, the phenotype ranged from testicular hypoplasia to complete sex reversal. FGF-9 mutants also displayed a reduction in coelomic epithelial proliferation and decreased numbers of Sertoli and interstitial cells [159]. Because a certain proportion of Sertoli cells and interstitial cells comes from the coelomic epithelium, any reduction in cells of the coelomic epithelium would also affect the cell types derived from that affected tissue. FGF-9 was also demonstrated to induce mesonephric cell migration into XX gonads and cause increased Sox9 expression. Furthermore, organ culture experiments that blocked endogenous FGF-9’s actions resulted in impaired testis cord formation. FGF-9 mutant embryos had impaired expression of Sox9, and XX organ cultures treated with FGF-9 formed seminiferous cords with increased expression of Sox9. Therefore, FGF-9 may be involved in induction of mesonephric cells into the testis in order to elicit cord formation. Conversely, FGF-9 may affect the ability of Sertoli cells to aggregate or differentiate in order to upregulate Sox9 at cord formation [159]. In either scenario, FGF-9 is a potential candidate gene that could be downstream of Sry to elicit the actions within the testis that are Sry dependent during testis differentiation. 6. Platelet Derived Growth Factor and PDGFRa Platelet derived growth factors (PDGFs) mediate epithelial-to-mesenchymal interactions (cell proliferation, migration, and differentiation) and are generally expressed in epithelial or endothelial cells, whereas the receptors for PDGF are expressed in mesenchymal cells [190]. The PDGF family is composed of four ligands (A, B, C, D) and two distinct receptors: PDGFRα and PDGFRβ. PDGFRα can bind both A, B, and C homodimers and AB heterodimers, whereas PDGFRβ can bind only the B and D homodimers. PDGF-A is expressed in both XX and XY gonads at 11.5 dpc, and by 12.5 dpc it is strongly expressed in Sertoli cells within the seminiferous cords, whereas expression in the XX gonad is diminished. PDGF-B was expressed in both the XX and XY gonads in endothelial cells by 13.5 dpc [191]. In contrast, PDGF-C was expressed at 11.5 dpc in the coelomic epithelium [192], in the gonadal

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mesonephric boundary, and at lower levels in cells scattered through the gonad. Inhibitors to the PDGFR signal transduction pathway in rat testis organ cultures caused abnormal or “swollen” cord formation with increases in cord diameter and reductions in total numbers of cords [193]. Similar results were seen in PDGFRα homozygous mutant mice, demonstrating abnormal cord formation with no distinct division between seminiferous cords and interstitial compartments [191]. Furthermore, there were defects in vascular development, mesonephric cell migration, cellular proliferation, and Leydig cell differentiation [191]. Thus, it appears that PDGF ligands and PDGFRα may play important roles during testis development. 7. Hepatocyte Growth Factor Hepatocyte growth factor (HGF) mediates its effects through a tyrosine kinase receptor c-met [194]. In general, during mouse development HGF mediates epithelial-to-mesenchymal transitions [195, 196] and branching morphogenesis of the lung [197] and also appears to be critical in development of the placenta [198, 200]. HGF is expressed in the developing gonad at 11.5 dpc around the time of seminiferous cord formation. A procedural consideration is that mouse organ cultures generally are cultured with 10% fetal calf serum in order for seminiferous cords to develop in vitro. In the absence of serum, cord structures will not form in vitro. In contrast, in the rat, serum appears to inhibit cord formation, so organ culture experiments are conducted in serum-free defined media [201]. HGF has been demonstrated to support seminiferous cord formation in mouse organs that were cultured without serum [158]. Therefore, the presence of HGF may be necessary or permissive in order for cord formation to occur in vitro. In addition, HGF does induce mesonephric cell migration and could be a potential growth factor involved in the process of cord formation in the mouse [202]. 8. Neurotrophin Growth Factor Family A role for neurotrophins and their receptors has been implicated in many systems at sites of mesenchymalto-epithelial cell interactions [203–209]. The four neurotrophin ligands are neurotrophin-3 (NT3), nerve growth factor (NGF), brain derived growth factor (BDNF), and neurotrophin-4/5 (NT4/5) [210]. All neurotrophin ligands bind to a low-affinity receptor p75 neurotropin receptor (p75NTR) and each has more specific high-affinity trk receptors. NT3 binds to trkC,

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NGF binds to trkA, and both BDNF and NT4/5 bind to trkB [211–214]. In rat testis, the low-affinity receptor for neurotropins, p75NTR, had been determined to be present in a sex-specific manner in the testis at the time of seminiferous cord formation. Expression of p75NTR was present in the mesonephros prior to cord formation and then was localized to cells surrounding seminiferous cords at cord formation. Later at postnatal day 0 (P0), p75NTR was expressed in the single layer of peritubular cells that surrounding the seminiferous cords [156]. Similar expression profiles were determined in mice [215] and humans [216] for p75NTR during testis differentiation. p75NTR was determined to be a marker for early migrating mesonephros cells that develop into preperitubular cells [217]. NT3 and its high-affinity receptor trkC were expressed in the Sertoli cell and within specific cells of the mesonephros prior to cord formation. Subsequent experiments determined that anti-sense NT3 oligonucleotides [157] and trkC-IgG fusion proteins had partial inhibition of seminiferous cord formation [156] while both trk signal transduction inhibitors (K252A and tyrophostin AG879) inhibited seminiferous cord formation [156, 161]. The trk specific signaling inhibitor AG879 was shown to inhibit seminiferous cord formation by inhibition of mesonephric cell migration. Furthermore, NT3-coated beads induced mesonephric cells to migrate into XX gonads similar to FGF-9 [161]. Taken together, these results demonstrate that NT3 may be a Sertoli cell–derived chemotactic factor involved in mesonephric cell migration. Testicular cord formation in both trkC and trkA knockout mice appears to be developmentally delayed with reductions in the number of cords at E14 [160]. In addition, trkC knockout mice had reduced interstitial area at E13 and a reduction in the number of seminiferous cords at E19. Additional germ cell abnormalities were detected with both receptor mutants demonstrating reduced germ cell numbers during embryonic development. Furthermore, P19 trkA knockout mice had increased germ cell apoptosis when compared to wildtype controls. The trkA knockout mice also appeared to have increased aberrant branching of seminiferous cords at E14 and P19. Double knockout mice (trkC/trkA) did not have increased testis morphological abnormalities when compared to phenotypes of either knockout alone; however, the numbers of double knockouts were low, 2 in 80 embryos, one of which was female [160]. Therefore, absence of all trk receptors may be necessary to determine compensatory functions of the neurotropin ligands and receptors. A similar observation was found in triple insulin receptor knockout mice [218].

In human embryonic testis cultures, treatment with the trk receptor inhibitor K252A resulted in reduced numbers of peritubular cells, Sertoli cells, and gonocytes [216]. Observations support a role for neurotropins in testis cell growth and survival. Therefore, the neurotrophin family appears to have functions in both testis morphogenesis and germ cell development. Sertoli cell–expressed NT3 appears to be a chemotactic factor for the migrating mesonephros cells that are required for testis cord formation. 9. Estrogen or Estrogen receptor Estrogen receptor double knockout mice (those lacking both the alpha and beta receptor genes) have been demonstrated to display female-to-male sex reversal in XX mice. Furthermore, as discussed earlier, estrogen receptor double knockout mice have somatic cells that express Sox9, which is a Sertoli cell–specific gene. Researchers concluded from these experiments that estrogen was important in suppression of Sox9 expression in somatic cells and without its action this somatic cell linage was capable of expressing Sox9 and initiating the testis development pathway [137]. Both estrogen and androgen receptors are present around the time of seminiferous cord formation in mice [219] and rats [220, 221]. Estrogen may also have adverse effects if expression occurs abnormally in males. Experiments utilizing rat testis organ cultures demonstrated that cord formation was inhibited in a doseresponsive manner when exposed to increasing levels of estrogen. Furthermore, cord formation was also inhibited when organs were exposed to compounds that were antiandrogenic [220]. The effects of estrogen may be directly on Sertoli cell function because estrogen receptor alpha is present in Sertoli cells at this developmental period. However, the antiandrogenic effects may be more indirect because the androgen receptor is localized to germ and interstitial cells around cord formation [220]. Steroid hormones and their receptors are potentially involved in critical morphological gonadal development and maintenance of cell differentiation within the testis and ovary. 10. Insulin Receptor Family The insulin receptor family composed of insulin receptor (Ir), insulin-like growth factor receptor 1 (Igfr1), and insulin-receptor–related receptor (Irr) is important in multiple cellular pathways to direct growth and metabolism. Single receptor null mutants for Ir die after birth due to ketoacidosis [222, 223] and Ir die at birth due to respiratory failure [224]; whereas Irr mutants are viable and show no phenotype [225]. Testis morphogenesis

Chapter 4 Embryonic Sertoli Cell Differentiation

and cord formation in single null mutant mice appear to be normal; however, gross morphologic sex reversal (testis to ovary) occurs in triple mutants Ir/Irr/Igfr1. Expression of testis-specific genes, Sox9 and MIS, are absent in mutant gonads with coordinate high expression of ovarian-specific genes, Wnt4 and Figα. In double null mutants Ir/Igfr1 and compound heterozygous Ir+/−/Irr−/−/Igfr1–/–, Ir−/−/Irr−/−/Igfr1+/− knockout mice the degree of sex reversal was more variable with lowered expression of testis-specific Sox9, MIS, and Insl3 and ovarian-specific genes, Wnt4 and Figα. Gonadal cells from the triple knockouts also proliferated more slowly, resulting in a smaller gonad than their wild-type littermates [218]. Reduced proliferation is also critical because proliferation of the coelomic epithelium is dependent on Sry and is necessary for normal development of the XY gonad. Therefore, this research indicates a critical role for the insulin receptor family in testis development and adds yet another growth factor family to the list that is involved in the complex pathway of gonadogenesis. 11. Similar Signal Transduction Pathways That Affect Seminiferous Cord Formation Several of the growth factors (FGF-9, HGF, NT3, insulin) that are involved in seminiferous cord formation have a common signal transduction pathway that is activated, which is PI3 kinase [226]. All of these growth factors also appear to induce mesonephric cell migration or have a potential role in induction of mesonephric cells into the differentiating testis. Experiments using a PI3 kinase inhibitor specific to the 110 isoforms of class I PI3 kinases demonstrated in rat testis organ cultures (in a dose responsive manner) an inhibition of seminiferous cord formation. These observations demonstrated that a PI3 kinase inhibitor and not a Map kinase inhibitor were critical to seminiferous cord formation through inhibition of mesonephric cell migration. Therefore, the PI3 kinase signal transduction pathway may be a common pathway that multiple growth factors induce through direct or indirect Sry actions to promote mesonephric cell migration and seminiferous cord formation [226]. An inhibitor of the protein kinase A signal transduction pathway, forskolin, also inhibited mesonephric cell migration in mouse gonad cultures. However, there were no effects on Sertoli cell differentiation because Sertoli cells were capable of expressing MIS, Sox9, and Dhh. Forskolin was utilized because the protein kinase A pathway is a signal transduction pathway activated by the Dhh gene within the testis [76]. The Dhh gene is expressed in Sertoli cells during testis development, and its receptor patched1 (Ptc1) is

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expressed in interstitial, peritubular, Leydig, and endothelial cells [77, 227]. The phenotype of Dhh null mice is variable depending on the genetic background. Some backgrounds have normal testis structure but defective spermatogenesis [227]. In contrast, Dhh mutants from other genetic backgrounds exhibit defects in development of peritubular cells, an absence of basal lamina, apolar Sertoli cells, anastomotic testicular cords [228], and defects in Leydig cell differentiation [77]. Therefore, Dhh may initiate critical steps during testis differentiation. In summary, both protein kinase A and PI3 kinase appear to be important pathways for regulation of seminiferous cord formation potentially through regulation of cell migration. Previous research has demonstrated that calphostin C, a potent protein kinase C inhibitor, did not have any effect on seminiferous cord formation [156]. Because testis morphogenesis is crucial for male adult testis function, multiple signal transduction pathways may be necessary to ensure proper regulation of seminiferous cord formation within the male gonad. 12. Pattern of Sertoli Cell–Specific Gene Expression and Cord Formation Sry expression is first detected in the central region of the gonad around 11 dpc and its expression spreads to the anterior and posterior poles within the testis [15, 113]. This suggests that pre-Sertoli cells within the central region of the testis are more advanced than other pre-Sertoli cells within the indifferent gonad at this time. There do not appear to be morphological changes in cells between the regions within the indifferent gonad at this stage. Therefore, there must be differential gene regulation or altered access to factors (potentially through closer vascular supply) that initiate Sertoli cell differentiation and Sry expression. Like Sry expression, seminiferous cord formation also occurs from the central region of the gonad into the anterior and posterior poles of the gonad. The chronology of testis cord formation also explains the morphology of ovotestes, which comprise ovarian tissue on either side of testicular tissue [15, 113]. Recent experiments have demonstrated different abilities of certain regions of the gonad to form seminiferous cords during development. The central region of the gonad (separated from both the anterior and posterior regions) can initiate cord formation at a much earlier stage in gonadal development (12–14 ts) when compared to both the posterior and anterior regions (15–17 ts) [229] (Fig. 4.3). Therefore, this supports previous conclusions that preSertoli cells in the central portion of the gonad are at a more advanced stage of differentiation, have greater

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Gonads

Anterior

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Anterior

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Leydig cell differentiation

FIGURE 4.3 Regional differences in somatic cell differentiation within the gonad. Arrows demonstrate how each somatic cell type differentiates: Sertoli cells differentiate from center to pole, whereas Leydig cells differentiate from anterior to posterior within the differentiating XY gonad [15, 229].

access to factors that allow them to develop differentiated functions earlier, or have a greater innate capacity to initiate events earlier during testis development. 13. Müllerian Inhibiting Substance/Anti-Müllerian Hormone MIS is a member of the transforming growth factor β (TGFβ) family of growth factors and is expressed specifically in the Sertoli cells of the testis around 11.5 to 12.5 [230] (Fig. 4.2). The MIS gene is the first Sertoli cellsecreted protein that is produced at or after the time of seminiferous cord formation. The role of MIS in the testis appears to be severalfold. MIS received its name initially because it was the factor that caused Müllerian duct regression and eliminated any precursors of the female reproductive tract. Around 11.5 dpc the mesonephros contains two duct structures that are precursors of the female, the Müllerian duct or paramesonephric duct, and the male, Wolffian duct or mesonephric duct, reproductive tract structures [231, 232]. In the female XX gonad, where no Sertoli cell differentiation occurs, the Müllerian duct will develop and eventually form portions of the female reproductive tract structures. In contrast, in the male, MIS expression by the Sertoli cell will cause regression of the Müllerian duct through initiation of Müllerian duct cell apoptosis. In MIS null male mice, the Müllerian duct fails to regress and both male and female reproductive tract develop [19, 233]. Recently, MIS has been identified as a factor that induces mesonephric cell migration. However, because no abnormalities were found in early testis development within MIS null mice, it is believed that this function of MIS may be redundant and shared with another member of the TGFβ family [234]. 14. Endothelial Cell Migration and Development of Vasculature Sex-specific vascularization and development of a large blood vessel supply occurs in the developing

testis (XY gonad) under the coelomic epithelium by 12.5 dpc in the mouse. There is no research available to suggest if testis vasculature develops at the time of seminiferous cord formation or after testis morphogenesis. However, because both endothelial cells and preperitubular cells migrate into the differentiating testis as testis morphogenesis is initiated, it is hypothesized that both seminiferous cord formation and vasculogenesis in the testis occur simultaneously. Therefore, in addition to surrounding the Sertoli cell–PGC aggregates, the migrating mesonephric cells may also initiate the formation of vasculature within the developing testis directed by Sertoli-secreted factors downstream of Sry expression [235]. At 11.5 dpc the endothelial cells express both venous (EphB4) and arterial (Ephrin B2) markers in both the XX and XY gonad, which suggests that vascular development at this time is not sex specific. However, just after 11.5 dpc, vasculature within the developing testis undergoes rapid reorganization, causing the formation of the coelomic blood vessel, organization of endothelial cells to the interstitium outside of the seminiferous cords, and extensive branching of vessels [235]. By 12.5 dpc the vasculature of the testis labeled positive for the arterialspecific marker Ephrin B2, whereas the ovary at this time still has both arterial and venous makers. Sertoli [235] cells have been determined to express many growth factors that cause endothelial cell migration such as vascular endothelial growth factor (VEGF) [236] and PDGF [191, 193]. Therefore, the development of the cell-specific vasculature around the time of cord formation is presumably mediated by Sertoli cell–secreted proteins, which are expressed downstream of Sry expression and Sertoli cell differentiation.

E. Stage V: Development of a Functional Testis After seminiferous cord formation is completed, the other cells within the testis initiate differentiation. It is proposed that the Sertoli cell continues to differentiate

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Chapter 4 Embryonic Sertoli Cell Differentiation

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In some species timing of seminiferous cord formation greatly impacts Leydig cell differentiation and initiation of steroidogenesis. 3β-HSD production, which is a requisite for androgen production, was detected after 15 dpc in the rat [239] and at 13 dpc in the mouse [240]. In the pig, Leydig cell differentiation was determined to occur at 28 dpc (due to positive staining of cells in the interstitium for P450c17), which was several days after Sertoli cell differentiation and cord formation (26 dpc) [142]. Recently, it was determined that although Sertoli cell differentiation occurs from center to pole (Fig. 4.3), the 3β-HSD–positive cells differentiate from the anterior region of the gonad to posterior. This supports earlier studies where Sf1-positive cells were found in the anterior region of the indifferent gonad and were thought to be precursors of both the adrenal steroidogenic cells and the Leydig cells of the differentiating testis [241]. Furthermore, cells in the mouse gonad stained positive for 3β-HSD at 11 ts, which is much earlier than has been reported. The presence of 3β-HSD indicates that the somatic cells of the testis destined to become Leydig cells are capable of steroidogenesis (specifically testosterone production) at a much earlier time point than previously thought. Testosterone is critical to the developing male because androgens stabilize the Wolffian duct derivatives for normal male duct development [242, 243]. Therefore, appropriate differentiation of somatic cell types in the testis prior to and around the time of cord formation is crucial not only to the normal development of the testis, but also for the continued presence of the Wolffian duct and future development of the male reproductive tract.

% PCNA Stained Cells

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of cord formation, the testis undergoes a dramatic sexspecific growth. At 15 dpc in the rat, the testis is twice the size of the same age ovary [244]. Proliferation of the Sertoli cell population is important because each Sertoli cell can support a finite number of germ cells in the adult testis [9, 245]. An increased number of Sertoli cells correlates with increased spermatogenic capacity. After puberty, the Sertoli cells are no longer mitotic and the population of Sertoli cells is established. The rodent has two critical developmental periods when Sertoli cells are proliferating—during late embryonic development and again during early postnatal testis development [9, 246]. The greatest rate of embryonic Sertoli cell proliferation occurs from 18 dpc to P0 in the rat [247] with proliferation in late gestation as well in the pig [248]. Other testicular cell populations also proliferate from cord formation to birth (from 14 dpc to P0) [247]. At 14 dpc the germ cells, interstitial cells, and peritubular cells are all proliferating rapidly. The Sertoli and peritubular cells continue to proliferate at a high rate at P0, whereas the germ cells experience a dramatic reduction in proliferation. After birth all the cells proliferate until the onset of puberty, when Sertoli cells become postmitotic [247] (Fig. 4.4). Many factors have been shown to affect cell growth in the embryonic testis and some of the growth factors also affect organization of seminiferous cords. Several growth

% PCNA Stained Cells

and proliferate after seminiferous cord formation and that interstitial and Leydig cells differentiate after initial Sertoli cell differentiation. In part, this may be due to factors produced by the Sertoli cells that direct interstitial, peritubular, and Leydig cell differentiation. The peritubular layer of cells becomes identifiable from the interstitial or Leydig cells at 15 dpc in the rat [237]. The peritubular cells become a single layer of cells surrounding the seminiferous cord in rodents and a multilayered cell layer in some domesticated livestock [238]. A major role of the peritubular cell is to form the outer layer of the seminiferous cord and to jointly form with the Sertoli cell the blood–testis barrier to protect haplotype germ cells from any attacking immune cells.

FIGURE 4.4 Proliferation of each cell type within the developing

2. Sertoli Cell Proliferation Growth and proliferation of cells within the testis are critical for testis morphogenesis. After the process

rat testis from E14 through postnatal day 10. Most cell types accomplish most of their proliferation during embryonic development. Proliferation was assessed by percentage of cells staining positively for PCNA [247].

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factors that regulate cell growth in the embryonic testis are discussed in the following sections. 3. Transforming Growth Factor Alpha Family The transforming growth factor α (TGFα) and epidermal growth factor (EGF) have been associated with multiple roles during testis development. EGF increases production of inhibin, androgen-binding protein, and transferrin by Sertoli cells [249, 250]. EGF also affects Leydig cell function by altering cholesterol transport and the number of LH receptors and by regulating testosterone production [251, 252]. TGFα has been demonstrated to stimulate peritubular cell proliferation and migration in culture [253, 254]. TGFα mRNA has been detected in cultured Sertoli and peritubular cells [253, 254] and TGFα protein was found in Leydig cells [255]. EGFR, the receptor for both EGF and TGFα was found at the protein level in adult Sertoli and Leydig cells, and at the mRNA level in prepubertal peritubular cells [253, 256, 257]. TGFα and EGF have been demonstrated to increase cell proliferation at the time of cord formation and during late embryonic testis development [247, 258]. Most of these experiments were in vitro utilizing TGFα and EGF antagonists and receptor signaling antagonists. Most of the proliferative effects of TGFα appeared to be on Sertoli cells and cells in the interstitium. Knockout mice for EGFR (the receptor for TGFα) demonstrated a reduction in interstitial cells surrounding the seminiferous cords at 18 dpc. These differences were compensated for by birth in these mice [247]. In addition to its effect in the testis, EGF is also important in mediating the effects of testosterone to stabilize the Wolffian duct and allow it to differentiate [242, 243]. Therefore, both TGFα and EGF appear to be important in proliferative and morphogenic processes within the developing testis and reproductive tract. 4. Transforming Growth Factor Beta Family TGFβ isoforms are growth factors that regulate growth and development in many different cell types. All three mammalian isoforms have been identified in the adult testis [181, 259–264]. TGFβ1, TGFβ2, and TGFβ3 have also been localized to the embryonic testis. TGFβ1 appears in the testis at 14.5 dpc in the rat (after cord formation) and has been localized to the seminiferous cords and interstitium. TGFβ2 was localized to the Sertoli cells at 14.5 dpc, with localization to the Leydig cells (along with TGFβ1) at 16.5 dpc [264, 265]. In contrast, TGFβ3 was localized to preperitubular cells at 14 dpc [181] and to germ cells from 14.5 dpc to birth [181, 266]. This localization of TGFβ3 suggests that it

may be involved in critical processes involved in the restructuring of the gonad and germ cell maturation/ proliferation during embryonic testis development. One role of TGFβ1, TGFβ2, and TGFβ3 from 16.5 to 20 dpc is to regulate LH stimulated steroidogenesis from the Leydig cells [266, 267]. TGFβ3 may be involved in seminiferous cord formation; however, it does not appear that TGFβ1 or TGFβ2 is involved in differentiation of the testis, only in testis proliferation. Gene knockout and overexpression experiments with TGFβ have demonstrated that precise regulation of each isoform is essential for survival. TGFβ1 null mice are phenotypically normal until approximately 3 wk after birth and then they develop a severe wasting syndrome [268]. Reproductive traits and organs within TGFβ1 knockout mice have not been extensively studied. However, there are significant deviations from normal mendelian ratios, resulting in decreased offspring for both heterozygotes and homozygotes carrying the allele with the TGFβ1 gene disruption. TGFβ2 knockout mice possess abnormalities in urogenital development. These alterations within the reproductive tract ranged from testicular ectopia to unilateral testicular hypoplasia to complete absence of the epididymis [269]. However, these investigators did not conduct an extensive examination of testis histology to determine what processes in testis development were affected with the TGFβ deletion. TGFβ signals through the interactions of TGFβ receptor I and TGFβ receptor II. TGFβs bind to TGFBRII, which phosphorylates TGFβRI, causing the [270] activation of the SMad family of transcription factors [271]. TGFβRI and II are present in germ cells of mice around the time of cord formation (11.5 dpc) [272]. In the rat TGFβII has also been localized to germ cells at the time of cord formation (13.5 dpc) and later in the Leydig cells (16.5 dpc) where TGFβs may regulate steroidogenesis [267]. A requirement for TGFβRIII has been demonstrated in endocardial cell transformation in the heart where a mesenchymal-to-epithelial transition takes place [273]. Therefore, all three receptor isoforms may be important in mediating the actions of the TGFβ ligand isoforms.

5. Neurotrophin Growth Factor Family As stated previously neurotropin growth factors, specifically nerve growth factor and neurotropin-3, are two neurotropins that are secreted by Sertoli cells early during testis development and prior to cord formation [157, 215]. After cord formation they are localized to other cells and may be involved in germ cell maturation or Leydig cell differentiation. In both trkC and trkA (receptors for NT3 and NGF, respectively) knockout

Chapter 4 Embryonic Sertoli Cell Differentiation

mice, reductions were observed in the area of the interstitium during embryonic development, which may suggest that the neurotropins are necessary for interstitial cell proliferation. In addition, reductions in germ cells during embryonic development were observed that indicate germ cell maturation/proliferation or survival may be dependent on neurotropins [160]. Both NGF and NT3 demonstrated a dose-dependent increase in incorporation of tritiated thymidine in day 0 testis cultures [157]. Therefore, NGF and NT3 are potential regulators of testis growth during the late embryonic period after cord formation. 6. PDGF PDGF appears to be important for formation of normal cords as previously discussed. PDGF also appears to affect the differentiation of the Leydig cell population [191]. PDGF was not determined to stimulate growth of P0 testis when experiments were conducted using tritiated thymidine [193]. Therefore, PDGF appears to affect early morphogenesis and cell proliferation events during embryonic testis development. 7. Steroid Hormones and Their Receptors Estrogens and androgens both have been demonstrated to affect cell growth within the embryonic testis [220]. There also is accumulating evidence that high levels of estrogens and antiandrogens will affect cell growth within the testis and have the potential to cause adult testis abnormalities [274, 275]. Estrogens and androgens, as discussed previously, can have dramatic effects on gonadal development, growth of specific cell populations, and differentiation of the reproductive tracts. Therefore, any abnormal production of these steroid hormones can influence embryonic testis development and adult testis function. 8. Thyroid Hormone A wealth of information has been collected on the effects of transient hypothyroidism on early postnatal Sertoli cell differentiation. Briefly, transient hypothyroidism postnatally increases Sertoli cell numbers, increases testis size, and increases daily sperm production in mice and rats [276–281] with similar effects found in rams [282, 283] and cattle [10]. Other experiments conducted in the rat to induce hypothyroidism in pregnant females during late gestation demonstrated no effects on fetal testis growth, but resulted in reduced serum levels of gonadotropins and delayed maturation of Sertoli cells and puberty [284]. Recently, two lines

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of pigs that have divergent adult testis size were evaluated to determine if alterations in endocrine hormones may elicit the differences in testis size. The Meishan Chinese breed of pigs have a shorter Sertoli cell proliferation period, which results in smaller seminiferous cord diameter, larger interstitial area, and earlier age at puberty when compared to European White Cross pigs [248, 285]. Meishan pigs were demonstrated to be hyperthyroid during the late fetal period, which correlated with the time when Sertoli cell proliferation was reduced. In addition to increases in thyroid hormone, expression of thyroid receptor β1 was also enhanced at this time, which suggested enhanced testis sensitivity to thyroid hormone. Therefore, high levels of thyroid hormone may shorten Sertoli cell proliferation during late fetal testis development and induce early Sertoli cell differentiation. These alterations in Sertoli cell differentiation [286] may also affect Leydig cell differentiation, resulting in greater interstitial area and reduced area of seminiferous tubules. Further studies are necessary to elucidate the exact mechanisms that regulate thyroid hormone actions on embryonic testis development and morphogenesis, specifically with regard to Sertoli cell proliferation and differentiation.

9. Gonadotropins FSH and LH receptors are present in the testis during late gestation in many species [10, 162, 163]. FSH and LH have been demonstrated to be critical for postpubertal development and proliferation of Sertoli and Leydig cell populations in the testis [287, 288]. Early experiments demonstrated partial gonadotropin dependence of Sertoli cells during fetal development [246, 289]. Recent experiments suggest that gonadotropins do not play a critical role in regulation of Sertoli and Leydig cell populations in the fetal period [290, 291]. Mice that lack the thyroid-specific enhancer-binding protein (T/ebp or Nkx2.1) gene do not develop the thyroid gland, lung, ventral forebrain, and pituitary gland. Because they do not develop a functional pituitary with normal synthesis and secretion of gonadotropin hormones, they are an excellent model to determine how gonadotropins affect morphological testis development and growth. Male mice lacking the T/ebp/Nkx2.1 gene have a normal male phenotype with normal seminiferous cord formation. The testes from these mice produce lower levels of testosterone, have smaller Leydig cells (after 18.5 dpc), and have slightly smaller testes [291]. These data support earlier reports that seminiferous cord formation is independent of gonadotropins.

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Hypogonadal mice demonstrate normal Sertoli cell and Leydig cell differentiation and proliferation during the fetal period [290]. Therefore, gonadotropins do not appear to be critical regulators of embryonic testis proliferation during the late fetal period. Most of the regulation of cell proliferation during fetal development must be through paracrine growth factors produced within the differentiating testis.

IV. SUMMARY The Sertoli cell has been described as the “organizer” of the embryonic testis. Probably a more descriptive title is “orchestrater” because the Sertoli cell initiates, as well as organizes, testis development. Normal testis development is dependent on Sertoli cell differentiation and expression of genes that direct morphological development along the XY pathway. The Sertoli cell is the somatic cell type that initiates differentiation by expressing Sry, which has been described as the master gene in male development. Certainly Sry is critical for testis development. Several other genes, however, such as Sox9, Dmrt1, and DAX-1, appear to need to be expressed appropriately in order for testis differentiation to occur in the indifferent gonad. In the past 12 years, since the identification of Sry multiple genes, morphological events have been elucidated to obtain a better understanding of how testis development progresses from the indifferent gonad. However, further research is necessary to understand (1) what genes are immediately downstream of Sry, (2) what additional paracrine growth factors regulate Sertoli cell differentiation and cord formation, and (3) how embryonic Sertoli cell differentiation impacts other events such as Leydig cell differentiation and adult testis spermatogenic function. Future research using technological advancements such as RNAi, laser capture microscopy, tissue-specific gene expression models, and mouse mutagenesis procedures should add to our understanding of factors regulating embryonic Sertoli cell differentiation and testis development.

Acknowledgments We would like to thank Rebecca Bott and Jill Griffin for assistance with preparation of this book chapter, and also all of the scientists whose research contributed to the writing of this chapter. “The question of how a testis or an ovary develops from its early embryonic primordium is given a rather simple answer in most elementary textbooks. The question is not as clear if one looks at original papers. For more than a century many very good biologists have discussed the question and debated theoretical interpretations without reaching a general agreement.” (Alfred Jost, 1973)

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C H A P T E R

5 Sertoli Cell Biology in Fishes and Amphibians JERRY BOUMA

JOSEPH G. CLOUD

The Jackson Laboratory, Bar Harbor, Maine

Department of Biological Sciences, Center for Reproductive Biology, University of Idaho Moscow, Idaho

animal species is more dynamic and appears to renew itself following each spawning period. Because of the uniqueness of anamniote classes, biological questions concerning the origin and cellular proliferation of the Sertoli cells during testicular recrudescence and the relationship of these events to the resultant fertility and fecundity of the animal during the subsequent spawning period can be addressed. In most fishes and amphibians, testes are paired reproductive organs suspended from the body wall by a mesorchium. Exceptions are the Agnatha (jawless fishes; hagfishes and lamprey), in which the testes are fused, and some bony fishes (Teleostei), which contain a single testis (reviewed in Pudney [1]). Vertebrate testes can be divided into two compartments: the germinal compartment, composed of germ cells and associated somatic or Sertoli cells, and the interstitial compartment, generally consisting of connective tissue, blood vessels, and Leydig cells. Contrary to the situation existing in reptiles, birds, and mammals, in which spermatogenesis occurs in a permanent structure, the seminiferous tubule, the spermatogenic unit in fishes and amphibians is the spermatogenic cyst. This spermatogenic cyst is comprised of Sertoli cells surrounding a cohort of synchronously developing germ cells, separated from interstitial tissue by a basement membrane [2–4]. Although the germ cells develop synchronously within each spermatogenic cyst, the resultant spermatids and spermatozoa within a spermatogenic cyst are not isogenic.

I. INTRODUCTION II. TESTIS STRUCTURE IN FISHES AND AMPHIBIANS III. ORIGIN OF SERTOLI CELLS DURING EARLY TESTIS DEVELOPMENT IV. TESTICULAR RECRUDESCENCE: ORIGIN OF SERTOLI CELLS DURING SPERMATOGENIC CYST FORMATION V. SERTOLI CELL FUNCTION DURING SPERMATOGENESIS VI. SERTOLI CELL PROLIFERATION VII. ROLE OF SERTOLI CELLS IN INITIATION OF MEIOSIS VIII. CONCLUDING REMARKS References

I. INTRODUCTION Sertoli cells in amniotes are differentiated somatic cells located within the seminiferous tubules of the testes. These cells do not proliferate in the sexually mature male, and the number of germ cells supported by a single Sertoli cell appears to be constant for each species; thus, the number of Sertoli cells within the sexually mature testes defines the maximal production of spermatozoa by an individual. Although Sertoli cells of anamniotes appear to have the same physiological functions in the support of spermatogenesis as in amniotes, the Sertoli cell population in many of these SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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Generally, spermatogenesis starts with formation of the spermatogenic cyst; cellular proliferation occurs within the cyst through both mitotic and meiotic divisions, and the resultant spermatozoa are released as a result of rupture of the cyst wall at spermiation. The chapter provides a review of the information concerning the origin of Sertoli cells, during both the initial development of the testes and testicular recrudescence, and their role in the control and support of spermatogenesis. This chapter highlights our current understanding of Sertoli cell biology in major anamniote classes: the elasmobranchs (cartilaginous fishes, Chondrichthyes), teleosts (bony fishes, Osteichthyes), anurans (frogs and toads, Amphibia), and urodeles (salamanders and newts, Amphibia). For information on Sertoli cell cytology in fishes and amphibians, the reader is referred to a number of excellent reviews by Pudney [1, 5], Grier [6], and Loir et al. [7].

II. TESTIS STRUCTURE IN FISHES AND AMPHIBIANS A major structural difference between the testes of anamniotes and amniotes is the biological compartment in which spermatogenesis occurs. In amniotes spermatogenesis is mediated by or involves a permanent germinal epithelium and takes place within a fixed structure, the seminiferous tubule. However, in many anamniotes, the spermatogenic unit is a transient structure, the spermatogenic cyst, and in most fishes and urodeles there does not appear to be a permanent germinal epithelium; spermiation usually accompanies Sertoli cell degeneration [1, 7–9]. Alternatively, anurans have a spermatogenic cycle intermediate to fishes, urodeles, and amniotes. During spermiation, the apical portion of Sertoli cells is shed into the lumen, whereas the basal portion remains and regenerates [1]. These animals are therefore said to have a permanent germinal epithelium. In the anamniotes, the Sertoli cells proliferate up to the spermatocyte stage [10]. In teleosts, two testicular types have been described [2, 3]. Most teleost fishes possess an unrestricted spermatogonial testis in that germ cells, and spermatogenic cysts, are distributed along the entire length of the lobules. In contrast, testes of the superorder Atheriniformes contain a second type, which is classified as a restricted spermatogenic gonad. In these fishes, spermatogonia are restricted to the distal periphery of the lobule, and during spermatogenesis, cysts migrate passively from the periphery toward the sperm duct. A similar form of testicular organization, in which regions contain distinct stages of spermatogenic cysts, are also present in elasmobranchs and urodeles.

In these animals spermatogenic waves occur in a linear fashion, with spermatogenic cysts containing synchronously differentiating germ cells localized to distinct regions of the lobules. This type of organization allows for the isolation and culture of spermatogenic cysts, making it possible to study germ cell–Sertoli cell interactions at specific stages during spermatogenesis.

III. ORIGIN OF SERTOLI CELLS DURING EARLY TESTIS DEVELOPMENT In mammals, the expression of the Y chromosome linked sex-determining gene Sry in the supporting somatic precursor cells in the genital ridge leads to the differentiation of Sertoli cells, which in turn are responsible for testis development [11]. Sertoli cells in mammals therefore are first to differentiate and direct testis differentiation [12]. In medaka (Teleostei; Oryzias latipes), the differentiation of somatic cells of the genital ridge (Sertoli cells in the male, granulosa cells in the female) does not appear to require the presence of germ cells [13], but a recent study demonstrates that the Y chromosome in these cells is important for differentiation of bipotential germ cells into spermatogonia [14]. Similar to mammals, a medaka sex-determining gene on the Y chromosome has been identified [15, 16]. This gene, called dmY or dmrt1Y, contains a DM (doublesex in Drosophila and mab-3 in Caenorhabditis elegans) DNA binding domain motif, and is present in the sexdetermining region on the Y chromosome. The transcript of the dmY/dmrt1Y gene appears to be present only in somatic cells surrounding germ cells of the developing male embryonic gonad. Because two naturally occurring mutations in dmY/dmrt1Y resulted in sex reversal from phenotypic male to female, it has been concluded that this gene is essential for and controls male sex determination in this species [15]. Phylogenetic analysis, however, indicates that dmY/dmrt1Y is not a universal sex-determining gene in fishes [17]. At present, it is unknown whether this gene is required for Sertoli cell differentiation in medaka. The genital ridge in anamniotes, mesodermal in origin, is derived from the coelomic epithelium. Few studies have investigated thoroughly the developmental origin of somatic cell populations of the resultant male gonad. Based on ultrastructural observations in medaka, Hamaguchi [18] described a close association of primordial germ cells following their migration into the genital ridge with flat somatic cells. These associated somatic cells are believed to originate from the coelomic epithelium and envelop the primordial germ cells [18–20]. Additionally, Kanamori et al. [21], using the

Chapter 5 Sertoli Cell Biology in Fishes and Amphibians

same species, reported the existence of two somatic cell types, both derived from lateral plate mesoderm during gonadal development. One somatic cell type is localized in the central region of the developing gonad in close contact with primordial germ cells, whereas the second cell type is present near the periphery. The centrally localized somatic cells appear to give rise to Sertoli cells in the testis and granulosa cells in the ovary, whereas peripherally localized somatic cells are precursors of interstitial cells. Recent morphological investigations on early gonadal development suggested nephrostomial tubules as the site of origin of somatic cells, including Sertoli cells, in sturgeon (Acipenser) [22, 23]. According to the authors’ descriptions, nephrostomial tubules, which are derived from intermediate mesoderm, connect to the coelomic cavity and give rise to the gonadal fold. Proliferating cells from the nephrostomial tubule replace flat mesothelium lining the coelomic cavity, and some of these cells associate with primordial germ cells. Although still a matter of controversy, it appears that amphibian somatic cell populations, including future Sertoli cells, are derived from coelomic epithelium [24, 25]. These findings in fishes and amphibians are therefore in agreement with those for mammalian vertebrates [26]. However, because all conclusions cited regarding anamniotes are based on ultrastructural observations and without specific cellular markers for Sertoli cells, the origin of these cells in fishes and amphibians remains a matter of speculation.

IV. TESTICULAR RECRUDESCENCE: ORIGIN OF SERTOLI CELLS DURING SPERMATOGENIC CYST FORMATION Generally, in anamniotes spermatogenic cyst formation occurs or is initiated as a consequence of Sertoli cells encompassing primary spermatogonia. Initially, Sertoli cells are poorly differentiated, with a flat nucleus, containing a nucleolus (reviewed in [5, 6]). Information regarding the origin of the precursor Sertoli cells in the recrudescing testes is accumulating. Sertoli cell precursors are thought to exist as mesenchymal-like cells in interlobular connective tissue [5]. In rainbow trout (Oncorhynchus mykiss), however, Billard [27] reports the persistence of Sertoli cells as a monolayer in the periphery of the lobule following spermiation. In elasmobranchs, precursor Sertoli cells and primary spermatogonia appear to be present in interstitial tissue of the testes [1]. In this class of fishes, a single Sertoli cell associates with a single primary spermatogonium

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to form a structure called a spermatoblast. Both cell types undergo nine mitotic divisions, after which the Sertoli cell stops dividing, whereas spermatogonia divide four more times. At this point the Sertoli cell-to-germ cell ratio is 1:16. Formation of the basement membrane occurs at this time and completes formation of the spermatogenic cyst; formation of a basement membrane precedes entrance of the germ cells into meiosis. Sertoli cells within spermatoblasts are connected via tight junctions at the basal side, as occurs in mammalian testes [7]. In teleosts, it is generally assumed that a permanent population of germinal stem cells is present in the wall of testicular lobules [1, 28, 29]. However, it has also been speculated that primary spermatogonia migrate annually from the interstitial tissue into the lobules [30, 31]. Protogynous fish are a useful model to study testis formation in adult fish. Gonads of these fishes undergo natural sex reversal following the conversion of an initially developed ovary into a testis. Recent work in the protogynous reef fish, the bluehead wrasse (Thalassoma bifasciatum), suggests that the outer layer surrounding the testis is a source of testicular Sertoli cells [32]. This outer layer, or outer wall, consists of an outer flat epithelial peritoneum and an inner layer containing collagen fibers, fibroblasts, myoid cells, blood vessels, nerve bundles, and mesenchymal-like cells. From an ultrastructural analysis, a continuum was observed between the outer wall and the testicular interstitium, suggesting that somatic cells may migrate into the testicular interstitium and differentiate into Sertoli cells. Similarly, in the protogynous swamp eel (Synbranchus marmoratus), epithelial cells derived from the germinal epithelium, which in turn originate from the peritoneal wall, are reported to differentiate into Sertoli cells based on their nuclear morphology and close association with primordial germ cells [29]. In teleosts, with the development of the spermatocyst, both Sertoli cells and primary spermatogonia are physically separated from interstitial tissue by a basement membrane. The cellular origin of the basement membrane is unknown, but may involve the Sertoli cells and myoid cells. In mammals both of these cell types have been shown to contribute to synthesis of this structure (reviewed in Skinner [33]). In fishes, pinocytotic vesicles have been observed on the basal side of Sertoli cells and in the periphery of myoid cells [7]. Spermatogenesis occurs within the spermatogenic cyst, which grows in size due to cell proliferation resulting from the mitotic divisions of germ cells and Sertoli cells. The number of mitotic divisions of the germ cells is species specific [34]. Sertoli cells that form the walls of the spermatogenic cysts are connected via complex interdigitations, desmosome-like intermediate adhering

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junctions, and associated tight junctions (reviewed in Loir et al. [7]). These connections are localized on the apical side of the Sertoli cells and are only observed after meiosis has been initiated. Small infrequent adhering junctions have also been reported between Sertoli cells and spermatogonia and spermatocytes [7]. Desmosome-like junctions between spermatocytes and Sertoli cells were also reported in the bluegill (Lepomis macrochirus) and bullfrogs (Rana catesbeiana) [35]. In urodeles, spermatogenesis starts at the anterior end of the testes and progresses toward the posterior end [36]. The process of spermatogenesis occurs in a similar fashion as reported in teleosts and eventually ends with rupture of the spermatogenic cyst at spermiation. Early developing Sertoli cells have a fibroblastlike appearance and are derived from invaginated peritoneal epithelia, which form epithelial cords and transform into Sertoli cells [37]. Spermatogenic cysts are formed when Sertoli cells surround primary spermatogonia and are joined by desmosomes. Such Sertoli cells have a crescent-like appearance with an irregularly shaped nucleus containing a nucleolus. Short microvilli appear on the apical surface facing the primary spermatogonia [37], suggesting a regulatory role for Sertoli cells in germ cell development.

V. SERTOLI CELL FUNCTION DURING SPERMATOGENESIS In mammals, Sertoli cells are crucial in regulating and maintaining the process of spermatogenesis. Junctional complexes provide a testis–blood barrier, generating a basal compartment separate from an adluminal compartment. As nurse cells, the Sertoli cells provide nutrients and growth factors to a local environment, which is required for spermatogonia to survive and differentiate into spermatozoa [38–40]. It is generally accepted that this physiological relationship of spermatogonia and Sertoli cells is also the case in anamniotes. Formation of spermatogenic cysts provides a local environment for germ cells to proliferate and differentiate into spermatozoa. Compared to the amniote, anamniote Sertoli cells generally appear poorly developed throughout much of the spermatogenic cycle. Their cytoplasmic projections, which form cyst walls, are often very thin and contain little cytoplasm. However, as spermatogenic cysts grow and mature, Sertoli cells differentiate and upon spermiation generally contain abundant smooth and rough endoplasmic reticulum, lysosomes, and lipid droplets [9, 37]. An important Sertoli cell function during spermiation is phagocytosis of residual bodies or cytoplasmic remnants of spermatids [6].

Control of spermatogenesis is ultimately regulated by the hypothalamic–pituitary axis in both amniotes and anamniotes. The gonadotropins, luteinizing hormone (LH) and follicle-stimulating hormone (FSH), act on somatic cells of the testis to regulate steroid synthesis. Within testes, androgens produced by Leydig cells in the interstitium appear to be the driving force behind initiation and maintenance of spermatogenesis; the major androgen in most teleost fishes is 11-ketotestosterone [41–43]. Moreover, in the Japanese eel, 11-ketotestosterone has been shown to induce all stages of spermatogenesis in vitro [44]. Two distinct androgen receptors (ARs) with different affinities for testosterone and 11-ketotestosterone within the testis of fish have been recently reported [45–48]. The ARs are clearly expressed in the somatic cells of the testis [48]; in the rainbow trout testis, however, immunohistochemical analysis also indicates the presence of ARs in germ cells [49]. In the elasmobranch testis, Sertoli cells appear to have considerable steroidogenic capacity as evidenced by the levels of 3β-hydroxysteroid dehydrogenase enzyme detected in Sertoli cells during spermatogenesis; the highest levels of enzyme activity occurred during postmeiotic stages [50, 51]. Sourdaine and Garnier [52], using isolated lobules containing Sertoli cells associated with distinct germ cell stages, provided evidence for Sertoli cell steroidogenic capability throughout the process of spermatogenesis. Their results show that dogfish Sertoli cells synthesize and secrete increasing amounts of progesterone, androstenedione, testosterone, and 17α,20β-dihydroprogesterone, with highest activity localized in lobules containing Sertoli cells associated with spermatids. In the fish testis, receptors for both FSH and LH are present in Leydig cells and Sertoli cells [53]. In addition to the ability of FSH to stimulate steroidogenesis, a specific function for FSH in the fish Sertoli cell is yet to be determined. Recent work suggests that FSH may, directly or indirectly via 11-ketotestosterone, regulate the synthesis of growth factors by Sertoli cells [42]. Studies in Japanese eel (Anguilla japonica) and rainbow trout (Oncorhynchus mykiss) show that activin B and insulin-like growth factor produced by Sertoli cells stimulate spermatogonial proliferation [34, 43, 54, 55]. Additionally, in white-spotted char (Salvelinus leucomaenis), fibroblast growth factor (FGF) has been shown to play a role in regulation of spermatogenesis [56]. Transcripts for FGF and its receptor MFR1 have been localized to Sertoli cells in medaka, and it was suggested that both might play an important role in initiation and progression of spermatogenesis by regulation proliferation and differentiation of Sertoli cells [57].

Chapter 5 Sertoli Cell Biology in Fishes and Amphibians

Sertoli cells appear to play a similar physiological role in the regulation and support of spermatogenesis in amphibians. For example, Sertoli cells in newt testis have been shown to contain FSH receptor, and the actions of FSH on spermatogonial proliferation have been proposed to be mediated through activin B released by Sertoli cells [36, 58, 59]. In mammals, stem cell factor (SCF) released by Sertoli cells and acting via its receptor c-kit is well known to play an important role in survival, proliferation, and differentiation of germ cells (reviewed in Loveland and Schlatt [60]). A similar role for SCF in spermatogonial proliferation in the newt was recently demonstrated by Abe et al. [61]. Using a newt testis organ culture, recombinant human SCF was demonstrated to promote spermatogonial proliferation up to the start of meiosis [61]. Eel spermatogenesis–related substance 21 (eSRS21) was recently discovered in the Japanese eel testis [62], and it has some similarity to Müllerian-inhibiting substance and is expressed in the Sertoli cells of the immature eel testis. Organ culture studies have shown that eSRS21 prevents initiation of spermatogenesis, because it suppresses spermatogonial proliferation [62]. Furthermore, 11-ketotestosterone suppressed eSRS21 mRNA levels in vitro, suggesting a regulatory role for 11-ketotestosterone in eSRS21 production by the Sertoli cells. Interestingly, fish testes also have the potential to synthesize estradiol-17β (E2) [50, 63, 64]. Thus far, the cellular origin of E2 synthesis is only known in shark testes [50]. These authors [50] have described the presence of aromatase activity in the Sertoli cells; because the Sertoli cells are the main steroidogenic cells in elasmobranch testes, which is contrary to most other fishes, it is unclear if Sertoli cells are the general source of E2 synthesis in the testes of all fish. The biological action of E2 appears to be stimulation of spermatogonial stem cell renewal as seen in the Japanese eel [65] or the stimulation of spermatogonial proliferation as reported in the salmonid, the Japanese huchen [64]. The estrogen receptor has been identified in the Sertoli cells of the Japanese eel [65], whereas estrogen receptor transcripts have been reported in germ cells of channel catfish testes [66]. Recent studies using organ cultures in rainbow trout have demonstrated that E2 has proliferative effects on testicular interstitial cells mediated by an intracellular estrogen receptor [67 and unpublished data], suggesting that this sex steroid has a general mitogenic effect on various testicular cells. In anurans, mitogenic effects of E2 on spermatogonia appear to be mediated by the proto-oncogene fos protein (reviewed in [36]). This proliferative effect

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of E2 reportedly involves activation of mitogenactivated protein kinase [68]. However, it remains unclear whether the Sertoli cells are the source of E2 in the amphibian testes.

VI. SERTOLI CELL PROLIFERATION Contrary to mammals, wherein Sertoli cells do not proliferate in adults, spermatogenesis in fishes and amphibians is associated with Sertoli cell proliferation. Proliferating Sertoli cells have been reported during the early stages of the reproductive cycle regression in a variety of fish species [28, 69]. Sertoli cell proliferation is important for testicular growth and determines eventual sperm count [10, 70, 71]. Recently, De Franca et al. [10] performed quantitative morphometric analyses in tilapia testes and clearly demonstrated that Sertoli cell numbers increased up to the spermatocyte stage and then stabilized during postmeiotic stages. However, the mechanism behind Sertoli cell proliferation in fishes is unknown. It has been suggested that FSH may play a role in Sertoli cell proliferation as in mammals [62]. In addition, insulin-like growth factor 1 (IGF-1) has been shown to stimulate Sertoli cell proliferation in the elasmobranch dogfish (Squalus acanthias), and an IGF-1 receptor was detected in rainbow trout Sertoli cells [72, 73]. Similarly, in the newt (Cynops pyrrhogaster) FSH was shown to stimulate DNA synthesis in Sertoli cells and spermatogonia [74]. Recently, Matta et al. [75] demonstrated that nonsteroidogenic hormones may also play a role in Sertoli cell proliferation. They found that induced hypothyroidism using the goitrogen 6-n-propyl-2-thiouracil (PTU) resulted in an increase in Sertoli cell number in tilapia (Oreochromis niloticus), suggesting that thyroid hormones may regulate Sertoli cell proliferation. This is in agreement with studies in mammals, in which neonatal treatment of rats with PTU also resulted in an increase in the number of Sertoli cells in adult testes [76].

VII. ROLE OF SERTOLI CELLS IN INITIATION OF MEIOSIS After a species-specific number of mitotic divisions, spermatogonia enter meiosis and are referred to as primary spermatocytes; completion of the two meiotic divisions results in spermatid formation. Although, meiosis has been well described morphologically, very little is known about physiological–biochemical

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mechanisms that control the initiation or entrance of the germ cells into the meiotic process during anamniote spermatogenesis. Obviously, the intimate association of Sertoli cells with germ cells suggests a role for Sertoli cells in regulating this germ cell-specific event. Additionally, in more ancestral fish species like the agnathans (lamprey and hagfish) and elasmobranchs (sharks, rays, and chimeras), spermatogonia do not enter meiosis until after the formation of a basement membrane and formation of a spermatogenic cyst [4]; this set of observations is consistent with the conclusion that a specific local environment must be created in order for meiosis to occur. Although the role of Sertoli cells in controlling the entrance of spermatogonia into meiosis is unknown, some correlative data suggest that 17α,20β-dihydroxy4-pregnen-3-one, the spermiation-inducing steroid in fish, may also be involved in the progression from spermatogonia to spermatocytes [64, 77, 78]. This conclusion is based on the observation that plasma levels of this steroid are low during most of the reproductive cycle except for a slight increase at or before the onset of meiosis, and that in vitro production of 17α,20βdihydroxyprogesterone was highest at the onset of meiosis. Furthermore, in vitro treatment using a testis organ culture resulted in increased spermatogonial proliferation toward meiosis in the Japanese huchen. Finally, a progesterone receptor has been localized mainly to the Sertoli cells in the presence of spermatogonia [78], suggesting that 17α,20β-dihydroxyprogesterone via Sertoli cells may regulate the onset of meiosis. Studies have also implicated the IGF-1 system as a regulator of meiosis in fishes and amphibians. In the newt, FSH increases IGF-1 mRNA expression in Sertoli cells at the secondary spermatogonial stage [79]. Furthermore, IGF-1 has been localized in tilapia Sertoli cells and primary spermatocytes [80], and IGF-1 receptor is present in Sertoli cells and primary spermatocytes in rainbow trout [73]. These results suggest that IGF-1 may have both an autocrine and paracrine role before and during meiosis in the spermatogenic cyst. Based on information obtained in newts, a model has been proposed for the existence of a checkpoint for the initiation of meiosis based on the concentration ratio of FSH to prolactin [81]. These authors showed that FSH was required for the final mitotic division preceding entrance into meiosis. When FSH levels were low, this final division did not occur, instead spermatogonia underwent apoptosis. Apoptosis in the newt testis at this stage could be induced by prolactin [82, 83]. If the FSH-to-prolactin ratio was high, the final mitotic division took place and the cells entered meiosis, whereas in situations when the FSH-to-prolactin ratio

was low (e.g., at environmental cooler temperatures), the spermatogonia underwent apoptosis. Because the Sertoli cells contain FSH receptor, it appears that they play an important role in progression of spermatogonia into meiosis.

VIII. CONCLUDING REMARKS In summary, many of the regulatory functions of Sertoli cells on germ cell proliferation are similar in amniotes (reptiles, birds, mammals) and anamniotes (fishes, amphibians). FSH appears to be the primary endocrine signal from the pituitary, controlling proliferation and release of growth factors from the Sertoli cells. Several growth factors have also been identified in spermatogonial proliferation in amniotes and anamniotes; however, little is known about their function in controlling meiosis. It is unknown whether Sertoli cells secrete stimulatory or inhibitory factors acting on late spermatogonia, allowing these cells to enter meiosis as primary spermatocytes. The testicular organization in anamniotes (i.e., the spermatogenic cyst) provides an excellent model to study Sertoli cell interactions with specific stages of germ cell development. The isolation of spermatogenic cysts at specific stages during spermatogenesis and the identity of the transcriptome of the Sertoli cells at these specific stages will contribute to the identification of stage specific factors involved in regulating distinct phases of the spermatogenic cycle. Contrary to amniotes, Sertoli cells in most fishes and amphibians do not appear to be a permanent cell type within the testis. Sertoli cells proliferate, especially during the initial stages of spermatogenesis, and degenerate following spermiation. A continuous supply of Sertoli cells is therefore of great importance for maintaining reproductive capability in these species. Although several studies have investigated the origin of Sertoli cells, all conclusions drawn are based on ultrastructural observations. This is also true for the reported investigations on the developmental origins of Sertoli cells. The availability and use of Sertoli cell–specific markers would greatly enhance our ability to investigate these processes and the identity of the somatic precursor-Sertoli cells. Alternatively, because germ cells of transgenic rainbow trout containing the green fluorescent protein gene linked to the vasa promoter can be positively identified (reviewed in [84]), this current animal model may provide valuable additional information in the origin of the germ cells and their associated somatic cells during testicular recrudescence.

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41. Borg, B. (1994). Androgens in teleost fishes. Comp. Biochem. Physiol. 109C, 219–245. 42. Schulz, R. W., Vischer, H. F., Cavaco, J. E. B., Santos, E. M., Tyler, C. R., Goos, H. J. Th., and Bogerd, J. (2001). Gonadotropin, their receptors, and the regulation of testicular functions in fish. Comp. Biochem. Physiol. Part B 129, 407–417. 43. Schulz, R. W., and Miura, T. (2002). Spermatogenesis and its endocrine regulation. Fish Physiol. Biochem. 26, 43–56. 44. Miura, T., Yamauchi, K., Takahashi, H., and Nagahama, Y. (1991). Hormonal induction of all stages of spermatogenesis in vitro in the Japanese eel (Anguilla japonica). Proc. Natl. Acad. Sci. USA 88, 5774–5778. 45. Sperry, T. S., and Thomas, P. (1999). Characterization of two nuclear androgen receptors expressed in Atlantic croaker: Comparison of their biochemical properties and binding specificities. Endocrinology 140, 1602–1611. 46. Takeo, J., and Yamashita, S. (1999). Two distinct isoforms of cDNA encoding rainbow trout androgen receptors. J. Biol. Chem. 274, 5674–5680. 47. Takeo, J., and Yamashita, S. (2000). Rainbow trout androgen receptor-α fails to distinguish between any of the natural androgens tested in transactivation assays, not just 11-ketotestosterone and testosterone. Gen. Comp. Endocrinol. 117, 200–206. 48. Ikeuchi, T., Todo, T., Kobayashi, T., and Nagahama, Y. (2001). Two subtypes of androgen and progesterone receptors in fish testes. Comp. Biochem. Physiol. 129B, 449–455. 49. Takeo, J., and Yamashita, S. (2001). Immunohistochemical localization of rainbow trout androgen receptors in the testis. Fisheries Sci. 67, 518–523. 50. Dubois, W., and Callard, G. V. (1989). Role of the Sertoli cell in spermatogenesis: The Squalus testis model. Fish Physiol. Biochem. 7, 221–227. 51. Callard, G. V. (1992). Autocrine and paracrine role of steroids during spermatogenesis: Studies in Squalus acanthias and Necturus maculosus. J. Exp. Zool. 261, 132–142. 52. Sourdaine, P., and Garnier, D. H. (1993). Stage-dependent modulation of Sertoli cell steroid production in dogfish (Scyliorhinus canicula). J. Reprod. Fertil. 97, 133–142. 53. Yan, L., Swanson, P., and Dickhoff, W. W. (1992). A two-receptor model for salmon gonadotropins (GTH I and GTH II). Biol. Reprod. 47, 418–427. 54. Miura, T., Miura, C., Yamauchi, K., and Nagahama, Y. (1995). Human recombinant activin induces proliferation of spermatogonia in vitro in the Japanese eel Anguilla japonica. Fisheries Sci. 63, 434–437. 55. Loir, M. (1999). Spermatogonia of rainbow trout. II. In vitro study of the influence of pituitary hormones, growth factors and steroids on mitotic activity. Mol. Reprod. Dev. 53, 434–442. 56. Watanabe, A., and Onitake, K. (1995). Changes in the distribution of fibroblast growth factor in the teleostean testis during spermatogenesis. J. Exp. Zool. 272, 475–483. 57. Watanabe, A., Hatakeyama, N., Yasuoka, A., and Onitake, K. (1997). Distributions of fibroblast growth factor and the mRNA for its receptor, MFR1, in the developing testis of the medaka, Oryzias latipes. J. Exp. Zool. 279, 177–184. 58. Maekawa, K., Ji, Z.-S., and Abe, S-I. (1995). Proliferation of newt spermatogonia by mammalian FSH via Sertoli cells in vitro. J. Exp. Zool. 272, 363–373. 59. Ito, R., and Abe, S. I. (1999). FSH-initiated differentiation of newt spermatogonia to primary spermatocytes in germ-somatic cell reaggregates cultured within a collagen matrix. Int. J. Dev. Biol. 43, 111–116. 60. Loveland, K. L., and Schlatt, S. (1997). Stem cell factor and c-kit in the mammalian testis: Lessons originating from Mother Nature’s gene knockouts. J. Endocrinol. 153, 337–344.

61. Abe, K., Jin, Y., Yamamoto, T., and Abe, S.-I. (2002). Human recombinant stem cell factor promotes spermatogonial proliferation, but not meiosis initiation in organ culture of newt testis fragments. Biochem. Biophys. Res. Commun. 294, 695–699. 62. Miura, T., Miura, C., Konda, Y., and Yamauchi, K. (2002). Spermatogenesis-preventing substance in Japanese eel. Development 129, 2689–2697. 63. Fostier, A., Le Gac, F., and Loir, M. (1987). Steroids in male reproduction. In “Proceedings of the Third International Symposium on the Reproductive Physiology of Fish” (D. R. Idler, L. W. Crim, and J. M. Walsh, eds.), pp. 239–245. St. John’s, Newfoundland, Canada: Marine Sciences Research Laboratory, Memorial University of Newfoundland. 64. Amer, M. A., Miura, T., Miura, C., and Yamauchi, K. (2001). Involvement of sex steroid hormones in the early stages of spermatogenesis in Japanese huchen (Hucho perryi). Biol. Reprod. 65, 1057–1066. 65. Miura, T., Miura C., Ohta, T., Nader, M. R., Todo, T., and Yamauchi, K. (1999). Estradiol-17β stimulates the renewal of spermatogonial stem cells in males. Biochem. Biophys. Res. Commun. 264, 230–234. 66. Wu, C., Patino, R., Davis, K. B., and Chang, X. (2001). Localization of estrogen receptor α and β RNA in germinal and nongerminal epithelia of the channel catfish testis. Gen. Comp. Endocrinol. 124, 12–20. 67. Bouma, J., Cloud, J. G., and Nagler, J. J. (2003). In vitro effects of estradiol-17β on myoid and fibroblastic cell proliferation in the immature rainbow trout testis. Fish Physiol. Biochem. 28, 191–192. 68. Chieffi, P., D’Amato, L. C., Guarino, F., Salvatore, G., and Angelini, F. (2002). 17β-Estradiol induces spermatogonial proliferation through mitogen-activated protein kinase (extracellular signal-regulated kinase 1/2) activity in the lizard (Podarcis s.sicula). Mol. Reprod. Dev. 61, 218–225. 69. Brown-Peterson, N. J., Grier, H. J., and Overstreet, R. M. (2002). Annual changes in germinal epithelium determine male reproductive classes of the cobia. J. Fish Biol. 60, 178–202. 70. Orth, J. M., Gunsalus, G. L., and Lamperti, A. A. (1988). Evidence from Sertoli cell-depleted rats indicates that spermatid number in adults depends on numbers of Sertoli cells produced during perinatal development. Endocrinology 122, 787–794. 71. De Franca, L. R., Hess, R. A., Cooke, P. S., and Russell, L. D. (1995). Neonatal hypothyroidism causes delayed Sertoli cell maturation in rats treated with propylthiouracil: Evidence that the Sertoli cell controls testis growth. Anat. Rec. 242, 7–69. 72. Dubois, W., and Callard, G. V. (1993). Culture of intact Sertoli/germ cell units and isolated Sertoli cells from Squalus testis. II. Stimulatory effects of insulin and IGF-1 on DNA synthesis in premeiotic stages. J. Exp. Zool. 267, 233–244. 73. Le Gac, F., Loir, M., Le Bail, P.-Y., and Ollitrault, M. (1996). Insulin-like growth factor (IGF-1) mRNA and IGF-1 receptor in trout testis and in isolated spermatogenic and Sertoli cells. Mol. Reprod. Dev. 44, 23–35. 74. Ji, Z.-S., and Abe, S.-I. (1994). Mammalian follicle-stimulating hormone stimulates DNA synthesis in secondary spermatogonia and Sertoli cells in organ culture of testes fragments from the newt, Cynops pyrrhogaster. Zygote 2, 53–61. 75. Matta, S. L. P., Vilela, D. A. R., Godinho, H. P., and Franca, L. R. (2002). The goitrogen 6-n-propyl-2-thiouracil (PTU) given during testis development increases Sertoli and germ cell numbers per cyst in fish: The tilapia (Oreochromis niloticus) model. Endocrinology 143, 970–978. 76. Hess, R. A., Cooke, P. S., Bunick, D., and Kirby, J. D. (1993). Adult testicular enlargement induced by neonatal hypothyroidism is

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accompanied by increased Sertoli and germ cell numbers. Endocrinology 132, 2607–2613. Depeche, J., and Sire, O. (1982). In vitro metabolism of progesterone and 17α-hydroxyprogesterone in the testis of the rainbow trout, Salmo gairdneri Rich., at different stages of spermatogenesis. Reprod. Nutr. Dev. 22, 427–438. Miura, T., Amer, M. A., Miura, C., Higashino, T., Ozaki, Y., and Konda, Y. (2003). Involvement of 17α,20α-dihydroxy-4-pregnen3-one on spermatogenesis in Japanese eel (Anguilla japonica) and Japanese huchen (Hucho peryii). 7th International Symposium on Reproductive Physiology of Fish, Mie, Japan, Abstract P-IV-8. Yamamoto, T., Nakayama, Y., and Abe, S.-I. (2001). Mammalian follicle-stimulating hormone and insulin-like growth factor I (IGF-I) up-regulate IGF-I gene expression in organ culture of newt testis. Mol. Reprod. Dev. 60, 56–64. Reinecke, M., Schmid, A., Ermatinger, R., and Loffing-Cueni, D. (1997). Insulin-like growth factor I in the teleost Oreochromis

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mossambicus, the tilapia: Gene sequence, tissue expression, and cellular localization. Endocrinology 138, 3613–3619. Yazawa, T., Yamamoto, T., Jin, Y., and Abe, S.-I. (2002). Folliclestimulating hormone is indispensable for the last spermatogonial mitosis preceding meiosis initiation in newts (Cynops pyrrhogaster). Biol. Reprod. 66, 14–20. Yazawa, T., Yamamoto, T., and Abe, S. (2000). Prolactin induces apoptosis in the penultimate spermatogonial stage of the testes in Japanese red-bellied newt (Cynops pyrrhogaster). Endocrinology 141, 2027–2032. Yazawa, T., Fujimoto, K., Yamamoto, T., and Abe, S.-I. (2001). Caspase activity in newt spermatogonial apoptosis induced by prolactin and cycloheximide. Mol. Reprod. Dev. 59, 209–214. Yoshizaki, G., Takeuchi, Y., Kobayashi, T., Ihara, S., and Takeuchi, T. (2002). Primordial germ cells: The blueprint for a piscine life. Fish Physiol. Biochem. 26, 3–12.

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C H A P T E R

6 Sertoli Cell Biology in Seasonal Breeders AMIYA P. SINHA HIKIM

ANDRZEJ BARTKE

Department of Medicine, Harbor-UCLA Medical Center and Research and Education Institute, UCLA School of Medicine, Torrance, California

Geriatrics Research, Department of Medicine, Southern Illinois University School of Medicine, Springfield, Illinois.

the basis for temporal dissociation of insemination and fertilization. In mammals inhabiting temperate and circumpolar regions, arrival of the young in the spring and early part of the summer is of obvious adaptive significance and is ensured by appropriate timing of the breeding season. Thus, in species with short gestation periods, the endocrine and gametogenic activities of the testes are either restricted to a specific period in the spring or exhibit annual fluctuations, with plasma androgen levels, testicular weight, and sperm production reaching maximal values during spring months. For example, different species of North American and Eurasian voles (Microtus, Clethrionomys) and wild mice (Peromyscus, Apodemus) breed primarily in spring and summer, with winter breeding generally restricted to a limited number of individuals in years with an exceptional abundance of food. An extreme example of seasonal breeding is provided by the brown marsupial mouse (Antechinus stuartii), which always breeds only once in each year in the seasonal forests of southeastern Australia, producing a single litters of 6 to 10 young [4]. In contrast, the house mouse (Mus musculus), progenitor of the various stocks of laboratory mice, which is commonly found in food storage facilities and human dwellings, is an opportunistic breeder with capacity to reproduce throughout the year if sufficient food resources are available. Carnivores with a somewhat longer gestation period achieve similar

I. SEASONALITY OF MAMMALIAN REPRODUCTION II. PHOTOPERIODIC CONTROL OF SEASONAL REPRODUCTION III. PHOTOPERIODIC CONTROL OF THE HYPOTHALAMIC–PITUITARY–GONADAL AXIS IV. STRUCTURAL RESPONSE OF THE SERTOLI CELL AT DIFFERENT STATES OF GONADAL ACTIVITY V. SUMMARY References

I. SEASONALITY OF MAMMALIAN REPRODUCTION Most mammalian species are seasonal breeders, exhibiting seasonal changes of their reproductive abilities with transitions from fertility to sterility timed to optimize the chances for survival of the offspring [1–3]. Seasonal breeding, thus, ensures that reproduction will occur in harmony with the environmental variations. Synchronizing energetically demanding biological functions to the most favorable time of the year enables individual species to cope with recurring stressors such as seasonal declines in food or water availability, annual fluctuations in ambient temperature, or annual changes in the day length (photoperiod). Reproductive competence of males and females also typically coincides in time, except when special mechanisms, such as longterm sperm storage in the female genital tract, provide SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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timing of births either by mating early in the spring or by mating in the fall with the fertilized eggs implanting only after a considerable delay. For example, the black bear (Ursus americanus) is a long-day breeder with a mating season extending from May to July [5]. The testes exhibit active spermatogenesis from mid-May until late July but regress in autumn before denning and begin to recrudesce during mid- to late denning [6]. The seasonal changes in testis size parallel the seasonal changes in serum testosterone concentrations [6]. Many ruminants, including white-tailed deer (Odocoileus virginianus), red deer (Cervus elaphus), reindeer (Rangifer tarandus), wild sheep (Ovis aries), and various breeds of domestic sheep, have longer periods of gestation (approximately 5 months in the sheep) and breed in the fall. In the horse (Equus caballus), the length of gestation is approximately 11 months and breeding season is in the spring. The primary environmental cue for stimulating gonadal activity and reproductive behavior during the appropriate time of the year for the majority of animals is the seasonal changes in day length [3, 7]. Supplementary cues such as temperature, food and water availability, and social factors from members of the same species (pheromones and vomeronasal mediation supplemented with tactile cues), through finetuning of the initial, general reproductive response to photoperiod, can also affect seasonal reproduction.

II. PHOTOPERIODIC CONTROL OF SEASONAL REPRODUCTION It is well known that regardless of whether seasonal breeding is required because of energetic, nutritional, or climatic variation, a mammal may opt to use a predictor of such variation. Photoperiod is the most widely used of such predictors, at least in the temperate and subpolar regions. Mechanisms responsible for photoperiodic control of testicular function have been studied in considerable details in several mammalian species, including the well-studied golden (Syrian) hamster [8–10] and the Siberian (Djungarian) hamster. In the male golden (Syrian) hamster (Mesocricetus auratus) during fall of the year or upon introduction to a short photoperiod, endocrine, structural, behavioral, and fertility parameters of reproductive function decline and remain at low levels until late winter or until reintroduction to a long photoperiod, when they regain their former status (recrudescence) in preparation for the spring and summer breeding season (Fig. 6.1). This seasonal reproductive rhythm can be elicited at any time of the year by artificial alteration in photoperiod [11]. Exposure of adult male hamsters to a short photoperiod (<12.5 hr of light per day) produces testicular atrophy [8, 12]. Reproductive function, however,

FIGURE 6.1 Cyclical changes in endocrine, testis weight, sperm production, copulatory behavior, and fertility in adult male golden hamsters exposed to varying natural photoperiod regimens. (Reprinted with permission from Sinha Hikim et al. [54].)

undergoes spontaneous recovery (recrudescence) after ~25 wk or can be restored much earlier by returning the animals to a long photoperiod [10, 13, 14]. The cyclic behavior of this species offers a physiological model system for “turning on” and “turning off” spermatogenesis and is used for studying the structure–function relationships of various testicular cell types, including the Sertoli cells [15–17]. Another very popular animal model for the study of seasonality of reproduction in mammals is the Siberian hamster (Phodopus sungorus). The Siberian hamster occupies vast regions of central Asia and matures rapidly when born early in the season under conditions of increasing photoperiod, but experiences a long delay in reproductive development if born later in the season when the photoperiod is decreasing [18]. Under laboratory conditions, transfer of adult Siberian hamsters from a long photoperiod consisting of 16 hr of light and 8 hr of darkness (16L:8D) to a short photoperiod leads to testicular atrophy followed by spontaneous gonadal recrudescence ~20 to 25 wk after the transfer [9, 19, 20]. Of note, naturally occurring or experimentally induced suppression of gonadal activity in this species is accompanied by a conspicuous change from a brownishgray summer coat to near white winter pelage [18, 21].

III. PHOTOPERIODIC CONTROL OF THE HYPOTHALAMIC–PITUITARY– GONADAL AXIS Spectacular progress has been made in our understanding of the photoperiodic control of the hypothalamic–pituitary–gonadal (HPG) axis in several

Chapter 6 Sertoli Cell Biology in Seasonal Breeders

mammalian species, including the Syrian (golden) and the Siberian (Djungarian) hamsters [22, 23]. Seasonal changes in day length impinge on the neuroendocrine axis via a retinal–hypothalamic–pineal pathway [24]. In brief, light entrains circadian oscillators in the suprachiasmatic nucleus (SCN), which via a polysynaptic pathway controls melatonin synthesis in the pineal gland. Circadian output from the SCN is transmitted to the parvocellular autonomic component of the hypothalamic paraventricular nucleus (PVN), which projects to the upper thoracic intermediolateral cell column via the medial forebrain bundle (MFB). MFB neurons innervate the superior cervical ganglion (SCG), which project β-noradrenergic sympathetic fibers to pinealocytes [24]. Information that originates in the SCN drives a circadian rhythm of synthesis of N-acetyltransferase (NAT), the rate-limiting enzyme in the conversion of N-acetyl-serotonin to melatonin. Consistent with the essential role of melatonin in mammalian photoperiodism, surgical excision of the pineal gland was found to be associated with a reduction in the rate and magnitude of seasonal gonadal regression in male Syrian hamsters [25]. Furthermore, lesions in any given component of the neural pathway that controls the circadian rhythm in pineal melatonin secretion eliminated the reproductive response to day length in hamsters [26–28]. Thus, the resulting changes in the daily pattern of melatonin secretion by the pineal gland alter the release of hypothalamic neurotransmitters, including gonadotropin-releasing hormone (GnRH) and dopamine, into the blood supply of the anterior pituitary. This affects the release of hormones, such as follicle-stimulating hormone (FSH), luteinizing hormone (LH), and prolactin (PRL), that are responsible for the control of gonadal activity. It is believed that of particular importance in the chain of events are the changes in the pulsatile pattern of GnRH release with stimulation of the “GnRH pulse generator,” leading to stimulation of the gonads, reproductive development or recrudescence and fertility, and inhibition of the GnRH pulse generator, leading to gonadal atrophy and reproductive quiescence [29–31]. It is well known that the pituitary gonadotropins, LH in particular, control the production of testosterone (T) by Leydig cells. This androgen is of prime importance in supporting spermatogenesis in adult mammals [32]. The role of FSH in the control of adult spermatogenesis in animals, such as the rat, is not well understood [32], but this gonadotropin is believed to be importantly involved in the seasonal activation of spermatogenesis in both golden and Djungarian hamsters [9, 33–37]. There is general agreement in the literature that Sertoli cells contain the receptors for both FSH and T and are the main targets of these hormones in the seminiferous epithelium [32, 38].

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In both male golden and Djungarian hamsters, exposure to short days results in a striking inhibition of testicular activity manifested by a marked decrease in serum levels of FSH, LH, PRL, and T and in the number of FSH, LH, and PRL receptors [9, 35, 39–41]. Concomitant reduction in circulating levels of T is believed to reflect suppression of pituitary LH secretion combined with loss of LH receptors due to hypoprolactinemia [42] and reduced ability of Leydig cells to produce T from nonsteroidal and steroidal precursors [33, 43]. Available evidence also indicates that exposure of the golden hamsters to an inhibitory photoperiod (6 hr of light, 18 hr of darkness) is associated with an increase in the concentration (femtomoles per milligram of protein), but a reduction in the total content (femtomoles per testis) of FSH receptors [35, 40, 44]. It has been suggested that the opposite changes in these measurements of FSH binding are related at least in part to changes in the cellular composition of the testes between active and inactive gonadal states, namely, a decrease in germ cell number and Sertoli cell size and an increase in the proportion of testicular mass contributed by the Sertoli cells [40, 44]. Subsequent morphometric analyses revealed that these seemingly contradictory changes in the concentration and total content of FSH receptors are related to the changes in the basal compartment surface areas of Sertoli cells, where most of the FSH receptors are presumed to be located [35]. For example, the decrease in the surface area of the basal compartment of Sertoli cells per testis during testicular involution (48.7%) was very similar to the changes in the total content (femtomoles per testis) of FSH receptors (55.7% reduction). Likewise, the observed increase in the surface area of the basal compartment of Sertoli cell per gram of testis paralleled the increase in the concentration of FSH receptors (femtomoles per milligram of protein) during testicular regression [35]. Consistent with this observation, the surface areas of the basal compartment of the Sertoli cells per gram of testis and per testis are significantly and positively correlated with the concentration (r = 0.79; P < 0.005) and the content (r = 0.77; P < 0.005) of FSH receptors [35]. Thus, it is reasonable to conclude that suppression of spermatogenesis at the end of breeding season is due to lack of adequate support of the Sertoli cells. Declining levels of circulating FSH and T would thus fail to provide the optimum stimulation of the Sertoli cells to maintain normal spermatogenesis. It is also well established that T, together with FSH, is a major regulator of germ cell survival, and the Sertoli cells are the main target of these hormones in the seminiferous epithelium [32, 38, 45, 46]. Indeed, studies have indicated that levels of spontaneous apoptosis (during active spermatogenesis) are increased as a result of

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seasonal declines in hormonal stimulation, and that this escalation in programmed germ cell death is an important determinant of inhibitory photoperiodinduced gonadal regression [47]. Further support for this conclusion stems from the findings that treatment with pregnant mare serum gonadotropin (PMSG), a hormone with predominantly FSH activity, followed by human chorionic gonadotropin (hCG), which has predominantly LH activity, can reinitiate spermatogenesis in a seasonal breeder, the woodchuck (Marmota monax), during gonadal regression [48]. It may be relevant in this connection to note that in golden hamsters with gonadal atrophy produced by exposure to a short photoperiod, spermatogenesis can be stimulated and fertility restored by treatment with an ectopic pituitary transplant that acts by releasing PRL and by stimulating gonadotropin release [49]. Seasonal changes in spermatogenic activity and, by implication, Sertoli cell function also underlie the endocrine changes during return of testicular activity from reproductive quiescence (recrudescence). For example, stimulation of testicular growth and spermatogenic activity that occurs either spontaneously or in response to experimental manipulation of photoperiod is also preceded by increases in circulating levels of gonadotropins and T with the increase in FSH being among the earliest detectable changes [14, 50]. Transfer of gonadally regressed adult golden hamster or pubertally delayed juvenile Siberian hamsters from a short (inhibitory) to a long (stimulatory) photoperiod can lead to stimulation of FSH release as early as within 3–5 days [16, 51–53]. Such photostimulated increases in FSH secretion reflect selective increases in pituitary FSH β and α-subunit mRNA levels [53]. Neither serum LH nor LH β mRNA levels changed significantly following long-day transfer. Taken together, these results suggest that long-day-associated increases in circulating levels of FSH reflect an underlying increase in pituitary FSH β and α mRNA accumulation. Likewise, the only hormone that reduced significantly during early regression after 4–6 wk of exposure to shortphotoperiod in golden hamsters was FSH [37]. The levels of LH and the T were not significantly altered. Thus, it can be concluded that the Sertoli cells of seasonally breeding mammals are normally subjected to changing hormonal environments, and that at different times of the year they are in very different functional states. Because cell function is often reflected in the structure of a cell, it is important to characterize the structure of the Sertoli cells at different functional states (active, early regressed, regressed, and early recrudesced) in order to develop a more complete

understanding of its structure–function relationships. In the golden (Syrian) hamster model, gradual changes in the Sertoli cell functional activity can be elicited at any time of the year by appropriate alterations of photoperiod and exposure time [35–37].

IV. STRUCTURAL RESPONSE OF THE SERTOLI CELL AT DIFFERENT STATES OF GONADAL ACTIVITY Among various seasonal breeders, the golden hamster is the only species studied so far that provides quantitative information on structural changes in the Sertoli cells at various states of gonadal activity [35–37, 54, 55]. Gradual changes in the Sertoli cell functional activity (active, early regressed, regressed, and early recrudesced) in this species can be elicited at any time of the year by appropriate alterations of photoperiod and exposure time.

A. Testis Size, Tubular Morphology, and Germ Cell Numbers The testes of gonadally active hamsters displayed active spermatogenesis similar to that seen in a nonseasonal breeder (e.g., laboratory rat or mouse). The seminiferous tubules contained Sertoli cells, spermatogonia, spermatocytes, and spermatids in different phases of development (Fig. 6.2, left panel). In contrast, testes of the hamsters exposed to short photoperiod exhibited a 90% decrease in testicular weight in comparison to gonadally active animals [55]. Seminiferous tubules contained primarily Sertoli cells and spermatogonia with occasional spermatocytes and spermatids (Fig. 6.2, right panel). From a structural standpoint, one would expect to see striking changes in hamster Sertoli cell structure during gonadal quiescence [54]. Indeed in our studies, we found a marked reduction in cell height and major organelle content, absence of lateral processes, and the presence of disorganized apical processes (Fig. 6.3). To determine the workload of the Sertoli cells, the number of germ cells (preleptotene and pachytene spermatocytes, and step 7 spermatids) supported by an individual Sertoli cell was determined in active and regressed testis. Sertoli cells in active states of spermatogenesis supported, on the average, 2.27 preleptotene spermatocytes, 2.46 pachytene spermatocytes, and 8.17 round spermatids, whereas the corresponding germ cell numbers supported in the regressed testes were 0.96, 0.20, and 0.04, respectively [55].

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FIGURE 6.2 Differences in the appearance of stage VII tubules between active (left panel) and inactive (right panel) states of spermatogenesis. Left panel: A stage VII tubule from a gonadally active hamster shows active spermatogenesis at all levels and contains spermatids soon to be released into the tubular lumen. Right panel: Seminiferous tubules from a gonadally regressed hamster show severe impairment of spermatogenesis. Sertoli cells (S) and spermatogonia (sg) are the main constituents of these tubules. Note also the complete obliteration of the tubular lumen. (Reprinted with permission from Sinha Hikim et al. [54].)

Two different but complementary approaches were also employed to document the early structural changes of the Sertoli cells in gonadal regression and in recrudescence [37]. Early gonadal regression was achieved by transferring gonadally active hamsters to a short photoperiod for 4–6 wk, whereas early recrudescence was achieved by transferring gonadally regressed hamsters to a stimulatory (long) photoperiod for 1–2 wk. During early regression, characterized by the presence of a small number of apoptotic germ cells exclusively at stage VII tubules (Figs. 6.4 and 6.5), there was a modest but significant (P < 0.05) decrease in the number of preleptotene and pachytene spermatocytes and step 7 spermatids supported by the Sertoli cells in comparison to gonadally active animals. No apparent changes in Sertoli cell structure were evident between the active and early regressed states (Fig. 6.4). During early recrudescence, testis size and tubule volume were increased [37]. The mean number of preleptotene spermatocytes per Sertoli cell was significantly (P < 0.02) increased in hamsters following photostimulation when compared to fully regressed animals (1.45 ± 0.15 vs. 0.96 ± 0.07), whereas no significant differences were detected in the relative number of pachytene spermatocytes and round spermatids between these two groups.

B. Sertoli Cell Numbers, Cell Size, and Plasma Membrane Surface Areas An average testis of gonadally active hamsters contained about 74 million Sertoli cells; the corresponding values in the early regressed, regressed, and recrudesced testes were about 67, 80, and 66 million, respectively [37, 55]. These values were not significantly different from each other, thus suggesting the numerical stability of the Sertoli cells at various states of gonadal activities. As shown in Figures 6.6 and 6.7, testicular regression in the golden hamster was accompanied by a marked decrease in the Sertoli cell size and the plasma membrane surface area. The mean volume of Sertoli cells in active state of spermatogenesis was 4,233 μm3, and their surface areas averaged 13,273 μm2. If a Sertoli cell of 4233 μm3 was spherical, it would be approximately 20 μm in diameter and would possess a membrane surface area of about 1256 μm2, as calculated by using the standard stereological equations V = 1/6 D3; S = π D2, where V is the mean volume of an individual Sertoli cell, S is the mean cellular surface area, and D is the diameter of the cell. However, the actual surface of an active Sertoli cell was 10.5 times greater. This high surface-to-volume ratio is reflective of the extremely irregular shape and extensive surface processes of these cells in the gonadally active state.

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The average volume of a Sertoli cell in the regressed testis was 1168 μm3 and the surface area of the entire cell was only 2152 μm2. Thus, the regressed hamster Sertoli cell had a volume of only 27.6% of its former size, primarily due to decrease in its cytoplasmic volume, and a surface area of only 16.2% of the surface of an active Sertoli cell. No significant changes in Sertoli cell size (3492 vs. 4233 μm3) or its plasma membrane surface area (9036 vs. 13,274 μm3) were noted during early testicular regression in comparison to the values measured in gonadally active animals. In contrast, transfer of gonadally regressed hamsters to a stimulatory photoperiod for 1 or 2 wk, led to a significant (P < 0.05) increase in both Sertoli cell size (2301 vs. 1169 μm3) and its surface area (4669 vs. 2153 μm3). The changes in the membrane surface area during gonadal quiescence and during early recrudescence most likely are responsive to changes in the germ cell population.

C. Structural Characteristics of Sertoli Cells during Various States of Spermatogenesis

FIGURE 6.3 Electron micrographs showing the characteristic features of the Sertoli cell in the regressed hamster testis. Upper panel: Basal aspect of these Sertoli cells shows sparse perinuclear cytoplasm, abundant secondary lysosomes formed from the degradation of germinal cells and lipids, and complete absence of lateral processes. A variety of other cellular organelles, including Golgi (G), mitochondria (M), and smooth endoplasmic reticulum (SER) are also indicated. Lower panel: Apical aspect of the Sertoli cells shows many disorganized apical processes. (Reprinted with permission from Sinha Hikim et al. [54].)

Stereological data obtained from hamster Sertoli cells at various phases of gonadal activities are summarized in Figures 6.6 and 6.7. The smooth endoplasmic reticulum (SER) was the most abundant organelle, occupying a volume of 286.9 μm3, and possessed a mean membrane surface area of 21,132 μm2. The rough endoplasmic reticulum (RER) occupied a volume of 46 μm3 in an average Sertoli cell cytoplasm, encompassed by an average surface area of about 5926 μm2, which was approximately one-quarter of the surface area

FIGURE 6.4 Typical appearance of the hamster Sertoli cells at stage VII tubules during early gonadal regression, achieved by transferring gonadally active hamsters to an inhibitory (short) photoperiod for 4 wk. Note the presence of an apoptotic step 7 spermatid (left panel) and a pachytene spermatocyte (right panel) among many nonapoptotic germ cells. The Sertoli cell appears structurally normal.

Chapter 6 Sertoli Cell Biology in Seasonal Breeders

FIGURE 6.5 Basal portion of a stage VII tubule from a hamster that had been exposed to a short photoperiod for 4 wk shows an apoptotic step 19 spermatid deep within the Sertoli cell cytoplasm. The mature spermatid is far removed from its normal position lining the tubular lumen and will eventually be phagocytosed by the Sertoli cells.

of the SER. The mitochondria occupied a volume of 203.4 μm3 in the cytoplasm of an average cell. Outer and inner membrane surface areas of mitochondria were 4180 and 6157 μm2, respectively. As the volume of the Sertoli cell decreased during photoperiod-induced gonadal regression, there was also a significant reduction in the absolute volumes of nearly all of its major subcellular organelles (Fig. 6.6), including mitochondria (73.4% reduction), SER (77.6%), RER (84.3%), and Golgi complex (74.3%). The absolute volumes of lipids and lysosome, however, were not significantly different. A significant decline in the membrane surface area of various subcellular organelles in inactive Sertoli cells (Fig. 6.7) was as follows: outer (84.3%) and inner (82.8%) mitochondrial membrane, SER (70.0%), and RER (88.4%). The Golgi surface area was reduced by 65.6%. The observed changes in the majority of the structural parameters of the Sertoli cells accompanying

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short-photoperiod–induced testicular regression were significantly and positively correlated with reductions in the levels of plasma gonadotropins and T, intratesticular T, and in the total content of FSH receptors in the testis [35]. Thus, the observed morphological manifestations of changes in the Sertoli cells during testicular regression appear to be the expected consequences of the reduced gonadotropin secretion and consequent decrease in T secretion during this period. Collectively, these results indicate that with the exception of the lysosomal and lipid compartments, which possibly reflect maintained or increased “phagocytic” activities, cellular inactivity was associated with an overall reduction in organelle volume and surface area. That virtually every measured structural parameter is substantially changed after loss of cellular activity indicates a general rather than specific shutdown of Sertoli cell function during testicular regression. However, note that in this study only gonadally active (similar to peak breeding) and maximally regressed (similar to nonbreeding) animals were used with no examination of interim time periods. It could be expected that studies during early gonadal regression and recrudescence may reveal differential susceptibility of Sertoli cell organelles to changes in the hormonal environment. Results of such studies are discussed later. A detailed account of the quantitative structural changes in the Sertoli cells during early testicular regression and recrudescence induced by alterations of photoperiod, so far, is only available for the golden hamsters [37]. No changes in the absolute volumes of nearly all of the major Sertoli cell organelles were noted during early gonadal regression, defined by the presence of a small number of apoptotic germ cells at stage VII tubules in animals transferred from a long to a short photoperiod (Figs. 6.4 and 6.5). No changes in the organelle surface areas, except for outer and inner mitochondrial membranes that showed a significant decline by 35%, were apparent during early regression. It is pertinent to note here that during early gonadal regression when numerous testicular parameters, including the number of germ cells supported by an individual Sertoli cell [37], the Leydig cell size, and the absolute volumes and surface areas of its organelles associated with steroid biosynthesis [41] had already declined, Sertoli cells showed no structural alterations. Of the endocrine changes, only plasma FSH levels had declined at this time and there were virtually no correlations between plasma FSH levels and the various structural parameters of the Sertoli cells during early testicular regression [37]. The general lack of Sertoli cell changes in response to a short-term exposure to inhibitory photoperiod is similar to the early response to hypophysectomy in the nonseasonal rat [56]. Transfer of gonadally regressed

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A

B

FIGURE 6.6 Summary of changes in absolute volume of Sertoli cells and their organelles in hamsters during active, early regressed, regressed, and early recrudesced states of the spermatogenic activity; increase in the volume of cell and SER (A) and RER (B) during recrudescence.

hamsters to a stimulatory photoperiod for 1 to 2 wk (early recrudescence) resulted in a significant increase in the volume of SER (Fig. 6.6A) and RER (Fig. 6.6B) and in the surface areas of outer and inner mitochondrial membranes, SER, and RER (Fig. 6.7). Unlike early regression, plasma levels of both FSH and T increased significantly during early testicular recrudescence. However, only the RER (both volume and surface area) was correlated with the hormonal changes during early recrudescence, suggesting that the initiation of the Sertoli cell synthetic processes was important in reestablishment of spermatogenesis. From these tests of correlation, one can reasonably conclude that there is a

causal relationship, direct or indirect, between hormone levels and the changes in the structural components of the Sertoli cell. Taken together, these results do indicate that the structural responses of the Sertoli cells during early regression and during early recrudescence do not involve the same organelles. Munoz et al. [57] also studied the changes in the Sertoli cell ultrastructure of another seasonally breeding rodent, the viscacha (Lagostomus maximus maximus), during gonadally active, regressed, and recrudesced states. During gonadal regression, which takes place during the short days of winter, the viscacha Sertoli cells exhibited a marked decrease in their size and in

Chapter 6 Sertoli Cell Biology in Seasonal Breeders

FIGURE 6.7 Summary of changes in the membrane surface areas of Sertoli cells and their organelles in hamsters during active, early regressed, regressed, and early recrudesced states of the spermatogenic activity.

the volume densities of the ER, mitochondria, and Golgi. There was, however, an accumulation of lipid and lysosomes. The volume densities of the ER, mitochondria, and Golgi were increased during gonadal recrudescence in spring. Gonadal regression in this species was accompanied by a marked decrease in the circulating levels of LH and T, and in the concentrations of LH, FSH, and PRL receptors in the testis [58]. Testicular restoration during spring is followed by a period of maximum activity during the long days of summer and autumn [58–60]. Available data also suggest that photoperiod synchronizes the viscacha reproductive rhythmicity through the pineal gland [59, 61, 62]. Thus, the observed morphological manifestation of the viscacha Sertoli cells during the annual reproductive

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cycles appears to be the expected consequence of the seasonal fluctuations of the gonadotropins and T. Lohiya et al. [63] examined multiple parameters of testicular function in langur monkeys (Presbytis entellus) throughout the year and found no evidence of seasonal fluctuations in sperm characteristics, seminiferous tubule diameter, or diameter of Sertoli cell nuclei. They concluded that males of this species lack reproductive seasonality. Sanford et al. [64, 65] examined seasonal fluctuations in serum levels of inhibin, an important product of Sertoli cells, in the ram. Animals exposed to alternating 4-month short (stimulatory) and long (inhibitory) photoperiods experienced fluctuations in testicular weight that resembled normal seasonal changes. In these animals inhibin levels increased in response to short photoperiods and were positively correlated with plasma FSH levels. Increases in the level of inhibin followed increases in the levels of FSH. Maximal inhibin levels were observed during the first or the second month of exposure to short photoperiods, i.e., during recrudescence of the testes, and these levels coincided with maximal increase in testicular content of FSH receptors. The number of FSH receptors was positively correlated with the levels of inhibin (r = 0.65). We have used Siberian hamsters to examine the effects of photoperiod on testicular levels of mRNA for two Sertoli cell products, inhibin α and androgen binding protein (ABP) [66]. In animals raised in short photoperiods, FSH was not detectable in the plasma and the testes remained very small, but expression of both inhibin α and ABP was clearly detectable. The levels of the corresponding mRNAs (normalized against 18S ribosomal RNA) were higher in the regressed testes of animals exposed to short photoperiods than in the active testes of animals exposed to long photoperiods. This may have been related to the absence of advanced germ cells and the relative enrichment of Sertoli cells in the testes of short-photoperiod– exposed hamsters. Majumdar et al. [67, 68] examined the effects of exposing Siberian hamsters to different photoperiods on the in vitro function of Sertoli cells isolated from these animals. There were no obvious effects of photoperiod history on the lactate production by Sertoli cells isolated from juvenile hamsters. Sertoli cells from both long- and short-photoperiod–exposed animals responded to FSH but not to vitamin D3 by increased lactate production. In both groups vitamin D3 potentiated the effects of FSH. However, functional differences were detected between Sertoli cells isolated from adult long-photoperiod–exposed gonadally active hamsters as compared to the cells isolated from adult short-photoperiod–exposed spermatogenically

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suppressed animals. Two-dimensional PAGE analysis of proteins secreted by these cells revealed greater abundance of two proteins and reduced abundance of one in the culture media of cells derived from longphotoperiod-exposed hamsters.

Acknowledgments Studies reported in this chapter were supported by NIH grants HD 20300 (L.D.R.) and HD 20033 (A.B.). We gratefully acknowledge the contribution of Dr. Lonnie D. Russell to those studies. We apologize to those whose publications pertinent to this topic may have been inadvertently omitted.

V. SUMMARY

References

This review has attempted to highlight Sertoli cell biology in seasonal breeders and to summarize the structural responses of Sertoli cells in relation to changing endocrine status at different phases of gonadal activity. The seasonally breeding golden (Syrian) hamster, which exhibits photoperiod-dependent transitions from reproductively active to quiescence and to recrudescence, was used as a model to study Sertoli cell structure during different phases of gonadal activity. Short-photoperiod–induced testicular involution was associated with a significant decrease in virtually all major morphological parameters of the Sertoli cells. The striking changes in Sertoli cell morphology between active and inactive states of spermatogenesis are structural manifestations of alterations in the function of these cells in response to concomitant endocrine changes in the testis and indicate a virtual shutdown of Sertoli cell function during short-photoperiod– induced testicular regression. In contrast, no apparent changes in the Sertoli cell structure, except for outer and inner mitochondrial membrane surface areas, were evident during early gonadal regression. The general lack of Sertoli cell structural response in response to short-term exposure to an inhibitory photoperiod is similar to early response to hypophysectomy in the nonseasonal rat. During early-photoperiod–related recrudescence, there were, however, significant increases in the volume of SER and RER, and in the surface areas of outer and inner mitochondrial membranes, SER, and RER. However, only the RER (both volume and surface area) was correlated with the hormonal changes during early recrudescence, suggesting that the initiation of the Sertoli cell synthetic process was important in reestablishment of spermatogenesis. We should emphasize that seasonal changes in testicular activity can be major, resulting in a succession of states of fertility and complete sterility that are naturally occurring and fully reversible. Studies of the regulation of Sertoli cell function in seasonal breeders are aimed not merely at elucidating the structure– function relationship of this fascinating somatic cell of the testis, but are also designed to use this model system to identify the underlying mechanisms, that control the activity of these cells.

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C H A P T E R

7 Sertoli Cell Gene Expression and Protein Secretion MICHAEL D. GRISWOLD AND DEREK MCLEAN Center for Reproductive Biology, Department of Animal Sciences, Washington State University, Pullman, Washington

may be a small number of genes that are uniquely expressed by Sertoli cells and thus provide functional capabilities that distinguish them from other cells. Interest in this group of unique genes has been stimulated by the technology of transgenic manipulation of gene expression that can benefit from knowledge of transcriptional promoters that allow cell-specific expression of transgenes. Five hundred to 1000 papers have been published in the past 10 years that deal with transcription and/or gene expression in Sertoli cells, and these studies have provided a number of insights into the functions carried out by Sertoli cells. Most of these involved the study of one or two gene products in Sertoli cells and were aided by the increased availability of routine polymerase chain reaction (PCR) technologies. In recent years the availability of gene arrays has allowed the relatively complete transcriptome of a tissue or cell type to be defined. In this chapter, we provide a broad overview of the recent progress in understanding the transcriptome of the Sertoli cells via conventional molecular techniques and a more detailed view of the results obtained from gene array analysis. Gene expression analysis in cells and tissues has moved from the single experiment–single gene approach (Northern blot, RNAse protection, in situ hybridization) to a higher throughput, single experiment– thousands of genes approach with the development of microarrays or gene arrays [1]. Microarrays have been successfully used to evaluate gene expression in a wide variety of species and tissues [2–4]. Analysis of gene array data provides not only a snapshot of the

I. INTRODUCTION II. OVERVIEW OF GENE EXPRESSION IN SERTOLI CELLS III. GENES THAT ARE HIGHLY EXPRESSED BY OR UNIQUE TO SERTOLI CELLS IV. GENE ARRAY ANALYSIS OF SERTOLI CELL GENE EXPRESSION V. GENE ONTOLOGY ANALYSIS OF SERTOLI CELL GENE EXPRESSION VI. CHROMOSOME DISTRIBUTION OF SERTOLI CELL EXPRESSED GENES VII. CONCLUSION VIII. RESOURCES ON THE WORLD WIDE WEB References

I. INTRODUCTION The timing of the expression of a specific subset of the 35,000 or so genes of mammals (the transcriptome) provides each cell type with the characteristics that define its unique functions. The interest in the genes expressed in Sertoli cells began with the assumption that this information would provide insight into their role in testis formation and spermatogenesis. The two basic parameters of interest are the subset of genes that are expressed and the developmental timing of their expression. Within this subset of expressed genes, many are genes that are commonly expressed in other tissues and cell types or, perhaps, are only expressed in a very small number of cell types that may play similar biological roles. Also, within this subset we expect that there SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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Copyright 2005, Elsevier Science (USA). All rights reserved.

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Michael D. Griswold and Derek McLean

genes expressed in a particular cell or tissue at a given time, but can also be used to compare gene expression following a treatment or in two different tissues. Databases of gene expression information from different cells or tissues are often available and can be used as resources for in silico expression analysis.

II. OVERVIEW OF GENE EXPRESSION IN SERTOLI CELLS Based on the known roles of Sertoli cells in testis formation and development and support of spermatogenesis, researchers proposed that the important gene products are those that interact with other Sertoli cells or with germ cells [5]. These gene products are likely secreted or are external components of the cell membrane. Equally important are the genes involved in regulating the production of the proteins involved in the interactions. The expression of many genes in the Sertoli cells is regulated by developmental factors and specific endocrine and paracrine factors. The important gene products were put into four major categories: transport or bioprotective proteins, proteases and protease inhibitors, membrane glycoproteins, and growth factors and paracrine factors [5]. Since 1993 substantial progress has been made on the identification of proteins in each of these categories. Early studies on gene expression in Sertoli cells involved the identification and study of the endocrine regulation of specific proteins produced by cultured cells. Many of the gene products that were first characterized as gene products from Sertoli cells were transport proteins such as androgen binding protein (ABP) or transferrin [6]. The discovery of these and similar gene products has led to the idea that a major function of Sertoli cells is the transportation of important nutrients and regulatory factors around or across tight junction barriers to the sequestered germ cells [7]. Another group of proteins secreted by Sertoli cells that have been extensively characterized over the years are the proteases and protease inhibitors [8]. Progress in this area is extensively reviewed in Chapter 9 of this book. Sertoli cells synthesize and secrete many different proteases and inhibitors, and one of the functions ascribed to this class of proteins involves the extensive tissue remodeling that occurs when tight junctions must be traversed or when spermatozoa are released from the seminiferous epithelium [8]. Investigations of membrane glycoproteins have included investigations of membrane hormone receptors such as the FSH receptor (reviewed in Chapter 16) [9, 10] and the components of the junctional complexes

between adjacent Sertoli cells or between germ cell and Sertoli cells [11–14]. The ckit ligand [15–19] and the Müllerian inhibiting substance (MIS) [20–22] have been the major focus of recent studies of the group 4 secretions that include growth factors and paracrine factors. Progress in this area is reviewed in Chapters 8, 14, and 18 of this book. It is necessary to add an additional group of gene products from Sertoli cells that have been the subject of many studies in recent years. This fifth group of proteins includes transcription factors and signaling molecules. The GATA family of transcription factors [23–26] and several helix–loop–helix transcription factors [27–31] have been shown to be present in Sertoli cells. Sry and Sox9 expressed early in testis formation are putative transcription factors that regulate several genes involved in this process [32–37]. Examples of signaling systems shown to exist in Sertoli cells include the desert hedgehog and tyro-3 families [38, 39]. In addition, an extensive study of Jak-Stat signaling in the regulation of Sertoli cell responses has been done [40–42]. Signaling and transcription factors in Sertoli cells are reviewed in Chapter 15 in this book, whereas the genes involved in sex determination are reviewed in Chapter 4.

III. GENES THAT ARE HIGHLY EXPRESSED BY OR UNIQUE TO SERTOLI CELLS The genes expressed by Sertoli cells that have been of the most interest to investigators are those that are unique to or are very highly expressed primarily by this cell type. The interest in these gene products is stimulated by the potential to understand more about the function of Sertoli cells in testis formation and spermatogenesis and in the identification of mechanisms that lead to cell-specific transcription. This latter information can be useful in alteration of gene expression via transgenic technologies in a very cell-specific manner in the intact animal. The first use of this technology was reported in 1992 and was used to generate a mouse Sertoli cell line that has been designated the MSC-1 cell line [43]. Transgenic mice carrying a fusion gene composed of human MIS transcriptional regulatory sequences were linked to the SV40 T-antigen gene. These mice specifically developed testicular tumors composed of a cell type that histologically resembled the Sertoli cell. A cell line was derived from one of the testicular tumors that expressed markers associated with Sertoli cells, such as transferrin, sulfated glycoprotein-2, and inhibin βB. The cell line does not

Chapter 7 Sertoli Cell Gene Expression and Protein Secretion

express detectable levels of inhibin-α, MIS, or FSH receptor, but has many characteristics of Sertoli cells [44]. Another pioneering study in the use of Sertolispecific promoters involved the ABP gene. The androgen binding protein was shown in the testis to be specific to the Sertoli cells in the rat [45, 46] and a 5.5-kilobase (kb) rat DNA fragment containing the ABP gene with all 8 exon sequences and 1.5 kb upstream of the transcription start site was used to make transgenic mice. Expression of the gene was observed in the testis and brain, but not in other tissues of the transgenic mice [47]. The study verified that ABP was made in Sertoli cells and the testicular ABP mRNA in the transgenic mice was shown to be translated by dihydrotestosterone (DHT)-binding assays and immunohistochemistry. The androgen binding activities in the testis and epididymis of the hemizygous transgenic mice were elevated 25- to 50-fold compared to activity in the wild-type tissues. The transgenic testicular ABP was shown by immunohistochemistry to be primarily in the cytoplasm of Sertoli cells and lumen of the seminiferous tubules. Increased ABP levels in the transgenic mice were associated with a variable degree of abnormal spermatogenesis and extensive structural changes in the transgenic testis [47]. Mice with the spermatogenic disorder had reduced epididymal sperm numbers and reduced male fertility. The homozygous transgenic male and female mice also had a serious motor dysfunction affecting their hind limbs. The extreme cell specificity and the importance of the FSH response to spermatogenesis have led to an extensive characterization of the promoter of the FSH receptor gene. It was proposed that an understanding of the FSH promoter could lead to new ways to alter gene expression in Sertoli cells. However, in both cell transfections and in transgenic mice, the promoter directed the expression of transgenes promiscuously. It was proposed that repression and activation of local chromatin structure are likely to play a major role in the regulation of the FSH receptor gene [10]. The expression of the FSH receptor gene is reviewed in more detail in Chapter 16. Both the Pem gene promoter and the cathepsin L gene promoter have been used very successfully to drive Sertoli cell–specific gene expression in transgenic mice [48–50]. The results from the cathepsin L gene promoter are described in detail in Chapter 9 of this book. The Pem gene encodes a homeodomain related to those in the Prd/Pax gene family. Pem transcripts were shown to be preferentially expressed in stages VII and VIII of the cycle of the seminiferous epithelium. In addition, expression depended on androgens and

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gonadotrophins since injection of either testosterone or luteinizing hormone (LH) into hypophysectomized and hpg/hpg mice restored Pem expression in the testes to normal levels [51]. The 0.6-kb 5’-flanking sequence of the Pem gene directed transgene expression specifically in the testis and the epididymis but not in any other tissues. The promoter fragment targeted the transgene expression specifically to Sertoli cells during stages IV through VIII of the cycle of the seminiferous epithelium and reproduced the expression pattern of the endogenous Pem gene. The promoter fragment also reproduced the postnatal expression pattern of the Pem gene. If a 0.3-kb fragment was deleted from the 5’-end of the transgene there was no effect on androgen-dependent Sertoli-specific expression, but the stage-specific expression and the timing of the postnatal expression were changed. These results demonstrated that there are regions in the Pem promoter that separately direct expression specifically to Sertoli cells and imparts stage-specific expression [48].

IV. GENE ARRAY ANALYSIS OF SERTOLI CELL GENE EXPRESSION The concept of microarrays is based on the placement of DNA fragments (probes) at a high density on a solid platform or support. Several different supports are currently being used including membranes, glass slides, and glass wafers. The DNA fragments can be cDNA, genomic fragments, or short oligonucleotides (20–70 bp). DNA probes are placed on the support by mechanical spotting or synthesized in situ on the support in the case of the glass wafer. Glass slide or membrane spotted microarrays can be produced “in house” provided the equipment to spot the microarray is available. This platform also requires the cDNA or oligonucleotide that will be used to spot on the slide or membrane to be prepared or purchased from an outside party. Usually, cDNA libraries are used as the template for PCR reactions to synthesize products that are eventually spotted on the glass slide or membrane. Spotting 25- to 70-bp oligonucleotides requires synthesis and cleanup of these products. Glass slide or membrane microarrays are also available commercially. As with in-house spotted cDNA microarrays, the genes found on commercially available glass slide or membrane microarrays are usually limited to a specific tissue or biological process (e.g., apoptosis, cell cycle, cytokines). Microarrays using short (25-bp) oligonucleotides synthesized in situ on the support require complex equipment to produce. The Affymetrix produced

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GeneChip line of products is an example of a commercially available microarray that uses short oligonucleotides synthesized in situ on a glass wafer using photolithography. The major advantage of this platform is the number of probes that can be synthesized on the support. Affymetrix produces microarrays with 18,000 transcript probes on a single chip. Several of the available GeneChips include all of the genes or open reading frames for individual species (see www.affymetrix.com for details). The major disadvantage of this platform is that major equipment must be available to process and scan the GeneChips. The sequences of the oliogonucleotide probes on GeneChips are determined from information in public databases; therefore, the completion of genome sequencing projects in additional species will allow extensive coverage of genes and limit the need for cell-specific microarrays. Affymetrix GeneChips have been used to analyze the gene expression of rat and mouse Sertoli cells from prepubertal animals in vitro and from day 6 postpartum mouse Sertoli cells isolated by Sta-Put techniques [52]. The rat Sertoli cells were isolated from day 20 postpartum rats, cultured using standard techniques, and cultured for 2 days prior to RNA purification [53]. The goal of this study was to identify genes expressed by Sertoli cells in culture and genes whose expression is induced or repressed by follicle-stimulating hormone (FSH). Thus, rat Sertoli cells in culture were treated with 25 ng/mL of ovine (o) FSH for 2, 4, 8, and 24 hr and gene expression was compared with pretreatment controls. The ultimate endpoint of this research was to identify a new or an improved pathway model for FSH stimulation of Sertoli cells. In addition, this approach would identify genes that may play a role in Sertoli cell differentiation during puberty. This discussion is limited to insights gained from the analysis of the control Sertoli cell samples. Other chapters provide details of FSH regulation of gene expression in Sertoli cells. Two types of analysis were performed with the Affymetrix Rat U34A GeneChip datasets. First, an absolute analysis of the rat Sertoli cell expressed genes in control analysis provided a list of genes expressed in cultured rat Sertoli cells. Second, comparison analysis was used to determine genes induced or repressed by FSH at each time point in comparison to untreated controls. This analysis provided a list of genes that are changing due to the treatment with FSH. Subsequent analysis of the lists of genes by clustering or grouping genes based on ontology provides a useful way to determine what genes are involved in Sertoli cell function. Absolute analysis of GeneChip data for rat Sertoli cells in culture provides a window into cell function by characterization of gene expression. First, GeneChip analysis indicates that approximately 32% of the

TABLE 7.1 Nonribosomal Genes with the Highest Expression in Cultured Rat Sertoli Cellsa Signalb

Accession number

Description

21295.8

J01435

Mitochondrial cytochrome oxidase subunit I, II, and III genes

14294.4

D16554

Polyubiquitin

14148

M34043

Thymosin β4

13881.9

U11071

Polyadenylate-binding protein

13537.2

NM_031144

Actin, β

13317.4

NM_138895

Polyubiquitin

12738.1

M64733

Clusterin, TRPM-2

9637.4

AA945152

Cytochrome c oxidase polypeptide III

9377

X04229

Glutathione S-transferase (GST) Y(b) subunit

8445.5

NM_017202

Cytochrome c oxidase, subunit 4a

8187.6

NM_017101

Peptidylprolyl isomerase A (cyclophilin A)

7978.9

J03752

Glutathione S-transferase

7824.4

X52815

Cytoplasmic-γ isoform of actin

7790.5

NM_013013

Prosaposin

7749.1

J02810

Prostate glutathione S-transferase

6486.2

X61654

Ad1-antigen, CD63

6484.6

NM_031972

Aldehyde dehydrogenase family 3

6231.3

M58404

Thymosin β10

5994.5

V01227

α-tubulin

5751.8

M12919

Aldolase A

5740.8

D10706

Ornithine decarboxylase antizyme

5705.8

X75253

Phosphatidylethanolamine binding protein

5629.4

S79304

Cytochrome oxidase subunit I

5415.8

E01534

Insulinoma

5118.7

X02231

Glyceraldehyde-3-phosphatedehydrogenase (GAPDH)

5054.4

S81353

Sulfated glycoprotein-1 [rats, testes, mRNA Partial, 1692 nt]

5044.3

M35826

NADH-dehydrogenase (NDI)

a RNA from Sertoli cells was used as a template for GeneChip analysis with the Affymetrix Rat U34A GeneChip. Expression analysis was determined with the use of Microarray Suite 5.0. b Rat genome U34A GeneChip data were scaled to a signal of 125. Signal is approximately equal to the overall abundance of a particular transcript within a cell. A signal value of 100 is approximately equivalent to one to five copies of a transcript per cell.

transcripts on the array are expressed in cultured rat Sertoli cells. The rat U34 GeneChip set utilized for this analysis has more than 9000 transcripts represented on each array. This set of GeneChips includes three arrays, A, B, and C, in which the A chip is comprised primarily of identified genes, whereas B and C chips are ESTs. Thus, the data indicates that approximately

Chapter 7 Sertoli Cell Gene Expression and Protein Secretion

V. GENE ONTOLOGY ANALYSIS OF SERTOLI CELL GENE EXPRESSION Gene ontology analysis is a technique used to group genes expressed in a sample in terms of their biological process, location in the cell, or function. The Gene Ontology (GO) Consortium (www.geneontology.org) has established three structured, controlled ontologies that describe gene products in terms of their associated biological processes, cellular components, and molecular functions in a species-independent manner. The three ontologies are biological process, cellular component, and molecular function and are available for download at the GO Consortium website. These main categories can then be refined based on, for example, physiological process or cellular component. Some genes will fall into more than one category, which improves the accuracy of the analysis. Most gene array analysis software provides gene ontology functional sorting. GeneSpring 5.0 (www.silicongenetics.com) and GeneSifter (www.genesifter.com) were used for gene ontology analysis with the rat Sertoli cell dataset. Gene ontology analysis of the 2750 genes expressed by cultured rat Sertoli cells provides valuable information about the biological process and molecular function of the cell (Fig. 7.1). Due to the number of ESTs present in the GeneChip dataset and the incomplete nature of the ontology datasets, a total of 1014 of the Sertoli genes are included in the ontology analysis (37% of the genes present). The largest number of expressed genes was placed in the biological process category (431 genes) followed by molecular function (354 genes) and cellular component (229 genes). Further categorization of these ontological groups into subontology categories is shown in Figures 7.2, 7.3, and 7.4, respectively. The biological process ontology group can be further categorized into four subcategories (Fig. 7.2): (1) physiological

500 450 400 Gene Number

2750 genes on each GeneChip are expressed in cultured rat Sertoli cells. Although some duplication of genes on the GeneChips does occur due to the presence of ESTs, 32% is probably an accurate estimate of the number of genes expressed by these cells. Therefore, extending this analysis and taking into account the estimation that higher mammals have 36,000 genes in their genomes, Sertoli cells would express approximately 11,500 genes. This evaluation does not take into consideration alternative transcripts or other transcriptional or post-transcriptional modification that may occur. Table 7.1 shows the 28 nonribosomal genes with the highest expression in rat Sertoli cells from the Affymetrix rat U34A GeneChip.

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350 300 250 200 150 100 50 0 Biological Process

Cellular Component

Molecular Function

FIGURE 7.1 Gene ontology analysis of Sertoli cell gene expression based on genes called present on the Affymetrix U34A GeneChip. The number of genes in each ontology is shown on the y-axis and the gene ontology category on the x-axis. Gene ontology categories are based on those of the Gene Ontology Consortium (www.geneontology.org).

processes, (2) cellular processes, (3) development, and (4) behavior. The development and behavior groups contained fewer than 20 genes, so further analysis of these datasets is not presented. These subcategories can be further refined into more descriptive groups. It is this third layer of ontology analysis that begins to provide useful information that can be used to gain a clearer understanding of how the expressed genes may affect cell function. For example, analysis of the subcategory physiological process from the biological process ontology demonstrates that almost half of the 431 genes expressed in Sertoli cells found in the biological process ontology can be subcategorized into the metabolism ontology (203 genes, Fig. 7.2B). It also indicates that another 87 genes expressed in Sertoli cells are found in the cell growth/maintenance category (Fig. 7.2B). Further analysis of the cell growth/maintenance category shows that more than half of the 87 genes expressed in Sertoli cells in this category are related to transport (45 genes), followed by 17 Sertoli cell transcripts in the cell organization and biogenesis category and 15 in the cell proliferation category (not shown in Figure 7.1). Sertoli cells express 140 genes found in the cellular process subcategory in the biological process main ontology that can be further refined to cell growth/maintenance, cell communication, cell death, and cell motility (Fig. 7.2C). An example of overlap between two ontologies is apparent when comparing the physiological process and cellular process subcategories within the overall biological process ontology. The physiological process and cellular process subcategories both contain a cell growth/maintenance subcategory that has a different number of genes in the group. This does not impair

100 A

Michael D. Griswold and Derek McLean A

Biological Process Ontology

300

160 140

Gene Number

Gene Number

250 200 150 100

Physiological Processes

Cellular Process Development

60 40

Intracellular

Biological Process – Physiological Processes Ontology

B

100

80

90

70

80

Gene Number

Gene Number

80

Behavior

90

60 50 40 30 20 10 Metabolism

100

Cell Fraction Cell Surface

Cellular Component – Cell – Intracellular Ontology

70 60 50 40 30

Cell Growth and/ Response to Response to or Maintenance External Stimulus Stress

10

Death

0

Biological Process – Cellular Process Ontology

Cytoplasm

90 80

C

70 60

40

Nucleus

Ribonucleo- Proton-transporting protein Complex ATP Synthase Complex Cellular Component – Cell – Membrane Ontology

35

50 40 30 20 10 0 Cell Growth and/or Cell Communication Maintenance

Cell Death

Cell Motility

FIGURE 7.2 Gene ontology subcategories of the biological process ontology for the Sertoli cell gene expression data from Affymetrix U34A GeneChip. The number of genes in the ontologies is shown on the y-axis and the gene ontology category on the x-axis. (A) Subcategories for the biological process ontology (see Fig. 7.1). (B) Further categorization of the physiological process subcategory from part (A). (C) Further categorization of the cellular process subcategory from part (A).

analysis because both groups of genes are analyzed independently. Sertoli cell expressed genes present in the cellular component ontology are primarily found in the cell subcategory (216 expressed genes) with only a small number of expressed genes in the extracellular (8) or unlocalized (5) subcategories (Fig. 7.3). Within the cell subcategory (Fig. 7.3B), the majority of the Sertoli cell expressed genes are in the intracellular group (152 genes) and the membrane group (64 genes). Further characterization of the intracellular and membrane ontologies is shown in Fig. 7.3C. The membrane group

Gene Number

Gene Number

Membrane

20

0

C

100

0

0

100

120

20

50

B

Cellular Component – Cell Ontology

30 25 20 15 10 5 0 Plasma Membrane

Integral to Membrane

Endomembrane Mitochondrial System Membrane

FIGURE 7.3 Gene ontology subcategories of the cellular component ontology for the Sertoli cell gene expression data from Affymetrix U34A GeneChip. The number of genes in the ontologies is shown on the y-axis and the gene ontology category on the x-axis. (A) Subcategories for the biological process ontology (see Fig. 7.1). (B) Further categorization of the physiological process subcategory from part (A). (C) Further categorization of the cellular process subcategory from part (A).

categorizes expressed genes in groups according to the membrane location within the cell. Information inferred from these datasets provides an example of how gene ontology analysis can lead to hypotheses about cell function. The plasma membrane and integral-to-membrane groups provide useful information on receptors whose genes are expressed in Sertoli cells and therefore may be present in the plasma membrane of these cells. Genes for

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Chapter 7 Sertoli Cell Gene Expression and Protein Secretion A

Molecular Function Ontology

180 160

Gene Number

140 120 100 80 60 40 20 0 Enzyme Activity B

60

Binding Signal Transporter Transcription Structural Enzyme Activity Transducer Activity Regulator Molecule Regulator Activity Activity Activity Activity Molecular Function – Enzyme Activity Ontology

Gene Number

50

40

30

20

10

0 Transferase Hydrolase Kinase Oxidoreductase Lyase Activity Activity Activity Activity Activity C

20

Ligase Activity

Isomerase Activity

Molecular Function – Binding Activity Ontology

18 16 Gene Number

receptors not known to be expressed in Sertoli cells can be identified and experiments designed to test the hypothesis that ligands to these receptors can contribute to cell function. The final gene ontology group is the molecular function category (Fig. 7.4). This ontology group contains the largest number of gene categories or subontologies. The largest number of genes expressed by Sertoli cells is in the enzyme activity subcategory (Fig. 7.4A). Further analysis of this subcategory (Fig. 7.4B) shows that genes for enzymes with transferase, hydrolase, kinase, and oxidoreductase activity are the most prevalent genes expressed in Sertoli cells. The other two subcategories with large numbers of Sertoli expressed genes in the molecular function ontology are binding activity and signal transducer activity. Further characterization of the Sertoli cell expressed genes in the binding activity subcategory is shown in Fig. 7.4C. This analysis indicates that the majority of the Sertoli cell expressed genes in this group are involved in nucleic acid binding and nucleotide binding activity. Lastly, the signal transducer group provides the number of Sertoli cell expressed genes involved in receptor and receptor signaling protein activity (Fig. 7.4D). The ontology classification of gene expression data provides broad or specific distribution of genes. How this distribution relates to the expected number of genes expressed in a cell is also of interest. This analysis can indicate unexpected trends in expression data. The expected ontology distribution of expressed genes in a cell type depends on the number of expressed genes found in each ontology on the gene chip. This information can be used to calculate a Z-score for each ontology group [54]. The Z-score indicates whether the specific GO term occurs more or less frequently than expected in the gene list. Extreme positive numbers indicate that the term occurs more frequently than expected, whereas an extreme negative number indicates that the term occurs less frequently than expected. The number of genes in each ontology category and subcategory present in Figures 7.2, 7.3, and 7.4 is listed in Table 7.2. The Z-score for the rat Sertoli cell data is also included for each GO term in Table 7.2. The Z-scores of the ontology data for rat Sertoli cell gene expression indicates that the number of genes expressed in several categories occurs more frequently than expected. Within the biological process GO group, metabolism has a Z-score of 7.0, the highest Z-score on the list. The highest Z-score in the cellular component GO group is the intracellular category with a value of 7.77. The enzyme category had the highest Z-score (5.22) in the molecular function group. The lowest Z-score in the three GO categories was signal transducer activity (–6.35) in the molecular function group.

14 12 10 8 6 4 2 0 Nucleic Acid Nucleotide Receptor Binding Protein Binding Neurotransmitter Binding Activity Binding Activity Activity Activity Binding Activity

FIGURE 7.4 Gene ontology subcategories of the molecular function ontology for the Sertoli cell gene expression data from Affymetrix U34A GeneChip. The number of genes in the ontologies is shown on the y-axis and the gene ontology category on the x-axis. (A) Subcategories for the molecular function ontology (see Fig. 7.1). (B) Further categorization of the enzyme activity subcategory from part (A). (C) Further categorization of the binding activity subcategory from part (A). (D) Further categorization of the binding activity ontology.

Interpretation of this information can be challenging due to the static nature of the data and biological broad categories the ontology represents. However, it is not surprising that expression of metabolism genes is higher than expected in Sertoli cells. These cells are known to have diverse active metabolic pathways.

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Michael D. Griswold and Derek McLean TABLE 7.2 Number of Genes Expressed in Rat Sertoli Cells Segregated in Each Gene Ontology Group and the Associated Z-Scorea Ontology term

Genes

Biological Process Physiological Processes

Z-score

Ontology term

431 Genes 273

Metabolism

Z-score

Transporter activity

30

–4.24

Transcription regulator activity

21

–1.15

203

6.89

Cell growth and/ or maintenance

87

–1.57

Response to external stimulus

15

–2.08

Response to stress

7

–1.21

Death

7

1.27

Chaperone activity

3

1.63

Homeostasis

6

0.69

1

1.34

Bone remodeling

2

0.14

Translation regulator activity

1

–1.25

Anticoagulant activity

1

0.42

Motor activity

1

–0.45

Defense/immunity protein activity

1

–1.56

Hemostasis Cellular Process

140 Cell growth and/or maintenance

87

–1.57

Cell communication

Structural molecule activity

21

2.2

Enzyme regulator activity

12

1.11

Cell adhesion molecule activity

4

–0.18

Apoptosis regulator activity

3

1.13

55

–3.51

Cell death

7

1.27

Transferase activity

56

3.49

Cell motility

6

0.25

Hydrolase activity

54

0.51

Kinase activity

36

2.07

Oxidoreductase activity

29

1.85

Cellular Component Cell Intracellular

229 Genes 216 152

Enzyme Activity

7.7

Lyase activity

13

1.52

Membrane

64

–6

Ligase activity

8

2.92

Cell fraction

41

–1.96

Isomerase activity

7

2.63

Cell surface

1

1.36

1

1.34

Cytoplasm

89

6.21

Sterol carrier protein X-related thiolase activity

Nucleus

57

2.53

1

1.34

Ribonucleoprotein complex

12

4.77

Small protein conjugating enzyme activity

Proton-transporting ATP synthase complex

2

1.93

Nucleic acid binding activity

19

Cell cortex

1

1.36

15

1.17

TCA cycle enzyme complex

1

1.36

Nucleotide binding activity Receptor binding activity

5

–2.98

Protein binding activity

3

–0.16

Neurotransmitter binding activity

2

–2.21

Intracellular

Binding Activity

Membrane Plasma membrane

36

–2.57

Integral to membrane

–0.7

25

–2.26

Endomembrane system

4

1.24

Metal ion binding activity

1

0.42

Mitochondrial membrane

3

0.44

Peptide binding activity

1

–3.28

Steroid binding activity

1

1.34 –5.93

Inner membrane

2

0.24

Extrinsic to membrane

1

1.36

Outer membrane

1

0.45

Receptor activity

22

5.22

Receptor signaling protein activity

10

0.28

5

–2.98

Molecular Function Enzyme activity

a

Genes

354 Genes 169

Binding activity

47

–3.61

Signal transducer activity

40

–6.35

Signal Transducer

Receptor binding activity

The three main ontologies—biological process, cellular component, and molecular function—are in bold. The subcategories within each of these groups are shown in italics. The Z-score indicates whether a specific GO term occurs more or less frequently than expected in the gene list. Positive Z-scores indicate that the term occurs more frequently, whereas negative scores indicate that the term occurs less frequently than expected.

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Chapter 7 Sertoli Cell Gene Expression and Protein Secretion

Likewise, prior to hormonal stimulation, Sertoli cells do not have high levels of signal transduction activity. The number of genes expressed in these groups would likely change on external activation of this cell type.

VI. CHROMOSOME DISTRIBUTION OF SERTOLI CELL EXPRESSED GENES The completion of the genome projects for model organisms such as the rat and mouse provides a useful database of information to evaluate the distribution of genes expressed in cells across the genome. The chromosomal distribution of genes expressed in rat Sertoli cells is shown in Table 7.3. This information was determined with the use of GeneSifter and NCBI genome databases (www.ncbi.nih.gov/Genomes). The limitation of this analysis is similar to the ontology analysis in that the presence of all genes is not known for the

rat (a total of 22,449 genes are present on the NCBI database) and the Y chromosome has not been sequenced. Similarly, a total of 1843 of the 2750 genes expressed in Sertoli cells (67%) from the U34A GeneChip can be located to the rat chromosomes. However, the initial analysis can indicate if genes expressed in Sertoli cells are clustered to a particular chromosome. Subsequent genomic experimentation could identify common elements consistent with Sertoli cell expressed genes located in a particular region of a chromosome. The analysis of the chromosomal distribution of Sertoli cell expressed genes was determined by comparing the percentage of the total of all genes (22,449) on each chromosome with the percentage of the total genes expressed in Sertoli cells found on each chromosome (Table 7.3). The number of genes on the Y chromosome for the mouse was substituted in this analysis. The difference in the percentages was calculated and trends were evaluated. The percentage of

TABLE 7.3 Chromosomal Distribution of Genes Expressed in Rat Sertoli Cellsa Chromosome

Genes

1

2933

13.1

260

14.1

2

1604

7.1

137

7.4

0.3

3

1657

7.4

106

5.8

–1.6

4

856

3.8

91

4.9

1.1

5

1395

6.2

106

5.8

–0.5

6

1050

4.7

97

5.3

0.6

7

1408

6.3

139

7.5

1.3

8

1200

5.3

114

6.2

0.8

9

816

3.6

73

4.0

0.3

10

1598

7.1

176

9.5

2.4

11

614

2.7

51

2.8

0.0

12

607

2.7

60

3.3

0.6

13

698

3.1

60

3.3

0.1

14

763

3.4

49

2.7

–0.7

15

889

4.0

39

2.1

–1.8

16

691

3.1

56

3.0

0.0

17

706

3.1

29

1.6

–1.6

18

567

2.5

31

1.7

–0.8

19

615

2.7

55

3.0

0.2

20

564

2.5

57

3.1

0.6

X

1082

4.8

54

2.9

–1.9 –0.4

Y (Mouse) Total

Percentage

Sertoli genes

Percentage

136

0.6

3

0.2

22449

100.0

1843

100.0

Percentage difference 1.0

a The number of genes present on each chromosome was determined from NCBI databases. The number of Sertoli genes present on each chromosome was determined with GeneSifter.

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Michael D. Griswold and Derek McLean

genes expressed in Sertoli cells on rat chromosome 10 was 2.5% higher than expected. In addition, the percentage of genes expressed in Sertoli cells on chromosomes 1, 4, and 7 was 1.0%, 1.1%, and 1.3% higher than expected, respectively. The number of genes expressed in Sertoli cells was lower than expected on several chromosomes as well. Chromosome 3, 15, and 17 had 1.6%, 1.8%, and 1.5% fewer Sertoli cell expressed genes on them than expected. Interestingly, the number of expressed rat Sertoli cell genes present found on the X chromosome was 1.9% lower than expected. Further genomic analysis must be conducted to determine the significance of this information, however, it may provide an interesting avenue of study.

VII. CONCLUSION Gene array analysis provides a unique opportunity to evaluate global gene expression in cells or tissues. The data presented here represent an initial step in determining the genes necessary for the proper functioning of a Sertoli cell. Additional analysis including changes in gene expression following hormone or growth factor stimulation over time and the influence of germ cells on Sertoli cell gene expression will aid in completing the picture. Indeed, these data were obtained from Sertoli cells in vitro; thus germ cell influence and changes due to the cycle of the seminiferous epithelium will have a profound impact on the genes expressed by Sertoli cells. Gene array and genomic approaches, along with proteomic and other high-throughput techniques, will significantly enhance our understanding of cellular function within the testis. The Sertoli cell transcriptome will provide a key piece to the puzzle of genes necessary for sperm production.

VIII. RESOURCES ON THE WORLD WIDE WEB Affymetrix Gene Ontology Consortium NCBI Genomes Gene Expression Omnibus(GEO) Mouse Genome Informatics Wellcome Trust Sanger Institute

www.affymetrix.com www.geneontology.com www.ncbi.nlm.nih.gov www.ncbi.nlm.nih.gov/Genomes www.ncbi.nlm.nih.gov/geo www.informatics.jax.org www.sanger.ac.uk

Weizmann Institute http://bioinformatics. of Science— weizmann.ac.il/cards GeneCards The Institute www.tigr.org for Genomic Research (TIGR)

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Chapter 7 Sertoli Cell Gene Expression and Protein Secretion

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C H A P T E R

8 Sertoli Cell Secreted Regulatory Factors MICHAEL K. SKINNER Center for Reproductive Biology and Center for Integrated Biotechnology, School of Molecular Biosciences, Washington State University, Pullman, Washington

I. II. III. IV. V.

as autocrine agents to influence Sertoli cells or paracrine factors to influence neighboring cells such as spermatogenic cells, Leydig cells, or peritubular myoid cells [1]. These regulatory factors are essential for the careful control of testis development, spermatogenesis and male fertility, and male reproductive endocrinology. Two major subcategories of regulatory factors are growth factors and hormones. Growth factors are defined as factors that influence cell proliferation and tissue growth. Although growth factors directly influence the cell cycle, growth factors can also have effects on a variety of other cellular functions and cell differentiation. Properties of a variety of growth factors are presented in Table 8.1. All the growth factors listed have been shown to be Sertoli cell secreted factors and are discussed in more detail later. These Sertoli cell secreted regulatory factors are all required for normal testis function and male fertility. Another subcategory of Sertoli cell secreted regulatory factors is hormones. The local actions of hormonelike factors are often required to directly influence cellular function and differentiation. In contrast to growth factors, hormones generally do not directly influence cell proliferation. However, indirect effects mediated through the production of a growth factor do occur. These hormones are also needed to act as an endocrine agent at distant tissue and organs to regulate the reproductive endocrinology of the male. The positive and negative feedback systems built into endocrine regulation require the local production of hormones. Sertoli cell secreted hormones are discussed later.

INTRODUCTION GROWTH FACTORS HORMONES OTHER REGULATORY FACTORS SUMMARY References

I. INTRODUCTION The Sertoli cell provides the microenvironment and cytoarchitectural support for the developing spermatogenic cells and directly regulates the reproductive endocrinology of the male. This is in large part accomplished through the secretion of a wide variety of proteins. These secretory products include transport proteins to provide nutrient support to the germ cells, extracellular matrix and cell adhesion molecules to promote appropriate cell–cell interactions, and proteases to allow tissue remodeling during spermatogenesis. These Sertoli cell secreted factors can be categorized as nutritional factors (e.g., transport proteins) that support the nutrient requirements of the germ cells, environmental factors (e.g., extracellular matrix) that influence the physical content and extracellular environment between cells, and regulatory factors (e.g., growth factors) [1]. Regulatory factors are defined as factors that, through receptor-mediated signal transduction events, influence cellular function, growth, or differentiation on a molecular level. These regulatory factors can act SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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Copyright 2005, Elsevier Science (USA). All rights reserved.

108

Michael K. Skinner TABLE 8.1 Properties of Several Common Growth Factors

Growth factor

Abbreviation

Approximate size (kDa)

Examples of physiological action

Receptor(s)

Insulin-like growth factor I

IGF-I

7.5

Skeletal growth

IGF-I receptor

Insulin-like growth factor II

IGF-II

7.5

Fetal development

IGF-I and IGF-II

Epidermal growth factor

EGF

6

Tissue growth

EGF receptor

Transforming growth factor α

TGFα

5

Tissue growth

EGF receptor

Transforming growth factor β

TGFβ

25/dimer

Growth inhibition/tissue repair

TGFβ, type 1, 2, and 3 receptors

Fibroblast growth factor

FGF

17

Angiogenesis/tissue growth

FGF receptor

Neurotropins

NT-3

13

Neuronal development

TrkC receptor

Interleukin 1

IL-1

17

Immune response/inflammation

IL-1 receptor

Kit ligand/stem cell factor

KL/SCF

30

Tissue growth/germ cells

KL receptor

Glial cell–derived neurotrophic factor

GDNF

34

Cell growth

Ret/GFRα receptor

The initial review of this topic was published more than 10 years ago in The Sertoli Cell [2]. The advances since that time have been significant and will be the focus of the current review. Both Sertoli cell secreted growth factors and hormones are discussed. These secreted factors play a critical role in the development, growth, and maintenance of testis function. Sertoli cells are the principal cell regulating the process of spermatogenesis and these secreted regulatory factors are essential for Sertoli cell function.

II. GROWTH FACTORS The development of the testis and maintenance of spermatogenesis require a precise growth regulation [3, 4]. All cell populations proliferate during embryonic testis development and in the early postnatal period. Sertoli cells become postmitotic and terminally differentiate during the early pubertal period once spermatogenesis is initiated [3–5]. The other somatic cells in the testis (e.g., Leydig, peritubular) have a slowed but continuous rate of growth in the adult [4–8]. Spermatogenic cells require a rapid rate of mitosis and meiosis at the onset of puberty that continues throughout adult life. The waves of spermatogenic cell proliferation require local control by Sertoli cells. The secretion of growth factors by Sertoli cells will have a role in the precise cell growth regulation required in the developing and adult testis.

A. Insulin-Like Growth Factors Growth factors with structural similarity to insulin are in a family of factors termed insulin-like growth

factors (IGFs). These include IGF-I and IGF-II. IGF-I is a critical factor required for cell cycle progression and DNA synthesis in all proliferating cells. For this reason IGF-I is expressed and acts on nearly all cells. The serum level of IGF-I is high (e.g., 100 ng/mL) and is a principal liver product [9]. A family of IGF binding proteins exists to regulate the level of IGF bioactivity available to a cell and influence the homeostasis of the extracellular growth factors. All testicular somatic cells express and respond to IGF-I including Sertoli cells [10–13]. IGF-I influences DNA synthesis and increases lactate and transferrin production by Sertoli cells [14, 15]. IGF-I also can influence the actions of FSH on Sertoli cells [16, 17] through unique signaling events [17]. Although the IGF-I produced by Sertoli cells [10–13] can act as an autocrine factor on the Sertoli cells, the high concentration of IGF-I in the interstitial fluid is available to the basal surface of the Sertoli cell. The blood–testis barrier created by Sertoli cells sequesters the developing spermatogenic cells. IGF-I available in the interstitium does not pass this barrier. Therefore, IGF-I secreted by Sertoli cells will likely have a critical role as a paracrine factor on germ cells (Fig. 8.1). Germ cells appear to be a site for IGF-I actions [18, 19]. Local production of IGF binding proteins by Sertoli cells has been shown to influence IGF-I actions and will contribute to this proposed Sertoli cell–germ cell interaction [12, 20, 21]. The ability of IGF-I produced by Sertoli cells to influence Leydig cell function has also been proposed [22]. Although these interactions can be observed in vitro, the physiological importance in vivo is questioned due to the levels of IGF-I and binding proteins in the interstitium [23]. It is likely Sertoli cell secreted IGF-I will be critical for spermatogenic cells, but actions on other

109

Chapter 8 Sertoli Cell Secreted Regulatory Factors

testis somatic cells is questioned due to other sources of IGF-I for these cells. IGF-II has also been shown to mediate Sertoli cell– spermatogenic cell interactions [24–27]. The IGF-II receptor is present on both Sertoli cells and germ cells [25]. This receptor can also influence gene expression in the spermatogenic cells [26]. The IGF-II receptor is also termed the mannose-6-phosphate receptor and can bind a variety of additional ligands with appropriate carbohydrate specificity. Therefore, this IGF-II receptor system appears to mediate interactions between Sertoli cells and germ cells through a variety of potential ligands [27].

B. Transforming Growth Factor α Transforming growth factor α (TGFα) is a member of the epidermal growth factor (EGF) family and binds to the same EGF receptor as EGF [28, 29]. TGFα is expressed as a transmembrane precursor that is proteolytically processed to release the mature peptide (Table 8.1). TGFα is secreted by a large number of normal cell types and promotes cell proliferation in normal and transformed cells. EGF has been shown to influence spermatogenesis [30], and removal of serum levels of EGF reduces sperm numbers [31]. Circulating concentrations of EGF, however, are thought to be too low to have physiological influences [32]. EGF is not expressed in the rodent testis or in Sertoli cells [33]. In contrast, TGFα is expressed by Sertoli cells [33]. Both peritubular myoid cells and Leydig cells also produce TGFα [33, 34]. Peritubular cells and

Leydig cells proliferate in response to TGFα [33]. TGFα has been shown to influence Sertoli cell lactate and estrogen production [35] and can influence Sertoli cell DNA synthesis if the cells are derived from prepubertal animals [36, 37]. TGFα also influences transferrin production by Sertoli cells [38]. The actions of TGFα on Sertoli cells and the testis are dependent on the developmental stage. Embryonic testis growth requires TGFα expression and action on most cell types [4]. At the onset of puberty, the Sertoli cells differentiate and become postmitotic. Sertoli cell responsiveness to TGFα/EGF declines as the adult stage of development is obtained [39]. Other cell types continue to utilize TGFα for growth regulation. The spermatogenic cells also utilize TGFα, and Sertoli cell–derived TGFα is postulated to be an important paracrine interaction of this Sertoli cell regulatory factor [40] (Fig. 8.1). Sertoli production of TGFα is also capable of influencing adjacent peritubular cells [39, 40].

C. Transforming Growth Factor β Transforming growth factor β (TGFβ) has three mammalian isoforms (TGFβ1, TGFβ2, and TGFβ3) that act as inhibitors of cell proliferation and facilitate cellular differentiation in a wide variety of cells [41]. TGFβ acts by inhibiting the actions of growth stimulators (e.g., TGFα) and promoting the expression of extracellular matrix components. Most cell types contain the receptors for TGFβ isoforms. Although found in transformed cells, TGFβ regulates most normal cell types [41].

Leydig

Sertoli

Germinal

bFGF TGF-α TGF-β IL

Leydig Cell IGF1 KL Sertoli Cell

Peritubular Cell

TG bF F-α G ,T F, G N FT- β 3 ,

Peritubular

Germinal Cell IL-1, IGF, TGF-α, TGF-β, bFGF, FGF-4, KL, GDNF

FIGURE 8.1. Sertoli cell secreted regulatory factors.

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TGFβ isoforms are expressed by Sertoli cells [40, 42, 43] and Sertoli cells are one of the few tissues that express TGFβ3 [40, 44]. Sertoli cells express all three isoforms under distinct hormonal regulation [40, 45]. The developmental regulation of TGFβ expression in the testis of the different isoforms suggests distinct functions and/or differential regulation [45, 46]. TGFβ expression by Sertoli cells is critical for embryonic testis development [46, 47]. At the onset of puberty, TGFβ expression by Sertoli cells also corresponds to alterations in cell growth [45]. TGFβ1 is high prepubertally and then declines to low levels in the adult. TGFβ2 expression is similar and FSH appears to promote the decrease in expression by Sertoli cells [45]. TGFβ3 has a transient increase in expression at the onset of puberty [45]. The site of action for Sertoli cell secreted TGFβ is suggested from the analysis of TGFβ receptor localization. All cell types in the testis contain type I and II TGFβ receptors [48–50], including spermatogenic cells. Peritubular cells produce and respond to TGFβ with a decline in cellular proliferation induced by TGFα [42, 45]. Peritubular cells also produce more extracellular matrix components in response to TGFβ [42]. TGFβ also induce peritubular cell contractility that is needed for sperm transport in the seminiferous tubule [51]. TGFβ does not have major effects on Sertoli cell proliferation or functional genes, but does increase production of plasminogen activator [52]. TGFβ also appears to influence the tight junctions between Sertoli cells [53, 54]. Leydig cells are also a site of TGFβ production and action. TGFβ inhibits Leydig cell steroidogenesis [55–57]. Therefore, critical targets for the Sertoli cell secreted TGFβ isoforms are spermatogenic cells and peritubular cells (Fig. 8.1). Abnormalities in TGFβ expression have been suggested as causal in testicular diseases such as fibrosis [58]. The bioactivity of TGFβ produced by Sertoli cells is also a factor in the function of TGFβ [59]. TGFβ also can be inhibited by specific proteins such as Bambi [60], which can influence the actions of the Sertoli cell expressed TGFβ. TGFβ is a critical regulatory factor produced by Sertoli cells to influence testis development, function, and spermatogenesis.

D. Fibroblast Growth Factor Several members of the fibroblast growth factor (FGF) family have been shown to influence testis function. FGF family members regulate cellular growth and differentiation, as well as tissue angiogenesis [61]. FGFs have widespread tissue distribution and each of the family members has distinct receptors and specific functions. Basic FGF (bFGF) is one of the most studied FGF family members. Initially bFGF was identified in

whole testis [62, 63] and subsequently was found to be expressed by Sertoli cells [64, 65]. bFGF production by Sertoli cells was found to be increased by FSH [65]. bFGF has been shown to stimulate Sertoli cell growth prepubertally [66]. The ability of FSH to simulate Sertoli cell growth prepubertally may in part be indirectly mediated through bFGF production [65]. Early postnatal actions of FGF also suggest potential indirect effects on gonocyte proliferation [67]. Sertoli cells also respond to bFGF by an increase in the production of proteoglycans [68], lactate, and glucose metabolism [69]. Localization of the bFGF receptor (FGFR type 1) demonstrated receptors in Sertoli cells, Leydig cells, peritubular cells, and germ cells [70]. Therefore, the actions of bFGF secretion by Sertoli cells could mediate cellular interactions with spermatogenic cells and peritubular cells, as well as autocrine actions on Sertoli cells (Fig. 8.1). Another member of the FGF family, Hst-1/FGF-4, has been shown to be expressed by Sertoli cells [71]. A conditional knockout of FGF-4 resulted in impaired fertility and a knockin caused enhanced spermatogenesis [72]. This observation suggests a potential Sertoli cell–spermatogenic cell interaction mediated by FGF-4 (Fig. 8.1). The overexpression of FGF-4 was also found to protect the testis from the chemotherapeutic drug adriamycin [72]. Therefore, Sertoli cell expression of FGF-4 appears to be important for normal testis function. FGF-9 is another member of the FGF family that has been shown to be important for early embryonic testis development [73]. FGF-9 appears to be a downstream gene to Sry and is expressed by Sertoli cells to influence male sex differentiation through the adjacent mesenchymal tissue. A knockout of FGF-9 results in sex reversal and impaired testis development [73]. Functions of FGF-9 later in development have not been thoroughly investigated.

E. Neurotropins The neurotropin family of growth factors includes nerve growth factor (NGF), neurotropin 3 (NT-3), neurotropin 4/5 (NT-4/5), and brain-derived neurotrophic factor (BDNF). The high-affinity receptors for these factors are the trk receptors and the low-affinity ones are the LNGFR. Although these neurotropins are critical for neurons and associated cells, they also have been shown to be expressed by a number of nonneuronal tissues including the testis. NGF has been the most highly investigated neurotropin in the testis. NGF appears in the adult testis to primarily be expressed by spermatogenic cells [74–76], whereas the NGF receptor trkA is expressed in Sertoli cells [76–78]. The NGF precursor protein is highly expressed in round spermatids, while the NGF acts to

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promote Sertoli cell viability [76] and cellular functions such as lactate and estrogen production [78]. Although some studies have suggested NGF expression in the adult Sertoli cell [79], the majority have suggested NGF mediates a spermatogenic cell–Sertoli cell interaction and adult Sertoli cells do not express NGF [76–78, 80]. In contrast to adult Sertoli cells, NGF does appear to be expressed by late embryonic and early postnatal Sertoli cells [81]. The NGF receptor (trkA) was expressed in Sertoli cells, peritubular cells, and interstitial cells at these stages of development [81]. Therefore, NGF is a Sertoli cell secreted growth factor, but its expression by Sertoli cells appears to be more important in the perinatal period than in the adult. Other neurotropins expressed in the adult testis have not been extensively studied. NT-3 was found to localize in spermatogenic cells, whereas BDNF may be expressed by adult Sertoli cells [82]. In contrast, BDNF is not expressed in the embryonic or postnatal testis [81]. More thorough investigation of other neurotropins is needed to clarify roles in the adult testis. Observations suggest that Sertoli cell expressed neurotropins play a critical role during development, but in the adult, spermatogenic cells influence Sertoli cells through neurotropin production. NT-3 appears to be a critical neurotropin secreted by Sertoli cells during the initial stages of male sex determination and testis development [81, 83]. At the time of Sertoli cell fate differentiation by the testis determining factor Sry, Sertoli cells express NT-3 as a potential immediate downstream gene to Sry [81, 83]. This Sertoli cell secreted NT-3 acts on adjacent mesonephros cells as a chemotactic factor and promotes migration into the testis, and the migrating mesonephros cells become peritubular cells and promote seminiferous cord formation, which is the first morphological event in testis development [81, 83, 84]. The NT-3 receptor trkC is on the migrating mesonephros cells and blocking the actions of this receptor inhibits cord formation and testis development [83]. This has also been recently confirmed in the human fetal testis [85]. NT-3 also can act as a direct chemotactic agent to induce mesonephros migration into the developing testis [84]. Observations suggest Sertoli cell secretion of NT-3 is a downstream event to Sry and critical to early embryonic testis development by acting on early developing peritubular cells. In contrast, adult Sertoli cells do not appear to express NT-3, such that this is primarily an embryonic function of Sertoli cells.

However, these proteins also influence nonimmune cells and tissues. A number of different cytokines that have been shown to have functions in the testis are outlined next. The interleukins (ILs) are a family of cytokines produced by activated lymphocytes and macrophages. Interleukin 1 beta (IL-1β) is secreted by lymphocytes, whereas interleukin 1 alpha (IL-1α) is produced by a wide variety of nonimmune tissues. IL-1α has been shown to be expressed by Sertoli cells [86] and appears to influence spermatogenic cells [87]. The expression of IL-1α by Sertoli cells appears dependent on the presence of germ cells [88]. IL-1α can directly influence prepubertal Sertoli cell growth [89], transferrin gene expression [90], and lactate production [91]. Interestingly, Sertoli cells also express an antagonist form of IL-1α termed IL-1ra [92, 93]. These positive and negative acting forms of IL-1α are proposed to have a role in regulating cellular interactions between Sertoli cells and germ cells [92–94]. In contrast to IL-1α, IL-1β is not expressed by Sertoli cells [92, 93]. However, IL-1β can induce the expression of IL-1α by Sertoli cells [95]. In addition to IL-1α, Sertoli cells produce interleukin 6 (IL-6) in response to the autocrine actions of IL-1α [96–98]. Residual bodies induce Sertoli cell expression of IL-1α that in an autocrine manner induces Sertoli cell production of IL-6. Analysis of the hormonal regulation of IL-1 and IL-6 demonstrates distinct regulation [99] and secretion into different tubule compartments [100]. Although autocrine actions of IL-6 on Sertoli cells have been postulated [101, 102], an important function of Sertoli cell secreted interleukins is communication with the immune cells in the interstitial compartment [103, 104]. Leydig cells also produce these interleukins, probably for a similar function [105, 106]. Other cytokines produced by Sertoli cells include interferon α (IFN-α) [107] and α2-macroglobulin (α2MG) [108]. Both are proposed to interact with immune cells in the interstitium and possibly mediate other cell–cell interactions. Macrophage populations in the interstitium also are targets for macrophage migration inhibiting factor (MIF), which is primarily produced by Leydig cells [109, 110], but under appropriate conditions is also produced by Sertoli cells [109]. These and other cytokines [111] are likely to promote critical interactions with immune cells and Sertoli cell secreted cytokines will have a role in these cellular interactions.

F. Cytokines

G. Kit Ligand/Stem Cell Factor

Cytokines are defined as growth factors involved in immune cell communication and the immune system.

Kit ligand (KL), also known as stem cell factor (SCF), acts at the kit receptor and is known to be important

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for primordial germ cell viability and cell migration to the embryonic gonad. KL is required for normal germ cell proliferation and fertility of both the male and female. Sertoli cells are known to secrete KL and the spermatogonia express the kit receptor [112, 113]. Inhibition of KL reduces spermatogenesis and specifically the early mitotic events of germ cell development. Therefore, KL produced by Sertoli cells is needed in the embryo for germ cell survival and proliferation, early postnatally for gonocyte development [114], and in the adult for spermatogonia proliferation. This is a classic Sertoli cell–spermatogenic cell paracrine interaction in that the growth factor is produced only by one cell and receptor is in the target germ cell (Fig. 8.1). Gonadotropins regulate the expression of KL by Sertoli cells and various agents such as gonadotropinreleasing hormone (GnRH) [115], growth-hormone– releasing hormone (GHRH) [116], and somatostatin [117] can influence KL production. The physiological importance of this Sertoli cell secreted factor is shown in forms of male infertility. Impaired expression of kit [118] and mutated forms of KL [119] result in subfertility in human infertility patients. In addition to the Sertoli cell produced KL acting on spermatogonia, Leydig cells have been shown to potentially be a target during development or regeneration of the testis [120, 121].

H. Glial Cell–Derived Neurotrophic Factor Glial cell–derived neurotrophic factor (GDNF) is a member of the TGFβ superfamily and neurotropin family. GDNF has been shown to be secreted by Sertoli cells and acts on spermatogonia through the ret/GFR receptor specifically located on spermatogonia [122]. GDNF appears to promote spermatogonia stem cells to initiate development [122, 123]. Spermatogonia cell proliferation is influenced by GDNF [124, 125]. A related growth factor, neurturin, also has similar functions [125]. Therefore, similar to KL, GDNF is secreted by Sertoli cells to act on early-stage spermatogonia to promote cell proliferation and development (Fig. 8.1). Disruption of GDNF expression alters spermatogenesis [126] and overexpression of GDNF causes a nonmetastatic germ cell tumor by hyperproliferation of spermatogonia [127]. Although GDNF can influence Sertoli cell proliferation prepubertally, the spermatogonia cell is the primary target in the adult [128].

III. HORMONES Sertoli cells are endocrine cells in that they respond to hormones (e.g., FSH and androgen) and also produce hormones. In addition to the secretion of growth

factors by Sertoli cells, hormones are also important regulatory factors secreted. The major hormones produced by Sertoli cells are reviewed next.

A. Inhibin/Activin More than 70 years ago, Sertoli cells were shown to produce a regulatory agent termed inhibin [129] that inhibits FSH production by the pituitary [130, 131]. The two inhibin gene product subunits α and β form a dimer for inhibin, whereas a β homodimer produces activin that can stimulate FSH production. Both inhibin and activin are now known to have a wide variety of biological functions and are produced by a number of tissues. Sertoli cells under the control of FSH produce inhibin [132–135]. Although the major role of inhibin is to feed back information to the pituitary to regulate FSH production, local actions within the testis are also known [136, 137]. Sertoli cells can respond to activin to influence postnatal Sertoli cell proliferation [138]. Inhibin may influence germ cells [139, 140] and germ cells can influence the expression of inhibin by Sertoli cells [141]. The cellular interactions mediated by inhibin and activin in the testis are complicated by their expression by multiple cell types. Leydig cells can produce both inhibin and activin [142–145]. Recently, activin has been shown to be produced by peritubular cells and may regulate Sertoli cells through a paracrine interaction [146, 147]. Therefore, the specific paracrine and autocrine roles for inhibin and activin remain to be fully elucidated. However, the endocrine roles for these factors are better established. Abnormalities in inhibin levels correlating with abnormal pubertal development and testicular dysfunction have been documented [148–150].

B. Müllerian Inhibiting Substance Müllerian inhibiting substance (MIS), also termed anti-Müllerian hormone (AMH), is secreted by embryonic Sertoli cells and promotes the regression of the Müllerian duct and female reproductive tract development [151, 152]. MIS was first identified in the fetal and neonatal testes [153] and was subsequently cloned and found to be expressed by Sertoli cells [154–156]. MIS is a TGFβ family member. MIS expression is induced upon male sex determination by Sertoli cells and then declines to negligible levels in the adult [157, 158]. FSH inhibits the expression of MIS correlating with the decline during pubertal testis development [159]. MIS acts at specific receptors [151] to promote regression of the Müllerian duct and to regulate androgen production by Leydig cells [160]. The regulation of MIS expression has been examined on the transcriptional level

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and demonstrated that the SF1, GATA-4, and Sox8 transcription factors regulate the MIS promoter [161–163]. FSH, thyroid hormone, and androgen can regulate the expression of MIS in Sertoli cells [164, 165]. MIS/AMH appears to be a Sertoli cell secretory factor that has a critical role during embryonic development as an endocrine hormone. Genetic mutations in MIS or its receptor cause human mutations in premature developmental defects, persistent Müllerian ducts, and azoospermia [166–168].

C. Estrogen Sertoli cells express aromatase prepubertally so they can utilize androgen production by Leydig cells to produce estrogen. As Sertoli cells differentiate, aromatase expression drops such that negligible estrogen is produced in the adult testis. The knockout null mutations in the estrogen receptor (ER) demonstrated alterations in spermatogenesis and infertility [169]. The phenotype was found to be primarily mediated through rat testis and efferent duct abnormalities [170, 171]. However, estrogen production by Sertoli cells appears to also have direct effects on testis function and spermatogenesis [172]. Localization of the ERα demonstrated expression in Leydig cells and spermatogonic cells [173], whereas ERβ was found in Sertoli cells in the adult stage and in the fetal stage in Sertoli, Leydig, peritubular, and germ cells [174]. Estrogen can act on Sertoli cells and influence FSH actions [175]. As stated, FSH inhibits estrogen production and IGF-1 can alter these effects [16]. Sertoli cell secretion of estrogen appears to be primarily a role for the prepubertal period and as the testis develops in response to FSH, estrogen production becomes negligible. Estrogen may have a role in suppressing prepubertal androgen produced by Leydig cells and may influence germ cells at later stages. Specific roles for estrogen in the adult remain to be elucidated.

IV. OTHER REGULATORY FACTORS A. Uncharacterized Factors Several regulatory factors have been postulated to be produced by Sertoli cells and have specific roles in mediating interactions with other cell types. These factors have not been characterized beyond their biological activity. The first was seminiferous growth factor (SGF), which had mitogenic activity and was distinct from FGF and stimulated proliferation of several Sertoli cell lines [176–178]. The second was

Sertoli cell secreted growth factor (SCSGF), which had some similarities to TGFα and also stimulated the growth of several Sertoli cell lines [179]. These factors may well be some of the factors described earlier and have similar biological activities. A number of studies have suggested that Sertoli cells secrete a factor(s) that influences Leydig cell function and steroidogenesis. Abnormal physiological and environmental insults that affect Sertoli cell function have been shown to indirectly influence Leydig cell function [180–192]. Sertoli cell conditioned medium has been used to identify factors that stimulate cell proliferation and suppress steroidogenesis [193]. The evidence for such an activity is very convincing, but the factor(s) have not been characterized. Again a number of the Sertoli cell secreted factors mentioned earlier have this activity on Leydig cells (e.g., TGFα, bFGF). Sertoli cells have also been shown to produce factors such as erythropoietin [194], leukemia inhibitory factor (LIF), and ciliary neurotrophic factor (CNTF) [195], but the specific expression and actions of these factors needs to be assessed. Recently, the genomic microarray analysis of Sertoli cell gene expression has suggested the expression of a number of secreted regulatory factors [196]. Examination of these factors primarily provided the same list as that discussed earlier. It is likely that new factors secreted by Sertoli cells will be determined, but a more genomic approach should facilitate the identification and initial detection versus the more classic protein biochemistry approaches used in the past.

B. Lipids In addition to protein regulatory agents, nonprotein factors such as lipids are also found to have dramatic biological activities. Although many lipids have been primarily localized within the cell, some lipids such as lysophospholipids and thromboxanes are soluble and are known to bind specific G-protein coupled receptors on the surface of cells [197, 198]. These lipid-induced receptor-mediated events alter cellular functions in a similar manner to protein growth factors [197]. The testis has unique forms of metabolic enzymes generating these lipids [199–201], such that lipid factors may be secreted by Sertoli cells. Sertoli cells have been shown to produce soluble sphingosine molecules and other lysophospholipids [202–204]. The specific lipids secreted and functions remain to be elucidated, but due to the important functions of these extracellular soluble lipids it is likely they will be important regulators of testis function and spermatogenesis.

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V. SUMMARY The secretion of regulatory factors by Sertoli cells is critical for normal testis development and function. Abnormalities in the expression or action of these factors often result in infertility or endocrine defects. As shown in Figure 8.1 and Table 8.2, a number of growth factors are secreted by Sertoli cells to influence spermatogenic cells. Because Sertoli cells are the principal regulatory cells for spermatogenesis, it is not surprising that many of the growth factor’s primary targets are germ cells. Since the original review of this topic, several major observations have been made regarding Sertoli cell secreted growth factors [2]. Knowledge that the production of KL and GDNF by Sertoli cells directly regulates spermatogonia cell proliferation and development represents a significant advance in our understanding of the control of the initiation of the spermatogenic wave and regulation of the spermatogenic stem cell population. Recently, human male infertility conditions have been linked to abnormalities in the expression and action of these factors. Other specific factors that have the characteristics of the growth factor being expressed by Sertoli cells and actions only on a specific stage of germ cells are likely to be identified. The number of

TABLE 8.2 Sertoli Cell Secreted Growth Factors Growth factor

Site of action

Function

Insulin-like growth factor I (IGF-I)

Germ cells Sertoli cells Leydig cells

Cell cycle progression and growth (i.e., S-phase DNA synthesis)

Insulin-like growth factor II (IGF-II)

Germ cells

Cell–cell signaling

Transforming growth factor α (TGFα)

Germ cells Sertoli cells Peritubular cells

Cell growth initiation

Transforming growth factor β (TGFβ)

Germ cells Sertoli cells Peritubular cells

Growth inhibition and increase in cell differentiated function

Basic fibroblast growth factor (bFGF)

Germ cells Sertoli cells Peritubular cells

Cell growth initiation

Neurotropin 3 (NT-3)

Peritubular cells

Cell migration (i.e., mesonephros) and growth

Interleukin 1 and 6 (IL-1 and IL-6)

Germ cells

Cell–cell signaling

Kit ligand/stem cell factor (KL/SCF)

Germ cells Leydig cells

Cell growth (i.e., spermatogonial)

Glial cell–derived neurotrophic factor (GDNF)

Germ cells

Cell growth (i.e., spermatogonial stem cells)

TABLE 8.3 Sertoli Cell Secreted Hormones Hormone

Site of action

Function

Inhibin

Pituitary

Reduce FSH production

Müllerian inhibiting substance/antiMüllerian hormone (MIS/AMH)

Müllerian duct

Regress female reproductive tract development

Estrogen

Leydig cells

Reduce steroidogenesis and cell–cell signaling

growth factors listed in Table 8.2 will likely grow further during the next decade as well. The microenvironment within the testis and local cell–cell interactions mediated by regulatory factors such as those listed in Table 8.2 require elucidation for us to understand how hormones regulate testis function and how therapeutic treatments for male infertility can be developed. The Sertoli cells also secrete several critical hormones, as listed in Table 8.3. Sertoli cells are critical to male reproductive endocrinology because they are a target for several hormones (e.g., FSH and testosterone) and they produce hormones. Feedback pathways are always required in a biological process and endocrine system, so it is not surprising that a critical inhibitory feedback protein like inhibin is produced. In addition, the negative feedback estrogen provides prepubertally to suppress Leydig cell androgen production is also anticipated. The information that has developed during the past decade [2] indicates that these factors also have other roles in mediating regulatory interactions between cells within the testis. Although the specific functions of the local actions of the inhibin, activin, and estrogen need to be elucidated, clearly they have local paracrine functions independent of the endocrine role of these factors. This will likely be an intense area of research for a number of years.

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C H A P T E R

9 Proteases and Protease Inhibitors MARTIN CHARRON AND WILLIAM W. WRIGHT Division of Reproductive Biology, Department of Biochemistry and Molecular Biology, Bloomberg School of Public Health, Johns Hopkins University, Baltimore, Maryland

biochemical networks. These general discussions are prompted by our conviction that this information is a necessary foundation for subsequent discussions of the expression and potential functions of these proteins within the seminiferous epithelium. Sections II, III, and IV conclude with a review of the reported reproductive phenotypes observed in knockout mouse models. The reasons why, ultimately, a thorough analysis of the testes of these mice needs to be performed are described in Section V, and future research directions are discussed in Section VI. Finally, resources available on the World Wide Web are listed in Section VII.

I. II. III. IV. V.

INTRODUCTION THE METZINCINS THE PLASMINOGEN SYSTEM THE CATHEPSINS MORE THAN MEETS THE EYE: LESSONS FROM CATHEPSIN L–DEFICIENT MICE VI. FUTURE DIRECTIONS VII. RESOURCES ON THE WORLD WIDE WEB References

I. INTRODUCTION A. Scope of This Chapter

B. Proteolysis, Proteases, and Protease Inhibitors

During spermatogenesis, the seminiferous epithelium undergoes restructuring events that are essential for the production of sperm. These events likely involve the proteolysis of different protein substrates, controlled by the opposing actions of proteases and their complementary inhibitors. This chapter summarizes current knowledge about the expression and potential functions of proteases and protease inhibitors that are synthesized in the seminiferous epithelium. Particular emphasis is given to products of Sertoli cells. In this section, information about proteases and their targets in the seminiferous epithelium are discussed. Sections II, III, and IV examine proteases and their complementary inhibitors in the metzincin superfamily, the plasminogen system, and the cathepsins, respectively. Each section includes a discussion of the general biochemistry of these proteases and their respective inhibitors and, where appropriate, their integration into functional SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

Proteolysis is one of several important posttranslational modifications that a protein can undergo during its lifetime. Whereas modifications such as glycosylation and phosphorylation are reversible, proteolysis is irreversible and the only means available for rebuilding the intact protein is to translate mRNA. Proteolysis can be divided roughly into two types: limited proteolysis (e.g., the generation of mature forms from protein precursors) and complete proteolysis (breakdown) of proteins to amino acids. Proteolysis is the consequence of the opposing actions of proteases and their complementary inhibitors. Ubiquitously expressed in all biological tissues and fluids, proteases are enzymes capable of catalytically cleaving peptide bonds in proteins (reviewed in [1]). All proteases share in common the general mechanism of a nucleophilic attack on the carbonyl carbon of

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an amide group. Proteases are divided into two groups: proteinases (endopeptidases) and peptidases (exopeptidases). Proteinases catalyze the hydrolysis of internal bonds in polypeptides. They are divided into four classes: aspartic, metallo, cysteine, and serine proteases (reviewed in [2]). The first two classes of enzymes utilize aspartate residues and heavy metals, respectively, to immobilize and polarize a water molecule so that the oxygen atom in water becomes the nucleophile. The latter two classes of enzymes utilize their hydroxyl- and sulfhydryl-side chains, respectively, as nucleophiles. Peptidases attack only peptide bonds localized at or near the amino or carboxy terminus of peptide chains (reviewed in [3]). Accordingly, they are divided into two classes: aminopeptidases and carboxypeptidases.

C. Proteolysis in the Seminiferous Epithelium As a consequence of their enzymatic activities, proteases play major roles in the differentiation and function of all cells and tissues, and they can have a profound effect on the interactions of cells with their immediate microenvironments. These enzymes are essential for the remodeling or degradation of basement membranes and, consequently, the movement of cells within an epithelium [4, 5]. Specific proteases can also modify cell–substrate and cell–cell interactions by degrading or activating cell adhesion molecules, procytokines, and cell surface receptors [6–12]. A protease often acts in cooperation with a network of other proteins, including their complementary inhibitors. Numerous proteases and their complementary inhibitors are present in the seminiferous epithelium. Aspects of the dynamic morphology of this epithelium argue strongly that these proteins are functionally important to the process of spermatogenesis. Structures in the seminiferous epithelium that are known or proposed targets for proteases are illustrated in Figure 9.1A. Major targets are residual bodies (Fig. 9.1B), which are shed from spermatids at spermiation, engulfed by Sertoli cells, and then degraded [13, 14]. This proteolytic process is potentially important because it destroys any autoantigens contained in the residual bodies and, thus, prevents their release into the general physiological environment [15]. A second potential target is the ectoplasmic specialization, which tethers spermatids to Sertoli cells (Fig. 9.1C). Removal of this structure is one of the last changes in germ cell–Sertoli cell interactions that occurs prior to spermiation [16]. The basal ectoplasmic specializations and tight junctions forming the blood–testis barrier (Fig. 9.1D) have also been identified as potential

targets for proteases and their inhibitors [17–19]. Restructuring of this barrier is required for preleptotene spermatocytes to pass from the basal to the adluminal compartment of the seminiferous tubule. Finally, as with all other epithelia, proteases and their inhibitors must be important to the remodeling and turnover of the basement membrane of the seminiferous tubule (Fig. 9.1D). Such turnover might be particularly crucial during the prepubertal growth of this epithelium [20]. Relevant to this function, reduced protease activity may be the immediate cause of the thickened basement membrane of the seminiferous epithelium in aged rats [21]. The next three sections examine proteases and their complementary inhibitors in the metzincin superfamily (Section II), the plasminogen system (Section III), and the cathepsins (Section IV). Table 9.1 lists the names and EC numbers of the proteases discussed in these sections and Table 9.2 provides the chromosomal localization of the relevant genes in humans, mice, and rats. Finally, a note on terminology: Because all the proteases discussed in this chapter are endopeptidases, the term proteinase will be used from this point forward.

II. THE METZINCINS Members of the metzincin superfamily are metalloproteinases that require zinc at their catalytic sites. These proteinases are characterized by three conserved histidine residues at the catalytic site and a loop (the “metturn”), which faces this site. This loop binds a water molecule required for catalysis [5, 22]. The metzincin superfamily encompasses four families of proteinases, two of which, the matrix metalloproteinases (MMPs) and the ADAM-TS (A Disintegrin and Metalloproteinase Domain with Thrombospondin motifs), are expressed by Sertoli cells. The actions of the proteinases in these families are opposed by the tissue inhibitors of metalloproteinases (TIMPs), which are also expressed by Sertoli cells. This section examines the expression of specific MMP, ADAM-TS, and TIMP family members by Sertoli cells and identifies potential substrates in the testis for individual proteinases. This section ends with a synopsis of the reproductive phenotypes of male mice deficient in a specific MMP, ADAM-TS, or TIMP.

A. Matrix Metalloproteinases and Their Inhibitors 1. Introduction The MMPs expressed by Sertoli cells are matrix metalloproteinase-2 (MMP2), matrix metalloproteinase9 (MMP9), and membrane-type matrix metalloproteinase 1 (MT1-MMP) [19, 23–26]. All MMPs are active at neutral pH and may use as substrates basement

B

C

D

AA

FIGURE 9.1 Structures in the seminiferous epithelium that are known or proposed targets for proteinases. (A) Drawing of a rat Sertoli cell and associated spermatocytes and spermatids at stage VIII of the cycle of the seminiferous epithelium. Known or potential targets are labeled 1, 1’, 2, 3, and 4. Structures 1 and 1’ are residual bodies. The arrows labeled 2 identify ectoplasmic specializations, which mediate the adhesion of spermatids to the Sertoli cell. The arrows labeled 3 point to the blood–testis barrier, formed by the combination of basal ectoplasmic specializations and tight junctions. Structure 4 is the basement membrane of the seminiferous epithelium. PLS, preleptotene spermatocyte; PS, pachytene spermatocyte; S, step 8 spermatid. (Source: Modified from Figure 18 of Dym, M., and Fawcett, D. W. Biol. Reprod. Vol. 3, The blood-testis barrier in the rat and the physiological compartmentation of the seminiferous epithelium, pp. 308–326. Copyright 1970 Society for the Study of Reproduction.) (B to D) Electron micrographs of the structures illustrated in panel (A). In panel (B), the arrow labeled 1 points to a residual body that has been engulfed by a Sertoli cell. The arrows labeled 1’ identify residual bodies that have fused with lysosomes in a Sertoli cell (final magnification = 3750 ×). Panel (C) shows an ectoplasmic specialization (arrow labeled 2) facing a step 8 spermatid (final magnification = 18,750 ×). The basement membrane (arrow labeled 4) and the blood–testis barrier (arrow labeled 3) in the basal portion of a Sertoli cell are shown in panel (D) (final magnification = 12,360 ×). (Electron micrographs courtesy of Janet S. Folmer.)

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Martin Charron and William W. Wright TABLE 9.1 Proteinase Nomenclature

Name used in the text

EC number Alternative names

ADAM-TS4

3.4.24.14

TABLE 9.2 Chromosomal Localization of Various Genes that Encode Proteinases Expressed in Sertoli Cells Chromosome

Cathepsin A

3.4.16.5

Procollagen I N-endopeptidase, ADAMTS2

Name

Human

Mouse

Rat

Protective protein, lysosomal carboxypeptidase A

ADAM-TS4

5qter

11 (B1.3)

8q24a 10q21a

Cathepsin A

20q13.1

2 (96.0 cM)

3q42b

Cathepsin B

3.4.22.1

None

Cathepsin B

8p22

14 (28.0 cM)

15p12

Cathepsin D

3.4.23.5

None

Cathepsin D

11p15.5

4 (50.0 cM)

1q37

Cathepsin K

3.4.22.38

Cathepsin O, O2, or X

Cathepsin K

1q21

3 (47.9 cM)

2q34

Cathepsin L

3.4.22.15

None

Cathepsin L

9q21q22

13 (30.0 cM)

17p14

Cathepsin S

3.4.22.27

None

Cathepsin S

1q21

3 (42.7 cM)

2q34

Cathepsin V

3.4.22.43

Cathepsin L2 or U

Cathepsin V

9q22.2

Absent

Absent

Matrix metalloproteinase 2 (MMP-2)a

3.4.24.24

Gelatinase A

MMP-2

16q13-q21

8 (44 cM)

19p11

Matrix metalloproteinase 9 (MMP-9)

3.4.24.35

Gelatinase B

MMP-9

20q11.2-q13.1

2 (96 cM)

3q42

MT1-MMP

14q11-q12

14 (12.5 cM)

15p13

Membrane-type matrix metalloproteinase 1 MT1-MMP

None

Matrix metalloproteinase 14, MMP-14

Plasmin

3.4.21.7

Fibrinase, fibrinolysin, activated plasminogen

Tissue plasminogen activator (tPA)

3.4.21.68

T-plasminogen activator

Urokinase

3.4.21.73

Urinary plasminogen activator

aAbbreviations

used in the text are in parentheses

membrane proteins and secreted or transmembrane proteins involved in cell–cell interactions [5]. Consequently, within the seminiferous epithelium, some substrates of the MMPs may be localized to sites of specific interactions between opposing Sertoli cells, between Sertoli cells and spermatogenic cells, and between Sertoli cells, spermatogonia, or preleptotene spermatocytes and the basement membrane (Fig. 9.1). The 25 vertebrate MMPs can be divided into two major types, membrane anchored and secreted [5]. Nineteen MMPs are secreted, including MMP2 and MMP9; four are integral membrane proteins (MT-MMPs); and two are attached to the cell surface by glycosyl-phosphatidylinositol (GPI) anchors [4, 5, 11]. The secreted MMPs are released from cells as inactive proenzymes, which are then activated by cleavage of the N-terminal prodomain by other proteinases. In most cases, the prodomains of membrane-anchored MMPs are cleaved intracellularly by a proprotein convertase. Thus, they reach the cell surface as functional enzymes [4]. The MMP inhibitors, the TIMPs, form a threedimensional wedge that fits into the catalytic and substrate binding sites of an MMP [27]. Multiple TIMPs can act as high-affinity inhibitors of a given

Plasminogen

6q26

17 (7.3 cM)

1p11

tPA

8q12

8 (9.0 cM)

16q12.5

Urokinase

10q24

14 (2.5 cM)

15p16

aSequences

similar to ADAM-TS4 precursor found on two chromosomes. bSequence similar to the mouse protective protein precursor.

MMP (Ki ≤ 1 nM) [28, 29]. There is some selectively in TIMP–MMP interactions, however. TIMP-2 and -3 but not TIMP-1 are effective inhibitors of cell surface MT1-MMP [27]. TIMP-1 and -2 have been reported to be expressed by Sertoli cells [18, 23]. TIMP-3 is also expressed in the testis and is represented in a Sertoli cell cDNA library [30] (see Section VII for the library’s URL address). The proteinase targets expressed in the testis of TIMP-1, -2, and -3 are listed in Table 9.3. TABLE 9.3 Matching Members of the Metzincin Superfamily to Their Complementary Inhibitors Expressed in the Testis MT1-MMP MMP-2 MMP-9 ADAM-TS4 ADAM-TS5

TIMP-1

X

TIMP-2

X

TIMP-3

X

X X

X

Note: Information is compiled from the following references: Sternlicht, M. D., and Werb, Z. (2001). How matrix metalloproteinases regulate cell behavior. Annu. Rev. Cell. Dev. Biol. 17, 463–516; Brew, K., Dinakarpandian, D., and Nagase, H. (2000). Tissue inhibitors of metalloproteinases: evolution, structure and function. Biochim. Biophys. Acta 1477, 267–283; Kashiwagi, M., Tortorella, M., Nagase, H., and Brew, K. (2001). TIMP-3 is a potent inhibitor of aggrecanase 1 (ADAM-TS4) and aggrecanase 2 (ADAM-TS5). J. Biol. Chem. 276, 12501–12504.

Chapter 9 Proteases and Protease Inhibitors

Cell surface MT1-MMP is potentially important to the function of Sertoli cells for two reasons: (1) It acts in a cell’s immediate microenvironment and (2) it forms part of the cell surface sites that bind pro-MMP2 and process the proenzyme to an active proteinase. Thus, these sites concentrate MMP2 enzyme activity in the restricted physiological microenvironment around a cell. Two types of MMP2 binding sites have been defined and their components are expressed by Sertoli cells. The first type of binding site is created by a TIMP-2–MT1-MMP complex (see [4, 5] for details). The second type of binding site for pro-MMP2 is formed by the binding of any MT-MMP to a member of a transmembrane family of proteins, the claudins [31]. Claudin 11 is an essential component of the blood–testis barrier [32, 33], and it is interesting to speculate that claudin-11 concentrates MT1-MMP and MMP2 at the blood–testis barrier. This raises the possibility that MMP2 and MT1-MMP, bound to claudin-11, participate in the restructuring of the blood–testis barrier that is required for preleptotene spermatocytes to pass from the basal to the adluminal compartment of the seminiferous epithelium. 2. Expression in Sertoli Cells a. MT1-MMP In addition to Sertoli cells, MT1-MMP transcripts are expressed by peritubular myoid cells, Leydig cells, spermatocytes, and spermatids (Fig. 9.2A). Western blot analysis identified both pro- and mature MT1-MMP (63 and 60 kDa, respectively) in these cells (Fig. 9.2B). Additionally, a 45-kDa form of MT1-MMP was present in spermatogenic cells. This smaller form has been shown in other cell types to result from the N-terminal cleavage of MT1-MMP, which inactivates the enzyme [34]. Immunocytochemical analysis argues that the expression by Sertoli cells of MT1-MMP changes during testis maturation. In sexually immature rats, when the basement membrane is being continuously remodeled, all Sertoli cells express MT1-MMP [23]. In contrast, available data suggest that in sexually mature rats, this enzyme is expressed only by a subset of Sertoli cells. This raises the possibility that the expression of MT1-MMP is regulated in a stage-specific manner by the surrounding germ cells, as has been shown for other proteases [35, 36]. Finally, note that folliclestimulating hormone (FSH) does not stimulate expression of MT1-MMP by Sertoli cells [23]. b. MMP2 Unlike MT1-MMP, MMP2 is only expressed by Sertoli cells, peritubular myoid cells, and Leydig cells [26] (Fig. 9.3A). In Sertoli cells, expression of MMP2

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A

B

FIGURE 9.2 Expression of MT1-MMP mRNA and protein in rat testicular cells. (A) RT-PCR analysis of MT1-MMP mRNA expression in liver, Sertoli cells (Sc), peritubular myoid cells (Pc), Leydig cells (Lc), spermatocytes (SPC), spermatids (SPT), and a crude germ cell preparation (CGC). (B) Western blot analysis of MT1-MMP in purified testicular cells. Cell lysates were obtained from peritubular myoid cells isolated from 5-day-old rats (C.Pc) and Sertoli cells (C.Sc). Also analyzed were freshly isolated Sertoli cells (Fl.Sc), spermatocytes (Fl.SPC), spermatids (Fl.SPT), and mixed germ cells isolated in the presence or absence of trypsin [Fl.CGc (+) and Fl.CGc (–), respectively]. (Source: Figure 3 of Longin, J., Guillaumont, P., Chauvin, M.-A., Morera, A.-M., and Le Magueresse-Battistoni, B., J. Cell Sci., Vol. 114, MT1-MMP in rat testicular development and the control of Sertoli cell pro-MMP2 activation, pp. 2125–2134. Copyright 2001 Company of Biologists, Ltd.)

transcripts and enzyme activity are highly regulated. FSH and cAMP stimulate MMP2 mRNA expression by cultured, rat Sertoli cells isolated from 20-day-old rats [23, 24, 26] (Fig. 9.3B). FSH has a small effect on MMP2 protein secretion but a large effect on MMP2 enzyme activity [26]. This larger effect on enzyme activity is likely due to the simultaneous stimulation by FSH of TIMP-2 expression by Sertoli cells. Recall that TIMP-2 in cooperation with MT1-MMP creates cell surface binding and activation sites for pro-MMP2. Coculture of germ cells with Sertoli cells also increases activation of pro-MMP2 (Fig. 9.3C). Longin et al. [26] proposed that this stimulatory effect of germ cells was due to their expression of cell surface MT1-MMP. Thus, the amount of active MMP2 in the seminiferous epithelium is determined first by the hormonal stimulation of Sertoli cells and then by the total amount of cell surface MT1-MMP on both Sertoli and germ cells. The stimulatory effect of FSH on MMP2 enzyme activity is opposed by tumor necrosis factor α (TNFα), which is produced by spermatids and binds to

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of pro-TNFα by MMPs may be part of a feedback mechanism that inhibits MMP2 expression in the seminiferous epithelium. c. MMP9 MMP9 is secreted by Sertoli cells and inhibits the formation of occluding junctions between these cells in vitro. Contrary to its inhibitory effect on MMP2, TNFα stimulates expression of MMP9 [19]. d. TIMP-1, TIMP-2, and TIMP-3

FIGURE 9.3 Expression and regulation of MMP-2 and TIMP-2 in rat testicular cells. (A) RT-PCR analysis of MMP-2 and TIMP-2 expression in Sertoli cells (Sc), peritubular myoid cells (Pc), Leydig cells (Lc), spermatocytes (SPC), spermatids (SPT), and a crude germ cell preparation (CGC). (B) FSH and cAMP stimulate expression of MMP-2 mRNA in Sertoli cells isolated from sexually immature rats. Sertoli cells were isolated from 20-day-old rats and cultured in the absence or presence of FSH (50 ng/mL) or 8-bromo-cAMP (10–3 M) for 48 hr. MMP-2 mRNA levels were then estimated by Northern blot analysis and data normalized to GAPDH mRNA. (C) MMP-2 activation in Sertoli cell–germ cell cocultures. Data show MMP-2 enzyme activity in 80-μg (1) and 40-μg total protein (2) in culture media. Enzyme activity was estimated by sodium dodecyl sulfate– polyacrylamide gel electrophoresis followed by gelatin zymography. The sizes of the lytic bands are indicated on the right. The band at 62 kDa is the appropriate size for activated MMP-2. Sc, Sertoli cells; Sc+CGc, Sertoli cell–germ cell cocultures. (Source for panels A–C: Figures 5B, 6B, and 10C of Longin, J., Guillaumont, P., Chauvin, M.A., Morera, A.M., and Le Magueresse-Battistoni, B., J. Cell Sci., Vol. 114, MT1-MMP in rat testicular development and the control of Sertoli cell pro-MMP2 activation, pp. 2125–2134. Copyright 2001 Company of Biologists, Ltd.) (D) The expression of TIMP-2 transcripts is stimulated by FSH and inhibited by TNFα. Sertoli cells were treated for 48 hr with FSH (50 ng/mL) and/or TNFα (50 ng/mL). The 3.5-kb TIMP-2 and 0.9-kb TIMP-2 transcripts were assayed by Northern blot analysis, and data quantified by scanning densitometry. Data are expressed as the ratio of each TIMP-2 transcript to GAPDH mRNA. Data with different superscripts differ statistically. (Source: Figure 9.6 of Longin, J., and Le Magueresse-Battistoni, B., Mol. Cell. Endocrinol., Vol. 189, Evidence that MMP-2 and TIMP-2 are at play in the FSHinduced changes in Sertoli cells, pp. 25–35. Copyright 2002 Elsevier.)

receptors located on Sertoli cells [23, 37]. TNFα is secreted as a procytokine and MT1-MMP, MMP2, and MMP9, all expressed in Sertoli cells, have the capacity to proteolytically process this inactive precursor to an active cytokine [5]. Thus, processing and activation

TIMP-1, a secreted inhibitor of MMP9 enzyme activity (see Table 9.3), is a product of Sertoli cells and peritubular myoid cells but not of germ cells [30]. In mouse testes, levels of TIMP-1 transcripts increase threefold from 18 to 29 days of age and then decrease threefold by 41 days of age [38]. Thus, TIMP-1 mRNA levels are maximal in immature seminiferous tubules. TIMP-1 mRNA expression in Sertoli cells is also stimulated by FSH, interleukin 1 alpha (IL-1α), and residual bodies. Additionally, secretion of TIMP-1 is stimulated by cAMP [30, 39]. TIMP-2 mRNA is expressed by Sertoli cells, Leydig cells, peritubular myoid cells, and germ cells (Fig. 9.3A), and the protein is secreted by cultured Sertoli cells [23]. Thus, Sertoli cells express MT1-MMP, MMP2, and TIMP-2, all of which are required for cell surface localization and activation of MMP2. As with TIMP-1 and MMP2, FSH stimulates both expression of TIMP-2 mRNA and secretion of this protease inhibitor by Sertoli cells (Fig. 9.3D). Similar to MMP2, the stimulatory effect of FSH on TIMP-2 expression by Sertoli cells is opposed by TNFα [23, 26] (Fig. 9.3D). This dual inhibitory effect of TNFα on both MMP2 and TIMP-2 expression makes this cytokine an important candidate to regulate cell surface MMP2 activity in the seminiferous epithelium. TIMP-3 is represented in a cDNA library prepared from murine Sertoli cells (see Section VII for the URL address of the library). In the mouse, total testis levels of TIMP-3 mRNA increase approximately threefold from 18 to 49 days of age [38]. This result is consistent with an inherent effect of testis maturation on TIMP-3 expression by Sertoli cells and/or with an effect of increasing numbers of spermatogenic cells on expression of this gene by Sertoli cells. The expression of TIMP-3 by germ cells represents another alternative explanation for that result.

B. Potential Substrates for MMP2, MMP9, and MT1-MMP in the Seminiferous Epithelium Biochemical analyses demonstrate that each MMP has many potential substrates and that multiple MMPs may cleave the same substrate [5]. A major class of

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Chapter 9 Proteases and Protease Inhibitors

substrates for MMPs are the collagens. Table 9.4A identifies the collagens that are degraded by MT1-MMP, MMP2, and MMP9. The principal type of collagen in the basement membrane of the seminiferous epithelium is type IV [40, 41], which is a substrate for MMP2 and MMP9. Laminin and fibronectin, present in the basement membrane of the seminiferous tubule, are also substrates for these two secreted MMPs [5, 11, 40, 41]. Thus, MT1MMP–TIMP-2 complexes, by activating pro-MMP2 and mature MMP2 by degrading specific basement membrane components, may determine the rate of turnover of the basement membrane of the seminiferous epithelium. As has been documented in other epithelia, such basement membrane turnover should release trapped growth factors, potentially stimulating Sertoli cells or germ cells that express the appropriate receptors [5, 11, 12]. The MMPs have another mechanism for changing the local concentrations or activities of cytokines and growth factors. This is achieved by processing inactive precursors or by degrading the binding partners of an active growth factor or cytokine. It is, therefore, noteworthy that the testis expresses pro-TNFα, protransforming growth factor β (pro-TGFβ), and pro-IL1β, all known substrates for specific MMPs expressed in the seminiferous epithelium [37, 42–44]. Additionally, Sertoli cells express a receptor for fibroblast growth factor (FGF) and an insulin-like growth factor (IGF) binding protein, which are degraded by MMP2 (Table 9.4B) [45, 46]. Thus, MMP2, MMP9, MT1-MMP, and their corresponding TIMPs are predicted to modify both the architectural and paracrine environments of the seminiferous epithelium. As illustrated in Table 9.4B, many proteins in the seminiferous epithelium are substrates for more than one of these proteinases, possibly creating considerable redundancy in this system. The potential importance of MMPs and TIMPs to the seminiferous epithelium is also suggested by the effects of hormones, growth factors, and cell–cell interactions on their expression by Sertoli cells. Such regulated expression, discussed earlier in this chapter, suggests that these proteinases or inhibitors have a specific physiological function in this tissue.

C. The ADAM-TS Family 1. Introduction To date, 16 distinct mammalian ADAM-TS proteases have been identified in the mammalian genome. Type 1 collagen and aggrecan, a major component of the extracellular matrix of cartilage, are major substrates of this family [47, 48]. However, neither of these are known components of the basement membrane of the seminiferous epithelium.

TABLE 9.4 Substrates of MT1-MMP, MMP-2, and MMP-9 A. Substrate Collagens for MT1-MMP, MMP-2, and MMP-9 Metalloproteinase MT1-MMP

MMP-2

MMP-9

I, II, III

I, III, IV, V, VII, X, XI

IV, V, XI

Collagens that are known substrates for each metalloproteinase

B. Substrates of MT1-MMP, MMP-2, and MMP-9 Identified in the Seminiferous Epithelium Biochemical substrates identified in seminiferous tubules MT1-MMP

MMP-2

MMP-9

Collagen IV

X

X

Laminin

X

X

Fibronectin

X

Pro-MMP-2

X

α2-Macroglobulin

X

Plasminogen Pro-TNFα

X X X X

X

X

X

Pro-TGFβ

X

X

FGF R1

X

IGFBP3

X

Pro-IL-1β

X

X

Information is compiled from the following references: Sternlicht, M. D., and Werb, Z. (2001). How matrix metalloproteinases regulate cell behavior. Annu. Rev. Cell. Dev. Biol. 17, 463–516; McCawley, L. J., and Matrisian, L. M. (2001). Matrix metalloproteinases: they’re not just for matrix anymore! Curr. Opin. Cell. Biol. 13, 534–540; De, S. K., Chen, H. L., Pace, J. L., Hunt, J. S., Terranova, P. F., and Enders, G. C. (1993). Expression of tumor necrosis factor-alpha in mouse spermatogenic cells. Endocrinology. 133, 389–396; Janitz, M., Fiszer, D., Lukaszyk, A., Skorupski, W., and Kurpisz, M. (1995). Analysis of mRNa expression for interleukin-1 genes on human testicular cells. Immunol. Lett. 48, 139–143; Caussanel, V., Tabone, E., Hendrick, J. C., Dacheux, F., and Benahmed, M. (1997). Cellular distribution of transforming growth factor betas 1, 2, and 3 and their types I and II receptors during postnatal development and spermatogenesis in the boar testis. Biol. Reprod. 56, 357–37; Skinner, M. K., and Mullaney, B. P. (1993). Transforming growth factor-beta (beta 1, beta 2, and beta 3) gene expression and action during pubertal development of the seminiferous tubule: potential role at the onset of spermatogenesis Mol. Endocrinol. 7, 67–76; Yazama, F., Esaki, M., and Sawada, H. (1997) Immunocytochemistry of extracellular matrix components in the rat seminiferous tubule: electron microscopic localization with improved methodology. Anat. Rec. 248, 51–62.

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2. Expression in Sertoli Cells Three members of the ADAM-TS family, ADAM-TS1, 2, and 5 are represented in a cDNA library prepared from murine Sertoli cells. Additionally, ADAM-TS4 mRNA is expressed in the testis, but its cellular source has not yet been identified (see Section VII for URL addresses of both libraries). However, data indicate that in other cell types, ADAM-TS4 expression is stimulated by TGFβ. It is, therefore, noteworthy that proTGFβ is secreted by both Sertoli and peritubular myoid cells, that the procytokine is processed by MMPs (Table 9.4), and that Sertoli cells and germ cells express receptors for TGFβ [43, 44, 49]. Thus, ADAM-TS4 expression in the testis may be regulated by the combined activities of TGFβ and MMPs.

D. Reproductive Phenotypes Observed in Knockout Mice As already discussed, Sertoli cells express three MMPs with overlapping substrate specificities and thus, potentially redundant biological functions. The minimal data available from knockout animals reinforce this theory. Male mice that lack MMP2 or TIMP-2 are fertile [50, 51]. Thus, high pericellular concentrations of MMP2 are not required for the production of the minimum number of sperm necessary for fertility. However, as discussed in Section V, mice deficient in a specific proteinase may be fertile but nonetheless produce fewer sperm than wild-type mice. Thus, it is currently unknown whether deficiencies in TIMP-2 or MMP2 have an adverse effect on spermatogenesis. In addition, the genes that encode MMP9, TIMP-1, and TIMP-3 have also been deleted in mice without any reported effects on male fertility [52–54]. Because a major function of MT1-MMP is to concentrate MMP2 at the cell surface, one might anticipate that MT1-MMP–deficient mice would be fertile. It was surprising, therefore, that MT1-MMP–deficient male mice were dwarfs with craniofacial and skeletal abnormalities and “showed no signs of sexual maturation” [55]. Although the testicular histology of the MT1MMP–deficient mice was not evaluated, the failure of testis maturation suggests that as-yet-unidentified substrates still exist for this transmembrane metalloproteinase. Alternatively, the apparently widespread physiological deficits of these animals may have been sufficiently severe to inhibit the pubertal increase in leutenizing hormone (LH) secretion, depriving the animals of the testosterone required for testis maturation. Finally, because a major substrate for ADAM-TS4 is procollagen I and because collagen I is not a major

component in the basement membrane of any epithelia, including the seminiferous epithelium, it would be reasonable to expect that ADAM-TS4 does not have a major function in the seminiferous epithelium [40, 41, 48]. Thus, the finding that ADAM-TS4–deficient male mice are infertile was unexpected [56]. Histological evaluation of the epididymis showed very few sperm, and cross sections of seminiferous tubules revealed a large number of round spermatids but few elongate spermatids (Fig. 9.4A). Thus, ADAM-TS4, like MT1MMP, may have unidentified substrates within the seminiferous tubule whose proteolytic processing is required for fertility. In summary, data from these knockout mice argue that MMP1, MMP9, TIMP-1, TIMP-2, and TIMP-3 are

A

B

FIGURE 9.4 Morphological comparison of a seminiferous tubule from ADAM-TS4-deficient mice (A) with a tubule from wild-type mice (B). Testes were perfusion fixed with 4% buffered paraformaldehyde, embedded in glycol methacrylate, and 1.5-μm histological sections stained with 1% toluidine blue. [Source: Figure 7 of Li, S. E., Arita, M., Fertala, A., Bao, Y., Kopen, G. C., Langsjo, T. K., Hyttinen, M. M., and Helminen, H. J., and Prockop, D. J. Biochem. J., Vol. 355, Transgenic mice with inactive alleles for procollagen N-proteinase (ADAMTS2) develop fragile skin and male sterility, pp. 271–278. Copyright 2001 The Biochemical Society.]

Chapter 9 Proteases and Protease Inhibitors

not obligatory for spermatogenesis. In contrast, MT1MMP and ADAM-TS4 are essential for this process.

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biological role in the seminiferous epithelium and, thus, the process of spermatogenesis. 1. tPA and Plasminogen

III. THE PLASMINOGEN SYSTEM A. Components of the Plasminogen System The two plasminogen activators, urokinase and tissue-type plasminogen activator (tPA), are secreted proteinases that are active at neutral pH. Both are expressed in the seminiferous epithelium [57]. Thus, like the metzincin superfamily of proteinases, the plasminogen system is active in the extracellular environment of this tissue and may be active in specific morphological sites involved in cell–cell and cell– basement membrane interactions (Fig. 9.1). Both urokinase and tPA have one major substrate, plasminogen, which is also expressed in the seminiferous epithelium [58, 59]. Urokinase and tPA cleave a single peptide bond in plasminogen, leaving the two chains of the enzymatic product, plasmin, joined by disulfide bridges [60]. A major function of the plasminogen system is the degradation of fibrin and, as a result, the dissolution of blood clots (reviewed in [61]). Although fibrin is not reported to be in the seminiferous epithelium, the plasminogen system could nonetheless be important to this tissue because it is involved in many other normal and pathophysiological processes [62–66]. This section of the chapter discusses specific aspects of the biochemistry of the plasminogen system, the expression of individual components of this system by Sertoli cells and germ cells, and the reproductive phenotypes in male mice deficient in one or more components of this system. Although urokinase, tPA, and plasminogen are secreted proteinases, they bind to specific cell surface receptors, concentrating their activities in the immediate microenvironment around a cell. The receptors should be present in the seminiferous epithelium, because cell surface receptors for urokinase and tPA are transcriptionally expressed in this tissue (Section III.A.3). The final required component of the plasminogen system, inhibitors for urokinase, tPA, and plasmin, have also been detected in the testes (see Section III.A.4). Thus, the testis expresses all of the major components of the plasminogen system and this system has the capacity to concentrate proteinases on a cell’s surface and regulate their enzymatic activities. This fact and the highly regulated expression of urokinase and tPA (Section III.B.2) suggest that this system has a specific and important

tPA is secreted as a single-chain protein with a high affinity for plasminogen (Km = 65 μM) [60]. In contrast to the high substrate specificities of the plasminogen activators, plasmin has many substrates; one is tPA. The cleavage of tPA by plasmin generates a two-chain tPA, whose catalytic activity is approximately 30-fold greater than single-chain tPA [67]. Thus, tPA and plasminogen activate each other, a process similar to reciprocal proenzyme activation of prourokinase and plasminogen ([68]; see Section III.A.2). Recent evidence indicates that tPA not only binds fibrin but also cell surface Annexin A2 [69], which is represented in a Sertoli cell cDNA library (see Section VII for the library’s URL address). Thus, in the presence of plasminogen, the binding of tPA to Sertoli cells can potentially produce a high pericellular concentration of plasmin. Because many different proteins, including basement membrane components, are substrates of plasmin (Section III.A.5), this high plasmin concentration around the Sertoli cell could cause local degradation of the basement membrane of the seminiferous epithelium and release cytokines or growth factors trapped therein. 2. Urokinase and Plasminogen Urokinase is secreted as a proenzyme that has substantially reduced catalytic activity compared to the mature enzyme [70]. However, prourokinase is still able, albeit inefficiently, to convert plasminogen to plasmin. Like tPA, prourokinase is a substrate for plasmin. Initially produced by prourokinase, plasmin processes prourokinase to urokinase. This event, called reciprocal proenzyme activation, potentially provides for continuous production of enzymatically active urokinase [68]. The process is important because urokinase is rapidly and irreversibly inactivated by its inhibitor, PAI-1, which is expressed in the seminiferous epithelium [71]. Therefore, as long as prourokinase and plasminogen are present, reciprocal proenzyme activation will maintain enzymatically active urokinase and plasmin in a microenvironment within the seminiferous epithelium [68]. 3. Cell Surface Binding Sites for Urokinase and tPA Although it has been argued that the production of plasmin by soluble urokinase is physiologically

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important [68], plasminogen activation by urokinase occurs mostly on the cell surface. This process involves distinct cell surface binding sites for both urokinase and plasminogen. The urokinase plasminogen activator receptor (uPAR) is a high-affinity (Kd ~ 10–9 M) and abundant (~5 × 105/cell) binding site for urokinase [70, 72–74]. This receptor is expressed by spermatogenic cells and the binding of urokinase to mouse sperm demonstrates that the receptor is functional [75, 76] Many different cell types also express a relatively lowaffinity (Kd ~ 10–6 M) but very abundant (≥107/cell) cell surface plasminogen binding site [77]. Given the widespread cellular distribution of these sites, it is possible that they are also expressed in the seminiferous epithelium. The binding of both urokinase and plasminogen to a cell surface substantially increases the functional concentrations of urokinase and plasminogen and increases by approximately sixfold the efficiency of the enzymatic conversion of plasminogen to plasmin [70]. A final consequence of producing plasmin on the cell surface is the protection of plasmin from inactivation. It is estimated that the half-life of plasmin activity increases from 0.1 sec in solution to greater than 60 sec on the cell surface [70]. Recent data identify Annexin A2 as a cell surface binding site for tPA [69] and Annexin A2 mRNA is represented in a Sertoli cell cDNA library (see SectionVII for the library’s URL address). Thus, it is likely that uPAR and Annexin A2 facilitate the localized production of plasmin in the microenvironments surrounding germ cells and Sertoli cells, respectively. This production could have a marked effect on protein turnover and processing in these microenvironments and, as a consequence, modulate the interactions of Sertoli cells with each other, with spermatogenic cells, or with the basement membrane.

covalent inhibitor of urokinase and, to a lesser extent, of tPA ([80] and references therein). PCI is a third member of the serpin family, which also covalently inactivates its cognate enzymes [79]. However, in contrast to PAI-1 and -2, PCI inactivates multiple proteases including acrosin, tissue kallikrein, thrombin, prostate-specific antigen, urokinase, and activated protein C [81–83]. The two major inhibitors of plasmin activity are α2-antiplasmin and α2-macroglobulin. At low concentration (≤0.5 μM), α2-antiplasmin is only effective against soluble plasmin; at concentrations of 1 μM, it inhibits both soluble and cell surface plasmin. In contrast, current data argue that α2-macroglobulin, a rather nonspecific protease inhibitor, is only effective against soluble plasmin [77]. Thus, α2-antiplasmin is the primary inhibitor of plasmin bound to the cell surface. 5. Plasmin Substrates Plasmin has many physiologically important substrates besides fibrin. This enzyme processes prourokinase and single-chain tPA and degrades fibronectin, laminin, and proteoglycans within a basement membrane [70, 74, 84]. Via the production of plasmin on their cell surface, spermatogenic cells or Sertoli cells might also modulate their exposure to growth factors. For example, plasmin has been shown to liberate FGF2 from extracellular matrices and IGF from IGF-BP3 [85, 86]. Additionally, plasmin can release TGFβ1 from basement membranes and process pro-TGFβ1 to a mature, biologically active cytokine [87–90]. Thus, like the metzincin superfamily of proteinases, the plasminogen system may also remodel the seminiferous tubule basement membrane and release or activate growth factors within this tissue.

4. Inhibitors of the Plasminogen System Plasminogen activator and plasmin enzymatic activities in all tissue microenvironments are determined not only by the rates of enzyme synthesis and activation, but also by the rates of enzyme inactivation. This inactivation is the function of plasminogen activator inhibitors 1 and 2 (PAI-1 and PAI-2), protein C inhibitor (PCI), α2-antiplasmin, and α2-macroglobulin [58, 78, 79]. All except PAI-2 are expressed in the testis. PAI-1 is a member of the serpin family that reacts avidly with single-chain and two-chain tPA and with two-chain urokinase [58]. PAI-1 binds in an equal molar fashion with these plasminogens, and then irreversibly inhibits their activities by forming an ester bond with a serine residue in the enzyme’s active site [58, 78]. PAI-2 is another serpin, which is an effective

B. Expression of the Plasminogen System in the Seminiferous Epithelium 1. Urokinase, tPA, and PAI-1 Enzymatically active urokinase, tPA, and plasmin activities have been detected within rat seminiferous tubules [57, 59]. Urokinase has been identified as a product of Sertoli cells by Northern blot analysis, by identification of the secreted protein in Sertoli cell culture media, and by immunocytochemical analysis of testis sections (Fig. 9.5A). tPA is also expressed by seminiferous tubules and is localized to pachytene and diakinetic spermatocytes [57] (Fig. 9.5B). However, Sertoli cells from sexually immature rats secrete tPA in response to FSH [91]. PAI-1 is secreted by both

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rat peritubular myoid cells and Sertoli cells [71, 92, 93] (Fig. 9.6) and by monkey Sertoli cells [76]. 2. Regulated Expression of Urokinase, tPA, and PAI-1

A

B

FIGURE 9.5 Rat seminiferous tubules at stages VII to VIII of the cycle stained with anti-mouse urokinase (A) or anti-human tPA (B) (see color plate). Testis sections were fixed in 1% paraformaldehyde. Cryostat sections (6–8 μm) were cut and incubated with the antibody. Bound antibody was detected using the peroxidase–antiperoxidase method. In panel (A), arrows point to Sertoli cells. In panel (B), arrows point to pachytene spermatocytes. (Source: Figure 1 of Vihko, K. K., Kristensen, P., and Parvinen, M., Dev. Biol., Vol. 126, Immunohistochemical localization of urokinase-type plasminogen activator in Sertoli cells and tissue-type plasminogen activator in spermatogenic cells in the rat seminiferous epithelium, pp. 150–155. Copyright 1988 Elsevier.)

FIGURE 9.6 Expression of PAI-1 in rat testicular cells. RNA was obtained from Sertoli cells or peritubular myoid cells of 6-day-old rats and from crude or purified germ cells and residual bodies isolated from sexually mature rats. PAI-1 mRNA and GAPDH mRNA were sequentially assayed by Northern blot analysis. Molecular weight standards (kb) are shown on the left. (Source: Figure 2 of Le Magueresse-Battistoni, B., Pernod, G., Sigillo, F., Kolodie, L., and Benahmed, M., Biol. Reprod., Vol. 59, Plasminogen activator inhibitor1 is expressed in cultured rat Sertoli cells, pp. 591–598. Copyright 1998 Society for the Study of Reproduction.)

A remarkable aspect of the urokinase system in the seminiferous epithelium is that it is highly regulated by cell–cell interactions and by hormones. This regulation suggests that this system has a specific function in this tissue. Fritz and colleagues were the first to prove that a plasminogen activator was expressed by Sertoli cells and that its expression was precisely controlled by germ cells [94, 95]. Their pioneering work documented that this activity was primarily expressed by rat Sertoli cells at stages VII and VIII of the cycle of the seminiferous epithelium. Later studies by Parvinen and colleagues demonstrated that this stage-specific expression resulted from precise interactions between developing male germ cells and Sertoli cells [96]. This fact was established by examining stage-specific expression of plasminogen activator in tubules missing specific populations of spermatogenic cells. To obtain tubules without specific germ cells, the testes were irradiated and tubules analyzed at varying times thereafter. The rationale for this experimental approach is as follows. Irradiation causes death of proliferating spermatogonia. Consequently, with time, maturation depletion results in the absence of increasingly mature germ cells. However, the loss of the differentiated spermatogonia also activates the spermatogonial stem cells, which replicate and differentiate. These events give rise to a new cohort of differentiated spermatogonia, which then progress through spermatogenesis. Thus, at a given time following testicular irradiation, only one or two types of spermatogenic cells are missing. The effects of a given type of spermatogenic cell on stage-specific expression of plasminogen activator secretion can, therefore, be evaluated by examining stagespecific plasminogen activator secretion at varying times after irradiation. Using this experimental approach, Parvinen and colleagues demonstrated that preleptotene spermatocytes were required for stage-specific secretion of plasminogen activator by Sertoli cells in stage VII and VIII tubules (Fig. 9.7). Subsequent analyses identified urokinase as the predominant plasminogen activator secreted at these stages [97]. The complex cellular regulation in the seminiferous epithelium of the plasminogen system is also evident in the stage-specific expression of tPA mRNA [97]. In contrast to urokinase mRNA, Sertoli cells and possibly germ cells express low levels of tPA mRNA at stages VII and VIII. However, high levels of this transcript are detected in Sertoli cells at stages IX through XIII.

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Urokinase and tPA expression by Sertoli cells also differ in their responses to FSH. Treatment of cultured rat Sertoli cells with FSH suppresses expression of urokinase but stimulates expression of tPA [98]. Measurement of the total urokinase activity secreted by cultured tubules indicates, however, that the net effect of FSH is to increase the total amount of secreted plasminogen activator enzyme activity (Fig. 9.8A) [36]. Interestingly, FSH also suppresses expression of PAI1, potentially contributing to the stimulatory effect of FSH on total plasminogen activator enzyme activity (Fig. 9.8B).

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FIGURE 9.7 The effect of germ cell depletion and restoration on stage-specific expression of total plasminogen activator secretion in seminiferous tubules at different stages of the cycle. Rat testes were exposed to 3 Gy of radiation and, at varying times thereafter, seminiferous tubules at specific stages of the cycle were isolated and cultured for 24 hr. Total plasminogen activator secretion was assayed by incubating culture medium with 125I-fibrinogen and plasminogen. Proteolysis of 125I-fibrinogen was expressed as a percentage of total 125I-fibrinogen in the assay. (A) Plasminogen activator secreted by control tubules at stages VII to IX of the cycle of the seminiferous epithelium. (B) Plasminogen activator secreted by seminiferous tubules, which lack preleptotene spermatocytes (8 days postirradiation). (C) Plasminogen activator secreted by seminiferous tubules in which the preleptotene spermatocytes are partially restored but pachytene spermatocytes are reduced in number (21 days postirradiation). Note the partial restoration of stage-specific PA secretion. (Source: Figures 1B, 3B, and 4B of Vihko, K. K., Suominen, J. J., and Parvinen, M., Biology of Reproduction, Vol. 3, Cellular regulation of plasminogen activator secretion during spermatogenesis, pp. 383–389. Copyright 1984 Society for the Study of Reproduction.)

Although at these stages tPA localizes to spermatocytes, the total tPA mRNA levels in tubules are not reduced when these cells are lost due to irradiationinduced maturation depletion [97]. This observation suggested that the immunocytochemical localization of tPA protein to spermatogenic cells (Fig. 9.5B) was due to binding of tPA to these cells.

The precise biochemical and physiological regulation of plasminogen activator and plasmin activities and the presence of receptors for these proteinases in the testis argue that the plasminogen activators and/or plasmin have an important function in the seminiferous epithelium and, thus, in spermatogenesis. This predicts that inactivation of genes in this system would be highly detrimental to male reproductive function. Thus, the fertility of male mice that lack urokinase, tPA, or the urokinase receptor, uPAR1, was not anticipated [99, 100]. In contrast, after birth, mice deficient in both tPA and urokinase grew slower and died sooner than mice that lacked either urokinase or tPA [99]. Finally, fewer pups were born when doubleknockout male and female mice were bred. However, whether this effect on fertility was due to deficits in the reproductive function of males, females, or both sexes was not systematically investigated. The effect of inactivating the plasminogen gene is similar to the effect of inactivating both the urokinase and tPA genes in the same animal [101]. Plasminogendeficient mice grow more slowly and exhibit reduced fertility. However, detailed breeding studies indicate that an inability to form plasmin affects the fertility of the female but not male mice. Thus, neither urokinase, tPA, nor plasminogen is essential for male fertility. Note, however, that the effects of these single or double gene knockouts on testis weight, sperm production, or morphology of the testis were not reported [99, 101]. Therefore, whether or not urokinase, tPA, or plasminogen has quantitative effects on spermatogenesis remains to be addressed. Lack of PAI-1 or α2-macroglobulin also does not cause male infertility [102, 103]. However, PCI-deficient mice are sterile [104]. Light and electron microscopic studies of the testes and epididymides of PCI-deficient mice revealed sloughing of germ cells, substantially reduced sperm production, and vacuolization and

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− FIGURE 9.8 FSH stimulates plasminogen secretion and inhibits PAI-1 mRNA expression in Sertoli cells. (A) Effect of FSH on total plasminogen activator secretion in segments of rat seminiferous tubules at stages VII to VIII. Plasminogen activator in the culture medium was assayed as described in Fig. 9.7. (Source: Figure 1 of Vihko, K. K., Penttila, T. L., Parvinen, M., and Belin, D., Molecular Endocrinology, Vol. 3, Regulation of urokinase- and tissue-type plasminogen activator gene expression in the rat seminiferous epithelium, pp. 52–59. Copyright 1989 The Endocrine Society.) (B) Effect of FSH on PAI-1 mRNA in cultured Sertoli cells. Sertoli cells isolated from 20-day-old rats were cultured without FSH (control), with 50 ng/mL FSH (+FSH 50), or with 100 ng/mL FSH (+FSH 100). PAI-1 mRNA levels were assayed by Northern blot analysis and normalized to GAPDH mRNA levels in the same samples. (Source: Figure 6 of Le Magueresse-Battistoni, B., Pernod, G., Sigillo, F., Kolodie, L., and Benahmen, M., Biology of Reproduction, Vol. 59, Plasminogen activator inhibitor-1 is expressed in cultured rat Sertoli cells, pp. 591–598. Copyright 1984 Society for the Study of Reproduction.)

structural disorganization of the seminiferous epithelium (Fig. 9.9). Thus, the infertility of PCI-deficient male mice demonstrates that male fertility requires the appropriate balance between specific proteinases and proteinase inhibitors in the plasminogen system.

IV. THE CATHEPSINS A. Background The cathepsins are distributed among all four classes of proteases: aspartyl (cathepsins D and E), cysteine (the majority of the cathepsins), metallo (cathepsin III), and serine (cathepsins A and G) proteases (reviewed in [105]). They are either endopeptidases or exopeptidases, with some having overlapping activities (e.g., cathepsins B and H) [106]. As discussed in Section IV.H, the enzymatic activities of the cathepsins are inhibited by α2-macroglobulin and members of the cystatin superfamily. The first pure cathepsin was isolated in 1948 by Gutman and Fruton [107], but the identification of new

cathepsins progressed slowly during the next 20 years: By the early 1970s, only three additional family members (cathepsins B, H, and L) had been reported in the literature [106]. Today, a survey of the different sequenced genomes reveals that up to 21 cathepsins are present in humans, up to 20 in mice, and up to 13 in rats. Of the 21 human cathepsin genes, 16 have orthologs in mice and 11 in rats. Although several cathepsin genes map to different chromosomes, 4 are clustered on human chromosome 11, 9 on mouse chromosome 13, and 4 on rat chromosome 17.

1. Biosynthesis Cathepsins are synthesized as inactive precursors that contain an amino-terminal signal peptide, a propeptide located toward the amino terminus, and a mature active domain (reviewed in [108–110]). The signal peptide mediates the transport of these proteinases across the endoplasmic reticulum (ER). As the proteinases pass into the lumen of the ER, the signal peptide is removed, giving rise to an inactive proenzyme.

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endopeptidases such as cathepsins L and S for their activation [111, 112]. Two lines of evidence demonstrate the importance of the propeptide in the folding and the enzymatic activity of the mature proteinases. First, the propeptide can inhibit the activity of the mature enzyme in vitro, suggesting that the propeptide blocks the active site of the enzyme until it is activated [113, 114]. Second, cathepsins expressed without their propeptide are not folded properly and remain inactive [115]. In the lysosomes, the single-chain enzymes can be processed into two-chain forms, comprised of one heavy chain and one light chain (reviewed in [108]). The subunits of the two chain forms are held together via noncovalent interactions or via disulfide bridges. Whether the single-chain enzymes undergo asymmetric cleavage into a light and a heavy chain appears to be cell type specific. Finally, very late during their processing, most proteinases undergo amino acid trimming at the cleavage site between the light and heavy chains and/or at their carboxyl termini (reviewed in [108]). 2. Biological Functions

FIGURE 9.9 Morphological comparison of the seminiferous epithelium of heterozygous PCI-deficient mice (A and C) with the seminiferous epithelium of homozygous PCI-deficient mice (B and D). Panels (A) and (B) are 0.5-μm epon sections stained with toluidine blue. Panels (C) and (D) are osmicated 80-nm epon sections stained with uranyl acetate. The arrows in panels (B), (C), and (D) identify Sertoli cells. (Source: Figures 4A–D from Uhrin, P. et al., Journal of Clinical Investigation, Vol. 106, Disruption of the protein C inhibitor gene results in impaired spermatogenesis and male infertility, pp. 1531–1539. Copyright 2000 Journal of Clinical Investigation.)

Carbohydrate chains containing mannose-6-phosphate residues are added to the proenzymes (the number of chains added varies with the particular proteinase). In most cases, the phosphorylated, asparagine-linked proenzymes are then recognized by mannose-6phosphate receptors located in the Golgi apparatus. These receptors ensure the association of the proteinases with vesicles that are targeted to the lysosomes. Following dissociation from the receptors in the prelysosomal compartments, the proenzymes are activated by the local acidic environment. The lysosomal membrane contains an ATP-dependent H+ pump that introduces protons into the lysosomal lumen, producing a pH of 4.5 to 5.5. The autocatalytic activation at acidic pH leads to the removal of the propeptide and the generation of active single-chain enzymes (reviewed in [108]). In vitro, only endopeptidases can be autoactivated, whereas exopeptidases require

Cathepsins are ubiquitously expressed and participate in the proteolytic degradation of proteins that have been imported by endocytic traffic to the lysosomes (reviewed in [116]). In addition to this housekeeping function, some cathepsins have been implicated in the proteolytic processing of specific and important bioactive molecules. Examples include the production of endostatin from collagen XVIII mediated by cathepsin L [117]; the processing of thyroglobulin into thyroid hormones T3 and T4 by the coordinate action of cathepsins B, K, and L [118]; the formation of angiostatin from plasminogen mediated by cathepsin D in prostate cancer cells [119]; the presentation of antigenic peptides on major histocompatibility complex (MHC) class II molecules mediated by cathepsins L and S [120–123]; the conversion of prorenin to renin by cathepsin B [124]; the production of the enkephalin peptide neurotransmitter by cathepsin L [125]; and the degradation of the epidermal growth factor (EGF) and its receptor by cathepsin B [126]. In the following sections, data regarding the cathepsins expressed in Sertoli cells (Sections IV.B through G) and their natural inhibitors (Section IV.H) are discussed. A more thorough examination of cathepsin L is provided (Section IV.B) because this protease is the most extensively studied member of the cathepsin family expressed in Sertoli cells. Readers interested in topics regarding cathepsins not covered in this chapter are referred to comprehensive reviews by McGrath [105] and Turk et al. [127].

Chapter 9 Proteases and Protease Inhibitors

B. Cathepsin L 1. Processing and Activation The human cathepsin L gene encodes a 333-aminoacid cysteine proteinase that contains a 17-amino-acid signal peptide, a 96-amino-acid propeptide, and a 220-amino-acid mature region. In the lysosomes, the 38-kDa procathepsin L is processed to mature, active cathepsin L and exists as a single-chain form of about 28 kDa and/or as a two-chain form of about 24 and 4 kDa (heavy and light chain, respectively) [128–130]. 2. Properties Cathepsin L plays a major role in the degradation of endocytosed proteins as well as intracellular proteins (reviewed in [105, 131]). Although procathepsin L is normally routed to lysosomes, it can also be secreted and, thus, can potentially degrade proteins in an acidic extracellular environment. Procathepsin L was originally named MEP (major excreted protein) because 30% of the proteins secreted by transformed mouse fibroblasts turned out to be procathepsin L [132]. An acidic pH is required to activate this proteinase. In vitro assays using synthetic substrates have established that cathepsin L is optimally active at pH 5.5 but inactive at pH 7.0 [133]. In vitro, it can degrade a variety of substrates including actin, albumin, collagen, elastin, fibronectin, and laminin (reviewed in [134]). 3. Expression in Sertoli Cells Procathepsin L was identified by two-dimensional gel electrophoresis of proteins secreted by intact seminiferous tubules at defined stages of the cycle of the seminiferous epithelium [135, 136]. Procathepsin L was the predominant secretory product in tubules at stages VI and VII, the protein being undetectable at most other stages. The stage-specific secretion was confirmed by radiolabeling and immunoprecipitation of the protein secreted by seminiferous tubules isolated from sexually mature rats [137]. The stage-specific secretion of procathepsin L was shown to reflect its rate of synthesis. The proenzyme represented 0.02% of the total protein synthesized in 1 hr by stage VI to VII tubules, its synthesis increasing more than 20-fold as germ cells that surround Sertoli cells progress from stage XII to VI [135]. Immunocytochemical studies corroborated these findings and established that in the seminiferous epithelium, detectable amounts of procathepsin L were only found in the somatic Sertoli cells, in regions concentrated around most elongate spermatids [138].

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In addition, cathepsin L was also detected in rat germ cells undergoing apoptosis [139]. As observed for the synthesis and secretion of the proenzyme, levels of cathepsin L mRNA in rodent Sertoli cells are highly regulated with respect to the stages of the cycle. In sexually mature rats, cathepsin L transcripts are expressed at high levels in Sertoli cells in tubules at stages VI to VII, and at low or undetectable levels at all other stages [136, 140]. As anticipated from the previous results on the cellular localization of procathepsin L in the testis, cathepsin L transcripts were not detected in germ cells or peritubular cells by Northern blot and in situ hybridization analyses. 4. Cell–Cell Interactions Regulate Stage-Specific Expression of the Cathepsin L Gene in Rat Sertoli Cells The stage-specific expression of cathepsin L mRNA and protein synthesis/secretion implied precise interactions between germ cells and Sertoli cells. Studies were conducted to gain insight into how germ cells regulate the stage-specific transcription of the rat cathepsin L gene. In one study, the effects of maturation depletion and repletion of germ cells on the stage-specific transcription of the rat cathepsin L gene in Sertoli cells was examined by exposing adult rat testes to gamma radiation in order to kill proliferating spermatogonia [141]. The effect of this treatment on the cellular composition of the seminiferous epithelium has already been discussed in Section III.B.2. The loss of specific germ cells from the seminiferous epithelium was correlated with changes in stage-specific expression of cathepsin L transcripts by Sertoli cells. In contrast to what was observed for the plasminogen activators (see Section III.B.2), this study showed that lack of preleptotene spermatocytes did not affect stage-specific transcription of the cathepsin L gene [141]. However, when pachytene spermatocytes through step 14 spermatids were depleted, the transcription of the rat cathepsin L gene was reduced by half at stages VI to VIII and increased 14-fold at stages IX to I. As a result, all Sertoli cells expressed similar levels of the transcript. Finally, stage VI to VIII tubules, depleted primarily in step 15 through 19 spermatids, had levels of cathepsin L mRNA that were 60% of that of control tubules at stages VI to VII [141]. However, stage-specific transcription of the cathepsin L gene was detected in these tubules. Collectively, these results suggested that pachytene and diplotene spermatocytes are responsible for suppressing the transcription of the cathepsin L gene at stages I to V and IX to XIV, and that elongate spermatids stimulated expression of

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5. Identification of a Promoter That Conveys Stage-Specific Expression in Sertoli Cells The molecular mechanisms responsible for stagespecific gene expression in Sertoli cells are unknown. A requirement for defining how male germ cells modulate stage-specific expression in these somatic cells is the identification of a promoter that confers accurate, stage-specific gene expression in vivo. Because the complex interactions between Sertoli cells and germ cells cannot be fully replicated in vitro, the characterization of such a promoter requires a transgenic mouse model. A β-galactosidase transgene that contained the 3-kb genomic region located immediately upstream from the translation start site of the rat cathepsin L gene was used to generate transgenic mice [144]. In two independent mouse lines, the only organ in which β-galactosidase

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cathepsin L transcripts in stage VI to VIII tubules. Consistent with this conclusion, results from another study indicated that elongate spermatids stimulated cathepsin L mRNA expression [142]. Analysis of the stage-specific expression of cathepsin L mRNA during maturation of the rat testis extended and confirmed the results described above. During the end of puberty in the rat (from 30 to 55 days of age), the amount of cathepsin L mRNA per Sertoli cell increases sixfold [139]. Simultaneously, germ cells at specific stages of the cycle of the seminiferous epithelium begin to stimulate or to repress expression of cathepsin L mRNA in Sertoli cells. At 30 days of age, in situ hybridization revealed low levels of this transcript in all rat Sertoli cells. However, by 40 days of age (when the most mature germ cell in a stage VII tubule is a step 7 spermatid), this transcript was only detected in Sertoli cells within stage VI to VII tubules. Upon completion of the first round of spermatogenesis (about 45 days of age), expression of this transcript markedly increased in Sertoli cells within stage VI and VII tubules, corresponding to the developmental period when elongate spermatids are first observed in the seminiferous epithelium [136, 139, 142, 143]. Based on these results, it appears that stage-specific transcription of the rat cathepsin L gene is repressed by signals from pachytene or diplotene spermatocytes at stages I to V and IX to XIV. However, at stages VI to VIII, the repressive signal ceases or is overridden. In addition, at these stages, signals from the elongate spermatids stimulate the expression of the cathepsin L gene. These results lend themselves to a model in which stage-specific expression of the cathepsin L gene results from sequential repression and stimulation of transcription (Fig. 9.10). If such a model is accurate, one would predict that the cathepsin L promoter contains cis-acting elements that respond to such signals.

Levels of rat cathepsin L transcripts (%)

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FIGURE 9.10 Stage-specific transcription of the cathepsin L gene in Sertoli cells of sexually mature rats. Based on data presented in Section IV.B.4, it is proposed that the stage-specific expression of the rat cathepsin L gene results from sequential waves of transcriptional activation and repression. The stage-specific transcription of this gene is repressed by signals from pachytene or diplotene spermatocytes at stages I to V and IX to XIV. However, at stages VI to VIII, the repressive signal ceases or is overridden. In addition, at these stages, signals from the elongate spermatids stimulate the expression of the cathepsin L gene.

enzymatic activity was detected was the testis. Reporter gene enzymatic activity in testes from transgenic mice was at least 250-fold greater than that measured in testes of control littermates. The fact that expression of the transgene was only detected in testes was surprising because high levels of endogenous cathepsin L transcripts are detected in kidney and liver [136]. The lack of expression in these tissues indicates that the regulatory elements that drive high levels of cathespin L mRNA in kidney and liver were missing from the 3-kb promoter region used in this study. In situ analysis revealed that all of the reporter gene enzymatic activity was in the seminiferous tubules and varied along the length of a tubule, consistent with stage-specific expression of the transgene [144]. Light microscopy analysis of testis sections from transgenic mice revealed that expression of the transgene was restricted to Sertoli cells (Fig. 9.11). To determine whether the transgene was expressed in a stage-specific manner, the stages of the cycle of the seminiferous epithelium of tubules that contained Sertoli cells expressing β-galactosidase were identified. Seventyfive percent of the Sertoli cells that expressed the transgene were at stages VI to VIII. The other Sertoli cells expressing the transgene were predominantly in stage V and IX tubules. As observed for the endogenous rat cathepsin L gene [139], expression of the transgene was also maturation dependent as

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FIGURE 9.11 The 3-kb genomic fragment located immediately upstream of the rat cathepsin L translation start site directs expression of the reporter gene, β-galactosidase, only in Sertoli cells of transgenic mice (see color plate). The blue staining is indicative of the expression of the LacZ reporter gene. The white arrowheads identify two Sertoli cells in a stage VII tubule. The black arrows labeled 1, 2, and 3 point to a pachytene spermatocyte, a round spermatid, and an elongate spermatid, respectively. The horizontal black bar is equal to 10 μm. (Source: Figure 4 of Charron, M., Folmer, J. S., and Wright, W. W., Biology of Reproduction, Vol. 68, A 3-kilobase region derived from the rat cathepsin K gene directs in vivo expression of a reporter gene in Sertoli cells in a manner comparable to that of the endogenous gene, pp. 1641–1648. Copyright 2003 Society for the Study of Reproduction.)

β-galactosidase activity increased 4.5-fold during the end of testis maturation [144]. Thus, the transgene was expressed at the same stages of the cycle and in the same maturation-dependent manner as the endogenous gene. The next step in the characterization of the cathepsin L promoter is the identification of the regulatory elements that are required for the stage-specific expression of this gene in Sertoli cells. The results from transient transfection assays and in vitro DNA–protein interactions support a working model in which the 3-kb cathepsin L promoter can be divided into at least three functional domains. The first domain is a ~120– base pair (bp) region that flanks the transcription start site of the rat cathepsin L gene. Transfection analyses using primary rat Sertoli cell cultures have established that this ~120-bp region derived from the rat cathepsin L gene is critical for promoter activity in Sertoli cells isolated from sexually mature rats [145]. Within this 120-bp region, a single 17-bp GC-box, which binds the transcription factor Sp3 (or an Sp3-related protein), was shown to be a critical regulatory element for promoter activity in mature Sertoli cells [145]. The second domain corresponds to the first intron of the cathepsin L gene.

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Transfection analyses in primary rat Sertoli cell cultures indicate that the first intron of the rat cathepsin L gene contains regulatory elements that activate transcription in rat Sertoli cells. Finally, the third domain appears to be responsible for the repressive effects mediated by male germ cells [35]. Transfection analysis demonstrated that two constructs that contain either 2065 or 244 bp upstream of the transcription start site of the rat cathepsin L gene have similar activities in Sertoli cells isolated from sexually mature rats. However, addition of pooled male germ cells to the transfected Sertoli cells only affected the activity of the longer construct, which was reduced by 30%. This result, coupled with the observation that transcription of the cathepsin L gene in mature Sertoli cells is repressed at most stages of the cycle of the seminiferous epithelium, is consistent with the hypothesis that male germ cells modulate transcription of the cathepsin L gene in Sertoli cells. Future experiments using a combination of both in vitro and in vivo experimental approaches should allow the identification of the specific regulatory elements and the transcription factors that confer stage-specific expression of the cathepsin L gene in Sertoli cells. 6. Cathepsin L–Related Genes In humans a gene closely related to the cathepsin L gene has been identified and designated cathepsin V [146, 147]. The cathepsin V gene encodes a 334-aminoacid cysteine proteinase that contains a putative 17-amino-acid signal peptide, a 96-amino-acid propeptide, and a 221-amino-acid mature region. The amino acid sequences between cathepsins L and V are quite similar, with a sequence identity of 78% for the proenzyme and 80% for the mature enzyme [147]. The high sequence identity and the fact that the cathepsin V gene is located adjacent to the cathepsin L gene on human chromosome 9 [147, 148] suggests that cathepsins L and V may have evolved from a common ancestral precursor by gene duplication. Despite these similarities, cathepsins L and V differ in their substrate specificity [147]. Also, expression of cathepsin V appears to be more restricted than that of cathepsin L. Northern blot analysis using RNA isolated from 16 human tissues revealed that cathepsin V transcripts are detected predominantly in the testis and the thymus [146]. Researchers have not yet established whether cathepsin V transcripts are expressed in Sertoli cells. It appears that the human cathepsin V gene has no rodent ortholog and the biological function of this protein is currently unknown. Three additional cathepsin L–related genes are also present on human chromosome 10 [149], but no data regarding

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the activity or function of the proteins they encode are currently available.

tissues examined. However, these mice were used to demonstrate that cathepsin B plays an important role in trypsinogen activation in acute pancreatitis [160].

7. Reproductive Phenotype Observed in Knockout Mice While precise, stage-specific expression of cathepsin L in Sertoli cells suggests an important function for this proteinase in the seminiferous epithelium, cathepsin L–deficient mice are fertile (reviewed in [150]). Absence of cathepsin L activity in mice does prevent proper MHC class II antigen presentation in the cortical epithelium of the thymus by impairing the normal degradation of the invariant chain (li) [120]. Absence of this proteinase also interferes with hair cycle control and leads to deregulation of epidermal homeostasis [120, 151].

C. Cathepsin B 1. Processing and Activation Together with cathepsin L, cathepsin B is the most abundant cysteine proteinase, with the lysosomal concentration for both proteinases reaching 1 mM [127]. The human cathepsin B gene encodes a 339-amino-acid cysteine proteinase that contains a 17-amino-acid signal peptide, a 62-amino-acid N-propeptide, a 254-aminoacid mature region, and a 6-amino-acid C-propetide. In lysosomes, the 42-kDa procathepsin B is stored as mature, active cathepsin B and exists as a single-chain form of about 30 kDa and/or a two-chain form of about 25 and 5 kDa (heavy and light chain, respectively). Procathepsin B is processed to the mature active form by autoactivation [152], as well as by cathepsin D [153], serine peptidases [154], and metallopeptidases [155]. This proteinase is optimally active at pH values between pH 5.5 and 6.5 [156]. The mature enzyme can activate, degrade, or inactivate a wide variety of proteins and enzymes (reviewed in [134]). 2. Expression in Sertoli Cells In one report, using reverse transcriptase–polymerase chain reaction (RT–PCR), cathepsin B transcripts were detected in Sertoli cells isolated from 20-day-old rats [157]. However, in a second study, using two different rat procathepsin B antibodies, this cathepsin was not detected in either Sertoli cells or germ cells [158]. Because of the disparity between these data, it is unclear if enzymatically active cathepsin B is expressed in Sertoli cells. 3. Reproductive Phenotype Observed in Knockout Mice Cathepsin B–deficient mice are fertile and indistinguishable from their wild-type counterparts [150, 159, 160]. No abnormalities were detected in the

D. Cathepsin K 1. Processing and Activation Contrary to the ubiquitous expression of cathepsins B and L, cathepsin K shows a restricted tissue distribution, being found predominantly in ovaries and osteoclasts [161–163]. The human cathepsin K gene encodes a 329-amino-acid cysteine proteinase that contains a 15-amino-acid signal peptide, a 99-amino-acid propeptide, and a 215-amino-acid mature region [164]. In the lysosomes, the inactive 38-kDa procathepsin K is converted to a mature, active form of about 27 kDa [161]. Procathepsin K is processed into the mature active form by autoactivation [165] or by treatment with pepsin at pH 4.0 [162]. Optimally active at pH values between 6.0 and 6.5 [162], cathepsin K can degrade, in vitro, type I collagen, fibrinogen, and elastin (reviewed in [134]). 2. Expression in Sertoli Cells Cathepsin K is also detected in Sertoli cells [166]. By immunocytochemistry, this proteinase was shown to be localized in small punctate lysosomes and in early and late residual bodies. Northern blot analysis established that cathepsin K is expressed in a stage-specific manner similar to that of cathepsin L, with maximal expression in tubules at stages V to VIII [166]. 3. Reproductive Phenotype Observed in Knockout Mice Cathepsin K–deficient mice appear normal and are fertile. No overt phenotypic abnormalities were detected during the course of the experiments (up to 10 months of age). However, these mice developed osteopetrosis due to impaired resorption of bone matrix [167].

E. Cathepsin S 1. Processing and Activation The human cathepsin S gene encodes a 331-aminoacid protein that contains a 16-amino-acid signal peptide, a 98-amino-acid propeptide, and a 217-amino-acid mature region. In the lysosomes, the 37-kDa procathepsin S is processed to a 24-kDa mature proteinase. Whereas cathepsins B and L are composed of two chains, the active cathepsin S is a single-chain enzyme. In vitro, procathepsin S is processed into the mature active

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form either by autoactivation or by treatment with subtilisin [168]. Cathepsin S is the second known cysteine proteinase to be active in the neutral pH range (the other one being cathepsin K). This proteinase is catalytically active in a broad pH range (from pH 4.5 to 7.5), but optimally active at pH 6.5 [134]. In vitro, cathepsin S can degrade a wide variety of proteins such as albumin, collagen, elastin, fibronectin, laminin, and proteoglycans (reviewed in [134]). 2. Expression in Sertoli Cells Using in situ hybridization, cathepsin S transcripts were detected near the basal lamina of seminiferous tubules, consistent with their localization in Sertoli cells and possibly primary spermatocytes [169]. Cathepsin S transcripts were detected in tubules at all stages of the cycle of the seminiferous epithelium but peaked in stage VII to VIII tubules. 3. Reproductive Phenotype Observed in Knockout Mice Cathepsin S–deficient mice are fertile and appear normal during the course of the experiments (up to 6 months) [121]. However, mice lacking cathepsin S failed to fully process the MHC class II associated invariant chain (li) in dendritic cells [121]. In contrast, cathepsin L is involved in li processing in the cortical epithelium of the thymus (Section IV.B.7).

F. Cathepsin A 1. Processing and Activation Cathepsin A is a member of the serine carboxypeptidase family. The human cathepsin A gene encodes a 480amino-acid protein that contains a 28-amino-acid signal peptide and a 452-amino-acid precursor [170]. Contrary to the cysteine proteinases, cathepsin A does not contain a propeptide near the amino terminus. Cathepsin A is synthesized as a 54-kDa inactive precursor. Shortly after its synthesis, the precursor dimerizes and is transported to the lysosomes [171], where it undergoes endoproteolytic removal of a 2-kDa peptide [172]. The excision peptide is located near the middle of the polypeptide (amino acid 285 to 298). The 52-kDa inactive precursor is then processed to the mature two-chain form of 32 and 20 kDa (heavy and light chain, respectively) [170, 173]. In the lysosomes, cathepsin A associates with and forms a functional high molecular weight complex with β-D-galactosidase and N-acetyl-α-neuraminidase [174–177]. In this enzyme complex, cathepsin A protects β-galactosidase against rapid proteolytic degradation [176] and facilitates the intracellular routing,

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lysosomal localization, and activation of N-acetyl-αneuraminidase [178, 179]. As a member of the serine proteinase family, cathepsin A exerts its own catalytic activity at an acidic pH but maintains its deamidase/ esterase activity at neutral pH [170, 180]. These two functions are fully separable from its protective function. The activity of cathepsin A is essential for the complete degradation of selected bioactive peptides within the lysosomes, including substance P, oxytocin, and endothelin I (reviewed in [181]). Cathepsin A deficiency in humans causes a lysosomal storage disease called galactosialidosis [174], which is characterized by an increase in the number and volume of lysosomes and reduced activity of both N-acetyl-α-neuraminidase and β-D-galactosidase. 2. Expression in Sertoli Cells In situ hybridization and immunocytochemistry revealed that cathepsin A is expressed in Sertoli cells [182, 183]. Expression of this proteinase was noted in Sertoli cells within tubules at all stages of the cycle of the seminiferous epithelium. It was highly expressed in the basal region of Sertoli cells in tubules at stages I to VIII and XIII to XIV, and weakly expressed in tubules at stages IX to XII [184]. In tubules at stages IX to XII, cathepsin A was detected in residual bodies near the base of the seminiferous epithelium, but not in residual bodies located closer to the lumen of the seminiferous epithelium. This pattern of expression was consistent with the loss of lysosomes at stages IX to XII, stages during which lysosomes fuse with the residual bodies within the Sertoli cells [13]. This fusion occurs in the middle and basal portion of Sertoli cells. In addition, cathepsin A was detected in lysosomes of Leydig cells, in interstitial macrophages, and in the vascular endothelium [184]. However, germ cells do not express this proteinase. Collectively, these results indicate that in Sertoli cells, cathepsin A is one of the proteases responsible for degradation of the residual bodies and the germ cell-specific antigens that they contain. 3. Reproductive Phenotype Observed in Knockout Mice Cathepsin A-deficient mice showed clinical symptoms similar to human patients with galactosialidosis [185]. The presence of large lysosomes was observed in kidney, intestine, liver, pancreas, brain, and pituitary gland (possibly a consequence of the abnormal accumulation of undigested metabolites). Analysis of cathepsin A-deficient mice also revealed that their reproductive capacity is affected [182]. Both male and female mice are fertile but they mate poorly. The frequency of pregnancies is less than that for wild-type mice and

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worsens with age. Litter sizes and delivery appear normal. In female mice that lack cathepsin A, macrophages infiltrated the stroma of the ovary and uterus [182]. In cathepsin A–deficient male mice, extensive vacuolation of the epididymis and Leydig cells was observed and large spherical macrophages were present in the interstitial space of the testis and large lysosomes accumulated in Leydig cells [182, 186]. However, it is important to point out that in the seminiferous tubules of the cathepsin A–deficient mice, both Sertoli and germ cells appeared normal [186].

G. Cathepsin D 1. Processing and Activation The human cathepsin D gene encodes a 412-aminoacid aspartyl endopeptidase that contains a 20-aminoacid signal peptide, a 44-amino-acid propeptide, and a 348-amino-acid mature region [187]. In the lysosomes, the 53-kDa procathepsin D is processed into an active proteinase of about 51 kDa [188]. Whereas both singleand double-chain forms are detected in human cells [109], the single-chain form of cathepsin D is predominantly present in rat hepatocytes [189] and mouse fibroblasts [190]. Found predominantly in intracellular vesicles, lysosomes, phagosomes, and late endosomes, cathepsin D is optimally active at pH 3.8 [191]. A unique feature of this proteinase is its regulation by estrogens and by growth factors in breast cancer cell lines [192, 193]. 2. Expression in Sertoli Cells In the rat testis, cathepsin D was detected in Sertoli cells, in Leydig cells, and over the acrosomes of spermatids [158]. Immunocytochemistry coupled with electron microscopy showed that cathepsin D was present in the lysosomes of the Sertoli cells. Surprisingly, cathepsin D was not detected in the residual bodies, excluding this proteinase as an important player in the degradation of these structures. Cathepsin D transcripts were detected in the basal compartment of rat seminiferous tubules by in situ hybridization [169], suggesting that this gene is not only expressed in Sertoli cells but also in spermatocytes. The reason for the discrepancy between these two reports regarding the expression of cathepsin D in germ cells is unclear. 3. Reproductive Phenotype Observed in Knockout Mice Cathepsin D–deficient mice develop normally during the first 2 weeks, stop thriving in the third week, and die 26 days after birth in a state of anoxeria [194].

Inactivation of the mouse cathepsin D gene caused atrophy of the intestinal mucosa as well as destruction of lymphoid cells [194]. However, the authors established that lack of cathepsin D did not impair bulk lysosomal protein turnover.

H. Cathepsin Inhibitors 1. The Cystatin Superfamily The cystatin superfamily is an evolutionarily related group of proteins that are divided into three families: family 1, the stefins; family 2, the cystatins; and family 3, the kininogens (reviewed in [195]). Members of the stefin family are generally unglycosylated proteins that lack disulfide bonds. The stefins have been shown to be potent, reversible, and competitive inhibitors of cysteine proteinases. Members of the kininogen family are single-chain glycoproteins characterized by three regions each containing two disulfide loops and a bradykinin sequence at the carboxyl terminus. There are three distinct types of kininogens: the high molecular weight kininogens (H-kininogens), the low molecular weight kininogens (L-kininogens), and the T-kininogen (also called major acute phase protein). The activity of cathepsin L is strongly inhibited by all three kininogens, whereas the activities of cathepsin B and H are weakly inhibited by H- and L-kininogens [195]. Members of the family 2 cystatins are secreted proteins that consist of approximately 120 amino acid residues. They share several features: a signal peptide, four characteristic cysteine residues located in the carboxyl-terminal domain, and three short conserved amino acid domains that coincide with those forming the inhibitory site of the protein (reviewed in [195]). Although evolutionarily related, members of the family 2 cystatins are only 30% to 50% homologous at the amino acid levels [195]. They can be divided into two groups. The first group includes cystatins C, D, E/M, F, S, SN, and SA, and the second group includes CRES, CRES2, cystatins SC, T, TE-1, and TE-2, and testatin (reviewed in [196]). Members of the second group share two common characteristics that distinguish them from members of the first group. First, lacking both the cystatin N-terminal region and the conserved Q-V-G domain, they possess only one of the three conserved domains in cystatins required to block the active site of cysteine proteinases. Second, they are all primarily expressed in reproductive tissues (reviewed in [196]). The functions of members of the family 2 cystatins expressed in Sertoli cells are currently unknown. Because they lack the crucial domains required to perform inhibitory

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functions, their biochemical activities may differ from that of the family 1 cystatins. For example, in vitro, CRES failed to inhibit cysteine proteinases but was shown to be a potent inhibitor of the serine proteinase prohormone convertase 2 [197]. a. Cystatin 2 Family Members Expressed in Sertoli Cells Immunohistochemistry revealed that cystatin C is localized in Sertoli cells and around the heads of compacted spermatids [198]. The latter could reflect the presence of cystatin C in the spermatids or in the surrounding cytoplasm of the Sertoli cells. Northern blot analysis revealed that cystatin C transcripts were present in Sertoli cells, in pachytene spermatocytes, and in spermatids [198], consistent with the possibility that the staining around the acrosome represented cystatin C in the spermatids. Analysis of total RNA from stage-synchronized testes demonstrated that transcription of the rat cystatin C gene in Sertoli cells is stage specific, with the highest expression in tubules at stages VIII to I and lowest in tubules at stages VI to VII [198]. This stage-specific expression pattern is opposite to the one described for the cathepsin L gene (Section IV.B.3), suggesting that cystatin C could limit the extracellular proteolytic activity of cathepsin L to stages VI and VII of the cycle. Cystatin C is a physiologically important inhibitor of cysteine proteinases such as cathepsins B, H, L, and S (reviewed in [134]). The affinity of cystatin C for these four cathepsins is very high, displaying Ki values in the nanomolar and picomolar range [134]. By its ability to inhibit several cathepsins, cystatin C may restrict their enzymatic and biological activities to a specific microenvironment within a tissue, such as the seminiferous epithelium in the testis. Three members of the second group of the family 2 cystatins are expressed in Sertoli cells: testatin and cystatins SC and TE-1 (also called CRES3). Northern blot analysis using RNA isolated from 14 mouse tissues revealed that cystatin SC transcripts are only detected in the testis, whereas cystatin TE-1 transcripts are detected at high levels in the epididymis and the testis and at low levels in the ovary and the prostate [199, 200]. In situ hybridization showed that cystatin SC transcripts were localized mainly in Sertoli cells, with low levels in tubules between stages VI to VIII, whereas cystatin TE-1 mRNA was expressed in Sertoli cells in tubules at all stages of the cycle [199, 200]. Levels of both cystatins SC and TE-1 transcripts vary during testis development [199]. They are detected at birth and become highly expressed between postnatal days 15 and 30. However, by postnatal day 35, both transcripts are barely detectable in testes. Note also that, contrary to the levels of cystatin SC mRNA, levels

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of cystatin TE-1 transcripts decreased between birth and postnatal day 10. The authors raised the possibility that in the absence of cystatin TE-1, unidentified proteases may be activated to participate in the migration of postnatal gonocytes [199]. Testatin (testis-specific cystatin-related gene) displays a very restricted pattern of expression. It is detected only in fetal gonads and in adult testes [201]. In situ hybridization analysis and immunostaining with a testatin-specific antibody on sections of fetal testis show that testatin mRNA and protein are expressed in pre-Sertoli cells and upregulated soon after the testisdetermining gene Sry is expressed [201]. Testatin mRNA is present in both XX and XY gonads at 11.5 days postcoitus (dpc). Between 11.5 and 12.5 dpc, transcription of this gene increases in XY gonads but decreases in the XX gonads. This expression pattern is in contrast to the expression of cystatin C and CRES, which are almost uniformly expressed throughout gonad development. Testatin is also expressed in Sertoli cells of adult testis [201]. b. Reproductive Phenotype Observed in Knockout Mice Of all the cystatin family members expressed in Sertoli cells, only the cystatin C gene has been inactivated in mice [202]. Cystatin C–deficient mice are born healthy and are fertile. Examination of several tissues did not reveal any abnormalities during the course of the experiments (up to 6 months). 2. The α-Macroglobulin Family a. Properties Unrelated to the members of the cystatin superfamily, the α2-macroglobulin is a member of a group of high molecular weight proteins known collectively as the α-macroglobulins. The α-macroglobulins are the only natural inhibitors of all four classes of proteinases (reviewed in [203, 204]). Mechanistically, the inhibition operates through a “molecular trap” in which the active site of the proteinases is preserved but access to it is reduced. Binding of proteinases to α2-macroglobulin is irreversible. Based on the determination of association rate constants, α2-macroglobulin appears to be a slower inhibitor than cystatin C [205]. b. Expression in Sertoli Cells Rat α2-macroglobulin is expressed in Sertoli cells in a stage-specific manner, secretion of α2-macroglobulin being maximal in stages II to VIII tubules [206–208]. c. Reproductive Phenotype Observed in Knockout Mice As stated in an earlier section, α2-macroglobulin– deficient mice are viable and fertile, producing normal

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sized litters. Homozygous mice show no phenotypic abnormalities during the course of the experiments (up to 15 months of age) [103, 209]. This result is very surprising in light of the fact that α2-macroglobulin inhibits a wide variety of proteinases.

V. MORE THAN MEETS THE EYE: LESSONS FROM CATHEPSIN L–DEFICIENT MICE A. Knockout Mice Producing No Reproductive Phenotype Understanding the normal physiological functions of proteinases and their complementary inhibitors has been aided by the production of knockout mouse models. As stated in Sections II, III, and IV of this chapter, many proteinases and proteinase inhibitors have been inactivated in mice. Because the expression or activity of many of these genes is highly regulated in the seminiferous epithelium (and, more specifically, in Sertoli cells), it was expected that mice in which individual genes had been inactivated would show defects during spermatogenesis, perhaps leading to male infertility. However, with the exception of mice carrying a disrupted ADAM-TS2 or MT1-MMP gene, all knockout mice produced to date are fertile. The absence of a reproductive phenotype raises more questions than answers. For instance, does the absence of a reproductive phenotype indicate that the function of these proteinases and proteinase inhibitors are dispensable in the testis? This may be a possibility, but it is difficult to accept the idea that proteins do not have any role in tissues in which their expression or activity is highly regulated. Another question is “Does the lack of reproductive phenotype suggest that a genetic redundancy may exist among members of the proteinase and proteinase inhibitor families?” This redundancy could be at the level of (1) a functional redundancy of another family member or (2) an upregulation in the expression of a related member that can functionally replace a missing gene when needed. If such a redundant system is operating, then crosses between mice in which a single gene has been disrupted may provide some clues as far as the functions of these proteins in the seminiferous epithelium are concerned. Such crosses have already been performed for some of the cathepsin family members. The reported phenotypes range from normal to lethal. For example, mice in which both cathepsins B and K are disrupted show no aberrant phenotype [118]. Cathepsin K- and L–deficient mice were reported to be smaller in size compared to

wild-type littermates [118]. Finally, mice that lack both cathepsin B and L genes die during the second to fourth week of life because of brain atrophy [210]. From these results, it appears that this strategy may not shed additional light on the possible roles of proteinases and their complementary inhibitors in the seminiferous epithelium. However, it is important to emphasize that in most studies fertility is measured by assessing whether or not knockout mice are able to produce a number of viable offspring similar to that of their wild-type counterparts. Is this criterion sufficient to rule out any potential defects during spermatogenesis in these mutant mice? Relevant to this point, it should be stressed that in rodents, sperm production can decrease as much as fivefold without affecting fertility [211]. This observation raises the following question: Would it be worthwhile to perform a thorough analysis of testes from mice in which the gene coding for a proteinase or proteinase inhibitor has been disrupted? Would such studies reveal new insights into the role of these proteins during spermatogenesis? A study, described next, which carefully examined the testes from mice deficient in functional cathepsin L, provides answers to these questions.

B. A Closer Look at the Testes of Cathepsin L–Deficient Mice The phenotype of mice in which the cathepsin L gene has been inactivated [120] recapitulates that of the spontaneous mouse mutant furless [151]. A glycine-toarginine mutation in the cathepsin L gene has been identified in these mice. This mutation at amino acid 149 abolishes the enzymatic activity of the protein. To evaluate whether or not cathepsin L is required for normal spermatogenesis, a thorough morphological analysis of the testes of furless mice was undertaken [140]. The results from this study can be summarized as follows: 1. Testes of 110- to 120-day-old furless mice are significantly (25%) smaller than those of wild-type and heterozygous littermates. 2. There was at least a 10-fold increase in seminiferous tubule atrophy in testes of furless mice. This atrophy was heterogeneous, ranging from vacuolization of the seminiferous epithelium and loss of one or more generations of germ cells, to a Sertoli cell-only phenotype (Fig. 9.12A). Most atrophic tubules were at least missing spermatids. 3. The nonatrophic (normal) tubules seen in testes of furless mice were also affected by the absence of cathepsin L enzymatic activity. In comparison to littermate controls, 16% fewer preleptotene

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maturation into pachytene spermatocytes. However, no detectable effect on the maturation of pachytene spermatocytes into round spermatids was observed. It can be concluded that the lack of cathepsin L affects a different subset of germ cells in nonatrophic (normal) tubules than in atrophic tubules and quantitatively reduces the efficiency of spermatogenesis.

A

The results summarized above suggest that cathepsin L has multiple functions in the seminiferous epithelium. However, little information is currently available about what these functions are. As mentioned in Section IV.B.3, immunocytochemical analysis has demonstrated that rat cathepsin L is concentrated around most elongate spermatids in tubules at stages VI to VIII [138] and in germ cells undergoing apoptosis [139]. In addition, it appears that cathepsin L is localized to residual bodies as soon as they are released from elongate spermatids and subsequently phagocytosed by Sertoli cells (Fig. 9.13). Collectively, these results indicate that cathepsin L may facilitate the release of elongate spermatids from Sertoli cells and/or the clearance of residual bodies and dying cells from the seminiferous epithelium. To explain why mice lacking cathepsin L are not infertile, it can be hypothesized that another proteinase(s) is localized to the same structures as

B

FIGURE 9.12 Cathepsin L is required for normal quantitative spermatogenesis. (A) Morphology of a tubule from the testis of a normal mouse (left panel) compared to an atrophic tubule from the testis of a furless mouse (right panel). The Sertoli-only phenotype indicates the absence of germ cells in the tubule. (Source: Figure 3 of Wright, W. W., Smith, L., Kerr, C., and Charron, M., Biology of Reproduction, Vol. 68, Mice that express enzymatically inactive cathepsin L exhibit abnormal spermatogenesis, pp. 680–687. Copyright 2003 Society for the Study of Reproduction.) (B) A significant reduction in the number per Sertoli cell of round spermatids (68% of control) and pachytene spermatocytes (69% of control), and in the total number of preleptotene spermatocytes formed (84% of control) is observed in nonatrophic tubules from testes of furless mice. (Source: Figure 5 of Wright, W. W., Smith, L., Kerr, C., and Charron, M., Biology of Reproduction, Vol. 68, Mice that express enzymatically inactive cathepsin L exhibit abnormal spermatogenesis, pp. 680–687. Copyright 2003 Society for the Study of Reproduction.)

spermatocytes per Sertoli cell and 32% fewer pachytene spermatocytes and round spermatids per Sertoli cell were observed in testes of furless mice (Fig. 9.12B). This result indicated that the absence of cathepsin L enzymatic activity reduced the production of early spermatocytes and their

FIGURE 9.13 Cathepsin L is localized to residual bodies after they have been engulfed by Sertoli cells (see color plate). Cathepsin L immunostaining is shown in green. Nucleic acids were stained with propidium iodide (red). Colocalization produces a yellow signal. Arrows identify residual bodies in a tubule at stage VII (left panel) or VIII (right panel). The arrowheads (left panel) identify the nuclei of step 19 spermatids. Twenty continous 0.5-μm sections were stacked together to generate each picture. The asterisk (*) points to the base of the tubule and the arrowheads identify elongate spermatids. The horizontal white bar is equal to 10 μm.

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cathepsin L and can degrade an overlapping subset of substrates normally processed by cathepsin L.

VI. FUTURE DIRECTIONS Since the first comprehensive review on this topic by Fritz et al. [212] in 1993, the growth of information regarding the expression of proteases and their complementary inhibitors in the seminiferous epithelium and, in particular, in Sertoli cells has been rapid. However, several lines of investigation are needed to further our understanding of the physiological functions of these enzymes in this specialized tissue. One line of investigation is the careful examination of testes from mice deficient in specific proteinases or proteinase inhibitors. The morphological analysis of the seminiferous epithelium of cathepsin L–deficient mice has demonstrated that, although a lack of this proteinase may not cause infertility, it nonetheless causes tubule atrophy in a subset of seminiferous tubules and also diminished the formation and maturation of preleptotene spermatocytes in other, apparently normal tubules (Section V). These results, combined with the stage-specific expression of the cathepsin L gene in murine Sertoli cells, support the hypothesis that this proteinase has important biological functions in the seminiferous epithelium. Based on the results obtained from the examination of testes from cathepsin L-deficient mice, careful morphological analyses of testes from the knockout mouse models already available (see Sections II, III, and IV) should be performed to establish whether or not the absence of a given proteinase or proteinase inhibitor leads to major defects in the seminiferous epithelium and, consequently, to reduced sperm production. These quantitative morphological analyses are a high priority for future research. The significance of the redundancy proposed for many proteinases and proteinase inhibitors expressed in Sertoli cells awaits to be tested experimentally. As pointed out in Sections II, III, and IV, mice deficient in a single proteinase or proteinase inhibitor are, in most cases, fertile. To explain the lack of a reproductive phenotype, it was suggested that a redundancy exists between specific proteinase or proteinase inhibitor members (Section V). If this redundancy model is accurate, it is reasonable to expect that an abnormal reproductive phenotype might be observed in mice lacking two or more proteinases or proteinase inhibitors. However, as discussed in Section V, mice carrying two disrupted proteinase genes can display severe abnormalities and prematurely die shortly after birth

(e.g., mice deficient in both cathepsins B and L). The use of conditional gene disruption technologies such as those based on the Cre-loxP system [213], combined with Sertoli cell–specific promoters, will be required to carry out these experiments. Note that transgenic mice in which the anti-Müllerian hormone promoter selectively drives expression of Cre recombinase in Sertoli cells have already been generated [214]. Although mice deficient in a single proteinase or proteinase inhibitor often appear fertile, it is important to recall that mice deficient in MT1-MMP, ADAM-TS4, or PCI are infertile because they produce few (if any) sperm (Sections II.D and III.C). However, one cannot conclude that these proteinases or proteinase inhibitors have a direct role in the seminiferous epithelium. It is important to emphasize that proteinases expressed in the seminiferous epithelium are also detected in many other organs. Thus, the reproductive phenotype observed in these three knockout mouse models could be secondary to lesions in other organs, such as the pituitary. This dilemma could be solved by conditional disruption of these genes in Sertoli cells. Another potential consequence of this apparent redundancy is that deficiency in one proteinase or proteinase inhibitor can lead to a compensatory increase in the expression of another family member or an unrelated protein with equivalent activity. Precedence for this scenario comes from a study that showed upregulation of cathepsin L expression in thyroids of cathepsin K–deficient mice [118]. It would be worthwhile to establish if upregulation of a proteinase(s) or a proteinase inhibitor(s) occurs in the seminiferous epithelium of the single knockout mouse models already available. Gene array analysis comparing transcript profiles between single knockout and normal mice should begin to address this point. In view of the fact that proteinases are under the control of their complementary inhibitors, experiments should be designed to determine if the interplay between these enzymes is crucial in the regulation of the seminiferous epithelium. As discussed in Section IV.H.1.a, Tsuruta et al. [198] reported that the stagespecific expression of cystatin C is the opposite of the one described for cathepsin L. The authors proposed that this “yin–yang” relationship could help cystatin C limit the extracellular proteolytic activity of cathepsin L to stages VI to VII of the cycle of the seminiferous epithelium. In view of these results, it might be interesting to alter the stage-specific expression of proteinase inhibitors in Sertoli cells and study the repercussions of such deregulation on the organization of the seminiferous epithelium. For example, altered expression of cystatin C in Sertoli cells at stages VI to VIII could be achieved by generating transgenic

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mice in which the coding sequence of cystatin C is fused to the 3-kb promoter region of the rat cathepsin L gene. The functions of testatin and cystatins SC and TE-1 in Sertoli are still a mystery. As discussed in Section IV, these members of family 2 cystatins share two common characteristics: (1) They possess only one of the three conserved domains in cystatins required to block the active site of cysteine proteinases, and (2) they are all primarily expressed in reproductive tissues (reviewed in [196]). Several questions regarding testatin and cystatins SC and TE-1 need to be addressed: Have they retained the ability to inhibit cathepsins? Have they evolved new functions? What are their target(s) in Sertoli cells? Finally, it will be important to establish the location of biologically important proteinases and their respective inhibitors in the seminiferous epithelium. Given the morphological complexity of the seminiferous epithelium, immunocytochemistry coupled with electron microscopy will have to be performed. Only by this means will potential targets (such as those illustrated in Fig. 9.1) be proven to be sites of action of proteinases or proteinase inhibitors. Of course, their cellular localization does not identify their substrate(s). Additional experiments such as comparing the proteome of seminiferous tubules from wild-type and relevant knockout mice or the identification of specific substrates for a given proteinase by coimmunoprecipitation experiments will need to be performed. In this chapter, it has been argued that proteinases and their complementary inhibitors are potentially important regulators of spermatogenesis by their ability to alter the architecture of the seminiferous epithelium and the paracrine environment of male germ cells. While good progress has been made toward proving the accuracy of this statement, it is clear that much work remains to be carried out. However, it is fitting that this chapter ends with the same enthusiastic sign off as the one found in the chapter on proteases and antiproteases written by Fritz et al. [212] in 1993: “Who knows what excitement lies ahead?”

VII. RESOURCES ON THE WORLD WIDE WEB Additional information on proteases and their complementary inhibitors are readily available through comprehensive and searchable databases [215–218]. Their URL addresses are as follows: Gene grouping and family nomenclature

http://ash.gene.ucl.ac.uk/ nomenclature/ genefamily.shtml

Interpro Merops Pfam

http://www.ebi.ac.uk/interpro http://merops.ac.uk http://www.sanger.ac.uk/ Software/Pfam/ As pointed out in different sections of this chapter, the expression of some proteinases and proteinase inhibitors in testes and/or Sertoli cells was inferred from data collected by searching two databases. Their URL addresses are as follows: Sertoli cell cDNA http://www.ncbi.nlm.nih.gov/ library (Unigene UniGene/library.cgi?all=yes& database) ORG=Mm&LID=12732 Testis cDNA library http://cgap.nci.nih.gov/ (Cancer Genome Genes/GeneFinder Anatomy Project database)

Acknowledgments Research on Sertoli cells conducted in the authors’ laboratory is supported by the NICHD (R01 HD044183). We thank Ms. Janet Folmer for the electron micrographs used in Figure 9.1. We also thank the following people for critical reviews or proofreading of the manuscript: Dr. Barry Zirkin, Dr. Terry Brown, Dr. Joel Shaper, Ms. Jeannie Chern, and Ms. Pamela Wright.

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Chapter 9 Proteases and Protease Inhibitors 174. D’Azzo, A., Hoogeveen, A., Reuser, A. J., Robinson, D., and Galjaard, H. (1982). Molecular defect in combined betagalactosidase and neuraminidase deficiency in man. Proc. Natl. Acad. Sci. USA 79, 4535–4539. 175. Verheijen, F., Brossmer, R., and Galjaard, H. (1982). Purification of acid beta-galactosidase and acid neuraminidase from bovine testis: evidence for an enzyme complex. Biochem. Biophys. Res. Commun. 108, 868–875. 176. Hoogeveen, A. T., Verheijen, F. W., and Galjaard, H. (1983). The relation between human lysosomal beta-galactosidase and its protective protein. J. Biol. Chem. 258, 12143–12146. 177. Yamamoto, Y., and Nishimura, K. (1987). Copurification and separation of beta-galactosidase and sialidase from porcine testis. Int. J. Biochem. Cell. Biol. 19, 435–442. 178. Bonten, E., van der Spoel, A., Fornerod, M., Grosveld, G., and d’Azzo, A. (1996). Characterization of human lysosomal neuraminidase defines the molecular basis of the metabolic storage disorder sialidosis. Genes Dev. 10, 3156–3169. 179. van der Spoel, A., Bonten, E., and d’Azzo, A. (1998). Transport of human lysosomal neuraminidase to mature lysosomes requires protective protein/cathepsin A. EMBO J. 17, 1588–1597. 180. Jackman, H. L., Tan, F. L., Tamei, H., Beurling-Harbury, C., Li, X. Y., Skidgel, R. A., and Erdos, E. G. (1990). A peptidase in human platelets that deamidates tachykinins. Probable identity with the lysosomal “protective protein.” J. Biol. Chem. 265, 11265–11272. 181. Hiraiwa, M. (1999). Cathepsin A/protective protein: An unusual lysosomal multifunctional protein. Cell. Mol. Life. Sci. 56, 894–907. 182. Rottier, R. J., Hahn, C. N., Mann, L. W., del Pilar Martin, M., Smeyne, R. J., Suzuki, K., and d’Azzo, A. (1998). Lack of PPCA expression only partially coincides with lysosomal storage in galactosialidosis mice: Indirect evidence for spatial requirement of the catalytic rather than the protective function of PPCA. Hum. Mol. Genet. 7, 1787–1794. 183. Sohma, O., Mizuguchi, M., Takashima, S., Satake, A., Itoh, K., Sakuraba, H., Suzuki, Y., and Oyanagi, K. (1999). Expression of protective protein in human tissue. Pediatr. Neurol. 20, 210–214. 184. Luedtke, C. C., Andonian, S., Igdoura, S., and Hermo, L. (2000). Cathepsin A is expressed in a cell- and region-specific manner in the testis and epididymis and is not regulated by testicular or pituitary factors. J. Histochem. Cytochem. 48, 1131–1146. 185. Zhou, X. Y., Morreau, H., Rottier, R., Davis, D., Bonten, E., Gillemans, N., Wenger, D., Grosveld, F. G., Doherty, P., Suzuki, K., and et al. (1995). Mouse model for the lysosomal disorder galactosialidosis and correction of the phenotype with overexpressing erythroid precursor cells. Genes Dev. 9, 2623–2634. 186. Korah, N., Smith, C. E., D’Azzo, A., El-Alfy, M., and Hermo, L. (2003). Increase in macrophages in the testis of cathepsin A deficient mice suggests an important role for these cells in the interstitial space of this tissue. Mol. Reprod. Dev. 64, 302–320. 187. Faust, P. L., Kornfeld, S., and Chirgwin, J. M. (1985). Cloning and sequence analysis of cDNA for human cathepsin D. Proc. Natl. Acad. Sci. USA 82, 4910–4914. 188. Hasilik, A., von Figura, K., Conzelmann, E., Nehrkorn, H., and Sandhoff, K. (1982). Lysosomal enzyme precursors in human fibroblasts. Activation of cathepsin D precursor in vitro and activity of beta-hexosaminidase A precursor towards ganglioside GM2. Eur. J. Biochem. 125, 317–321. 189. Isidoro, C., Demoz, M., De Stefanis, D., Mainferme, F., Wattiaux, R., and Baccino, F. M. (1995). Altered intracellular processing and enhanced secretion of procathepsin D in a highly deviated rat hepatoma. Int. J. Cancer 60, 61–64. 190. Isidoro, C., Demoz, M., De Stefanis, D., Baccino, F. M., Hasilik, A., and Bonelli, G. (1997). Differential targeting and processing

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of procathepsin D in normal and transformed murine 3T3 fibroblasts. Int. J. Cancer 70, 310–314. Srivastava, P. N., and Ninjoor, V. (1982). Isolation of rabbit testicular cathepsin D and its role in the activation of proacrosin. Biochem. Biophys. Res. Commun. 109, 63–69. Xing, W., and Archer, T. K. (1998). Upstream stimulatory factors mediate estrogen receptor activation of the cathepsin D promoter. Mol. Endocrinol. 12, 1310–1321. Rochefort, H., Chalbos, D., Cunat, S., Lucas, A., Platet, N., and Garcia, M. (2001). Estrogen regulated proteases and antiproteases in ovarian and breast cancer cells. J. Steroid Biochem. Mol. Biol. 76, 119–124. Saftig, P., Hetman, M., Schmahl, W., Weber, K., Heine, L., Mossmann, H., Koster, A., Hess, B., Evers, M., von Figura, K., and et al. (1995). Mice deficient for the lysosomal proteinase cathepsin D exhibit progressive atrophy of the intestinal mucosa and profound destruction of lymphoid cells. EMBO J. 14, 3599–3608. Turk, V., and Bode, W. (1991). The cystatins: protein inhibitors of cysteine proteinases. FEBS Lett. 285, 213–219. Cornwall, G. A., and Hsia, N. (2003). A new subgroup of the family 2 cystatins. Mol. Cell. Endocrinol. 200, 1–8. Cornwall, G. A., Cameron, A., Lindberg, I., Hardy, D. M., Cormier, N., and Hsia, N. (2003). The cystatin-related epididymal spermatogenic protein inhibits the serine protease prohormone convertase 2. Endocrinology 144, 901–908. Tsuruta, J. K., O’Brien, D. A., and Griswold, M. D. (1993). Sertoli cell and germ cell cystatin C: Stage-dependent expression of two distinct messenger ribonucleic acid transcripts in rat testes. Biol. Reprod. 49, 1045–1054. Li, Y., Friel, P. J., Robinson, M. O., McLean, D. J., and Griswold, M. D. (2002). Identification and characterization of testis- and epididymis-specific genes: Cystatin SC and cystatin TE-1. Biol. Reprod. 67, 1872–1880. Hsia, N., and Cornwall, G. A. (2003). Cres2 and Cres3: New members of the cystatin-related epididymal spermatogenic subgroup of family 2 cystatins. Endocrinology 144, 909–915. Tohonen, V., Osterlund, C., and Nordqvist, K. (1998). Testatin: A cystatin-related gene expressed during early testis development. Proc. Natl. Acad. Sci. USA 95, 14208–14213. Huh, C. G., Hakansson, K., Nathanson, C. M., Thorgeirsson, U. P., Jonsson, N., Grubb, A., Abrahamson, M., and Karlsson, S. (1999). Decreased metastatic spread in mice homozygous for a null allele of the cystatin C protease inhibitor gene. Mol. Pathol. 52, 332–340. Sottrup-Jensen, L. (1989). Alpha-macroglobulins: structure, shape, and mechanism of proteinase complex formation. J. Biol. Chem. 264, 11539–11542. Salvesen, G., and Nagase, H. (1989). Inhibition of proteolytic enzymes. In “Proteolytic enzymes: A practical approach” (R. J. Beynon and J. S. Bond, eds.), pp. 83–104. IRL Press, Oxford, England. Peloille, S., Esnard, A., Dacheux, J. L., Guillou, F., Gauthier, F., and Esnard, F. (1997). Interactions between ovine cathepsin L, cystatin C and alpha 2-macroglobulin. Potential role in the genital tract. Eur. J. Biochem. 244, 140–146. Cheng, C. Y., Grima, J., Stahler, M. S., Guglielmotti, A., Silvestrini, B., and Bardin, C. W. (1990). Sertoli cell synthesizes and secretes a protease inhibitor, alpha 2-macroglobulin. Biochemistry 29, 1063–1068. Kangasniemi, M., Cheng, C. Y., Toppari, J., Grima, J., Stahler, M., Bardin, C. W., and Parvinen, M. (1992). Basal and FSHstimulated steady state levels of SGP-2, alpha 2-macroglobulin,

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Martin Charron and William W. Wright and testibumin in culture media of rat seminiferous tubules at defined stages of the epithelial cycle. J. Androl. 13, 208–213. Mruk, D., Zhu, L. J., Silvestrini, B., Lee, W. M., and Cheng, C. Y. (1997). Interactions of proteases and protease inhibitors in Sertoli-germ cell cocultures preceding the formation of specialized Sertoli-germ cell junctions in vitro. J. Androl. 18, 612–622. Umans, L., Serneels, L., Overbergh, L., Stas, L., and Van Leuven, F. (1999). Alpha2-macroglobulin- and murinoglobulin-1- deficient mice. A mouse model for acute pancreatitis. Am. J. Pathol. 155, 983–993. Felbor, U., Kessler, B., Mothes, W., Goebel, H. H., Ploegh, H. L., Bronson, R. T., and Olsen, B. R. (2002). Neuronal loss and brain atrophy in mice lacking cathepsins B and L. Proc. Natl. Acad. Sci. USA 99, 7883–7888. Chapin, R. E., Sloane, R. A., and Haseman, J. K. (1997). The relationships among reproductive endpoints in Swiss mice, using the reproductive assessment by Continuous Breeding database. Fundam Appl Toxicol 38, 129–142. Fritz, I. B., Tung, P. S., and Ailenberg, M. (1993). Proteases and antiproteases in the seminiferous tubule. In “The Sertoli Cell” (L. D. Russel and M. D. Griswold, eds.), pp. 217–235. Cache River Press, Clearwater, FL.

213. Sauer, B. (1998). Inducible gene targeting in mice using the Cre/lox system. Methods 14, 381–392. 214. Lecureuil, C., Fontaine, I., Crepieux, P., and Guillou, F. (2002). Sertoli and granulosa cell-specific Cre recombinase activity in transgenic mice. Genesis 33, 114–118. 215. Barrett, A. J., Rawlings, N. D., and O’Brien, E. A. (2001). The MEROPS database as a protease information system. J. Struct. Biol. 134, 95–102. 216. Barrett, A. J., Tolle, D. P., and Rawlings, N. D. (2003). Managing peptidases in the genomic era. Biol. Chem. 384, 873–882. 217. Mulder, N. J., Apweiler, R., Attwood, T. K., Bairoch, A., Barrell, D., Bateman, A., Binns, D., Biswas, M., Bradley, P., Bork, P., Bucher, P., Copley, R. R., Courcelle, E., Das, U., Durbin, R., Falquet, L., Fleischmann, W., Griffiths-Jones, S., Haft, D., Harte, N., Hulo, N., Kahn, D., Kanapin, A., Krestyaninova, M., Lopez, R., Letunic, I., Lonsdale, D., Silventoinen, V., Orchard, S. E., Pagni, M., Peyruc, D., Ponting, C. P., Selengut, J. D., Servant, F., Sigrist, C. J., Vaughan, R., and Zdobnov, E. M. (2003). The InterPro Database, 2003 brings increased coverage and new features. Nucleic Acids Res. 31, 315–318. 218. Studholme, D. J., Rawlings, N. D., Barrett, A. J., and Bateman, A. (2003). A comparison of Pfam and MEROPS: Two databases, one comprehensive, and one specialised. BMC Bioinformatics 4, 17.

P A R T

IV SERTOLI CELL ENDOCRINOLOGY AND SIGNAL TRANSDUCTION

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C H A P T E R

10 FSH Regulation at the Molecular and Cellular Levels: Mechanisms of Action and Functional Effects ILPO HUHANIEMI

JORMA TOPPARI

Institute of Reproductive and Developmental Biology, Imperial College, London, United Kingdom; and Department of Physiology, University of Turku, Turku, Finland

Departments of Physiology and Pediatrics, University of Turku, Turku, Finland

I. INTRODUCTION II. STRUCTURES OF THE FSH AND FSHR GENES AND PROTEINS III. MOLECULAR ASPECTS OF FSHR FUNCTION IV. MODIFICATION OF FSH ACTION IN SERTOLI CELLS BY SPERMATOGENIC CELLS References

II. STRUCTURES OF THE FSH AND FSHR GENES AND PROTEINS A. FSH Genes and Protein Like the other glycoprotein hormones, luteinizing hormone (LH), thyroid-stimulating hormone (TSH), and human chorionic gonadotropin (hCG), FSH is composed of two protein subunits, the common α-subunit and the hormone-specific β-subunit (Fig. 10.1), coupled by noncovalent interactions. The common α-subunit gene is localized in humans on chromosome 6q12.21 and it consists of four exons, of which the first one is noncoding [1]. The α-subunit propeptide has a 24-amino-acid signal peptide in its N-terminal end, which is cleaved during the process of synthesis and secretion of the mature hormone dimers. The mature α-subunit is 92 amino acids long, and it contains 10 cysteines participating in intrasubunit disulfide linkages and two N-linked carbohydrate side chains are attached to Asn52 and Asn78. The FSHβ gene is localized on chromosome 11p13 and it consists of three exons, the first one being noncoding [1]. Its signal sequence and mature peptide are 18 and 111 amino acids in length, respectively. Like the other β-subunits, that of FSH contains 12 cysteines for disulfide formation, and its two N-linked carbohydrate side chains are located on Asn7 and Asn24. About 20% of the weight of FSH is accounted for by the carbohydrate side chains. Typical of all glycoprotein

I. INTRODUCTION Follicle-stimulating hormone (FSH) is the main trophic hormone regulating Sertoli cell function and, along with paracrine action of the Leydig cell product, testosterone (T), these two hormones maintain Sertoli cell functions to ensure the maintenance of qualitatively and quantitatively normal spermatogenesis. FSH action on Sertoli cells is mediated by a specific receptor, the FSH receptor (R). Besides this chapter, functions of FSH and FSHR are reviewed in Chapters 11, 16, and 23, which concentrate on the physiological aspects of FSH action, structure and regulation of expression of FSHR. The purpose of this chapter is to elucidate the molecular aspects of FSH action on Sertoli cells, including FSHR function, the biochemical responses evoked by FSH stimulation, and their dependence on the paracrine regulatory milieu determined by the functional state of the seminiferous epithelial cycle. SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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156

Ilpo Huhtaniemi and Jorma Toppari Common α-subunit

Gene 1 kb

Exon 1

2

3 4

Protein 10 aa −24

1

92 52

78

FSH β-subunit Gene

Exon 1

2

3

Protein

−18

1

111 7

24

FIGURE 10.1 Schematic presentation of the human common α and FSHβ subunits. The top parts of each scheme depict the gene structures. The open bars indicate noncoding, and the filled bars coding sequences of exons. The genes are drawn to scale. The dotted lines combine the exons to the approximate regions of the proteins that they encode, presented in the bottom parts of the schemes. The shaded areas indicate the signal peptides and the open bars the mature proteins. The numbers below the bars indicate the starts and ends of the signal peptides and the lengths of the mature proteins. The triangles below the proteins indicate the positions of the N-linked glycosylation sites.

hormones, the carbohydrate side chains of FSH show considerable microheterogeneity between individual molecules [2]. Besides prolonging the circulatory halflife, the carbohydrate composition apparently plays a role in heterodimer stability and in the intrinsic bioactivity of a molecule, which may vary according to the functional state of pituitary gonadotropin synthesis and secretion [3, 4]. The functional significance of this phenomenon has not been thoroughly studied, but there are differences between the in vitro and in vivo bioactivities of the different glycoforms. Their composition in circulation is dependent on the hormonal status of an individual, so the glycoform composition of circulating FSH may play a role in the fine-tuning of its actions. Deglycosylation of gonadotropins does not affect receptor binding but abolishes their signal transduction at the receptor site [5, 6]. The crystal structure of the human FSH was recently published [7] (Fig. 10.2). Like hCG, which was

FIGURE 10.2 The three-dimensional structure of FSH. The α-subunit is depicted with purple and β-subunit with green (see color plate). Ball-and-stick models present the carbohydrate side chains and disulfide bonds. The “seat belt” of the β-subunit that extends around the α-subunit is depicted with an arrow. (From [7], with permission.)

crystallized earlier [8], FSH belongs to the superfamily of cystine knot growth factors characterized by a cluster of three cystine disulfide bonds in each subunit. Similar folding is found in some protein growth factors, such as nerve growth factor (NGF), transforming growth factor β (TGFβ), and platelet-derived growth factor β (PGFβ). Despite dissimilar amino acid structures, the α- and β-subunits have remarkably similar three-dimensional structures, including two β-hairpins on one side, and one on the other side of the cystine knot. The β-hairpins are stabilized by additional disulfide bridges. The subunits are associated in a head-to-tail orientation forming an elongated slightly curved structure (Fig. 10.2). The subunit dimer is stabilized by a “seat belt” structure that is formed by the C-terminal amino acids of the β-subunit turning around the α-subunit and stabilized by one of the disulfide bonds. Compared with the crystal structure of hCG [8], FSH has some special features that

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might be important for the specificity of hormone action. These include conformational changes and/or differential distribution of polar or charged residues at the ends of the hairpin loops, in the cystine noose (a short loop in the middle of the molecule), and differences in the surface characteristics of hydrophobic patch areas. Many of these differences are in the concave side of the molecule, resulting in a differential face from that of hCG in the area participating in receptor binding. Glycosylation does not have global effects on gonadotropin conformation [7, 8].

B. FSHβ Subunit Mutations and Their Functional Consequences Concerning mutations in the FSH subunit genes, the only ones detected so far have been found in the β-subunit. Common α mutations may be lethal because of concomitant inactivation of hCG and TSH production. Concerning the FSHβ subunit mutations, three have been detected in men presenting with azoospermia [9–11]. The phenotype is surprising because neither FSHβ and FSHR knockout mice [12–14] nor men with inactivating FSHR mutations [15] are azoospermic. Detailed analysis of mutations at the molecular level has increased our understanding of the structure– function relationships of FSH action. One of the mutations, detected in a hypogonadal woman, induced a 2-bp deletion in the codon of Val61 [16], which gives rise to a completely altered amino acid sequence between codons 61 and 86, followed by a premature stop codon. The translation product of the mutated gene was unable to associate with the common α-subunit and to form bioactive FSH, apparently because the C terminus of FSHβ, contributing to the dimer stabilizing “seat belt,” one of the cysteines contributing to the cystine knot, and an amino acid forming the cystine noose structure were missing [7]. Another FSHβ mutation was a missense T-to-G transversion, causing a Cys51 to Gly mutation [17]. Cells transfected with this mutant FSHβ along with wild-type α-subunit failed to produce immunoreactive FSH, apparently because of the loss of a cysteine participating in formation of the cystine-knot structure, which is essential for organizing the core of the protein and determining its folding. A third FSHβ mutation with functional impact had a point mutation that changed codon 76 from Tyr to a stop codon [9]. The interference of the missing C-terminal amino acids with the cystine-knot, cystinenoose, and seat belt formation explains the lack of synthesis of functional FSH in the affected individuals. An additional inactivating mutation, Cys82Arg, also affects the cystine-knot formation [10]. Hence, the inactivation mechanisms of the FSHβ mutations

detected thus far can be explained by dramatic alterations in the key tertiary structures of the wild-type hormone.

C. FSHR Gene and Protein The FSHR gene is structurally very similar to those of LHR and TSHR, belonging to the superfamily of G-protein–associated receptors (GPCRs), sharing with all of them the seven-time transmembrane structure, but differing from the other GPCRs by their long extracellular domain (Fig. 10.3). FSHR is encoded by a single gene that has the same chromosomal location as LHR, that is, chromosome 2p21 [3, 18]. The FSHR gene is about 215 kb in size, and it consists of 10 exons and 9 intervening introns. As with LHR, the first 9 exons (10 in LHR) encode the extracellular domain, which constitutes about half of the size of the mature receptor protein. The last long exon encodes the C-terminal end of the hinge region, the transmembrane region, and the intracellular tail. The Sertoli cell is undisputedly the only site of FSHR expression in the male, although there are some older reports of its expression in spermatogonia [19], and in the female in extragonadal tissues [20]. The FSHR gene is discussed in more details in Chapter 16. The mature human FSHR polypeptide chain is composed of 678 residues, and the primary sequence contains an additional 17-amino-acid leader/signal peptide [3, 18] (Fig. 10.3). The molecular mass of the

FSH receptor Exon

1

2

3 4 5,6

7,8

9

10

Gene

Protein −17 1

678 174 182

FIGURE 10.3

274 301

Schematic presentation of the human FSH receptor gene and protein. The top part of the scheme depicts the gene structure. The open bars indicate noncoding, and the filled bars coding sequences of exons. The total length of the gene is about 215 kb. The dotted lines combine the exons to the approximate regions of the protein that they encode. Notably, the long 10th exon encodes about half of the protein, including its entire transmembrane region. The bottom part of the scheme presents the protein structure. The horizontal hatched part of the protein indicates the signal peptide, and the crosshatched bars signify the seven segments of the transmembrane domain. The numbers below the protein bar indicate the start and end of the signal peptide and the length of the mature protein. The triangles below the protein bar indicate the positions of the N-linked glycosylation sites.

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native receptor protein expressed at the target cell surface is 85–90 kDa, which is larger that the predicted size due to glycosylation. The extracellular domain of the human receptor has four consensus sites for glycosylation signals, and when compared with the other glycoprotein hormone receptors, only two are conserved. Glycosylation is most likely concerned with stability of the receptor, the process of folding of the nascent protein on its synthesis, and transit to the cell membrane. Site-directed mutagenesis studies on abolition of some or all of the glycosylation sites indicate that carbohydrates are essential for receptor folding and transport to the cell surface, but not for hormone binding [21]. The receptor is very sensitive to conformational changes, and even a conserved change in a single amino acid may cause its sequestration inside the cell, probably due to misfolding, thereby bringing about inactivation of FSHR action [22, 23]. With regard to the structural organization of the FSHR, there are the N- and C-terminal cysteine-rich regions of the extracellular domain, nine leucine-rich repeats between them, the transmembrane serpentine domain with seven transmembrane α-helixes joined by three intracellular and extracellular loops, and the intracellular tail. Upon functional characterization of the different domains of FSHR, it has been found that the N-terminal cysteine cluster is important for proper trafficking of the receptor to the cell membrane [3]. The middle leucine-rich repeats participate in ligand binding and determine its specificity [24].

D. FSHR Mutations and Their Functional Consequences Both inactivating and activating FSHR mutations are known in humans (Fig. 10.4). One inactivating mutation of FSHR has been found in five men, and it impairs spermatogenesis without causing azoospermia [15]. This mutation, a C566T transition, bringing about Ala189Val mutation, causes sequestration of the synthesized FSHR protein inside the cell, thereby blocking its entry to the site of action at the cell membrane [22]. Another inactivating mutation, Ala419Thr, was found in one woman as compound heterozygote with the Ala189Val mutation [25]. This mutation had minimal effect on ligand binding, but it totally abolished signal transduction as evidenced by absent cAMP generation upon stimulation of cells expressing the mutant receptor. Additional inactivating mutations of the FSHR include Pro348Ala and Pro519Thr substitutions, and they cause complete inactivation of receptor binding and signal transduction [26, 27]. The other inactivating FSHR mutations so far detected

−NH2

Ala189Val

Ile160Thr

(Asn191Ile ) )

Asp224Val

*

Val341Ala

*

Thr307Ala

Pro346Arg

Thr449Ile

Leu 601Val

Arg573Cys Ala419Thr Asp567Gly? Asp567Asn/ Gly

−COOH

*

Asn680Ser

FIGURE 10.4 Currently known mutations in the human FSHR. Schematic structures of the FSHR protein and localization of the inactivating (open squares) and activating (closed circles) mutations and polymorphisms (asterisks) are depicted. The short lines across the amino acid chain separate the 10 exons. The activating mutations Thr449Ile and Asp567Asn are of a special type in that they make the FSH receptor responsive to hCG. For further details, see the text.

concerned two compound heterozygous women with primary or secondary amenorrhea [28, 29]. Two of the mutations were detected in the extracellular domain, Ile160Thr and Asp224Val. They abolished signal transduction almost totally, and confocal microscopy demonstrated intracellular sequestration of the mutated receptor proteins. The other two mutations, Arg273Cys and Leu601Val, were localized in the transmembrane region, and in these cases signal transduction was partially compromised. Despite the rarity of the currently known inactivating FSHR mutations, a good correlation is seen between phenotype and degree of receptor inactivation, as well as the site of mutation and its functional consequences. All mutations in the extracellular domain cause defects in targeting of the receptor protein to the cell membrane. The mutations in the transmembrane region have minimal effects on ligand binding, and they impair to various extents, but not totally, the signal transduction. It also seems that the location, rather than the nature of amino acid substitution, determines the functional response. The only inactivating FSHR mutations in men so far detected have been

Chapter 10 FSH Regulation at the Molecular and Cellular Levels

in brothers of women with such mutations and hypergonadotropic hypogonadism (see earlier discussion). The reason may be that FSHR inactivation causes such a mild disturbance of Sertoli cell function that such men have at most mild phenotypes. One activating mutation in FSHR has been associated in a single male with maintenance of normal spermatogenesis in the absence of gonadotropin stimulation following hypophysectomy [30]. Because of the unusual nature of the subject (maintenance of some degree of testosterone production despite hypophysectomy) and the marginal constitutive activity of the mutated receptor in vitro, the final phenotypic importance of the mutation remains somewhat unclear. Very recently, two activating mutations in the FSHR were reported to make FSHR responsive to hCG and cause pregnancyassociated ovarian hyperstimulation syndrome [31, 32]. What is particularly intriguing in these mutations is that they can alter ligand specificity of the receptor. Whether similar mutations have phenotypic effects in men is not yet known.

III. MOLECULAR ASPECTS OF FSHR FUNCTION A. Activation and Signal Transduction of the FSHR As with LHR, the best characterized, and probably main signaling pathway of FSH action is the cAMPmediated activation of adenylyl cyclase and protein kinase A (PKA) [3, 18]. The first step after FSH binding to the receptor is its association with the Gs protein (Fig. 10.5). The Gs protein exists in inactive state as an α/β/γ heterotrimer binding GDP. Upon FSHR activation, it catalyzes the release of tightly bound GDP, permitting GTP to bind, which results in dissociation of the complex into Gsα-GTP and Gbγ. Gsα-GTP binds to and activates the plasma membrane associated adenylyl cyclase enzyme. The regulatory action is short lived due to the intrinsic GTPase activity of the α-subunit, which leads to reassociation of the heterotrimer into an inactive GDP associated complex. The Gβγ complexes are known to have other regulatory functions [33], but they have not been studied in connection with FSH action. The activated form of adenylyl cyclase catalyzes the conversion of ATP to cyclic adenosine-3’, 5’-monophosphate (cAMP), which is the best known second messenger of FSH action. It binds to the two regulatory subunits of the tetrameric PKA, liberating thereby the two catalytic subunits, which then phosphorylate

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structural proteins, enzymes, and transcription factors, altering in this way the functional state of target cells. FSH-dependent changes in phosphorylation of the cAMP responsive element (CRE) binding proteins (CREBs) and modulators (CREMs) belong to the latter category [34]. The CREM isoform ICER (inducible cAMP early repressor) is involved in downregulation of CREB expression [35]. FSHR activation is also linked to Gi protein activation, which may provide negative modulation of the PKA pathway [36]. The cAMP action is terminated by a group of phosphodiesterase enzymes that convert cAMP to adenosine monophosphate (AMP) [37]. Signaling mechanisms other than those related to cAMP generation have also been observed with FSHR activation in Sertoli cells, although their physiological significance remains under discussion. Protein kinase C (PKC) activity can be detected in Sertoli cells [38], but its coupling to FSH action is uncertain. Whereas FSHstimulated inositol phosphate turnover can be readily demonstrated in heterologous cell lines transfected with FSHR cDNA [39, 40], the results on activity of this pathway in primary Sertoli cells, with endogenous FSHR expression, have largely been negative [41, 42]. A recent study has demonstrated functionality of the FSH-stimulated phosphatidylinositol 3-kinase/protein kinase B signaling pathway in immature rat Sertoli cells [43]. The FSH-stimulated influx and increase in cytosolic free Ca2+ seem to be predominantly attributable to a serial signaling cascade after the generation of endogenous cAMP [44, 45]. Other signaling events that have been demonstrated in Sertoli cells in response to FSH are activation of phospholipase A2 [46], translocation of nuclear factor kappa B (NFKB) [47], activation of the mitogen-activated protein kinase (MAPK) pathway [48, 49], and membrane hyperpolarization with various ion fluxes [50].

B. Constitutive Activity of FSHR All receptors occur in two conformations: they are inactive in the absence of ligand and active when bound to the ligand. The role of ligand binding is to shift the balance of the receptor conformation from inactive to active. However, it seems that a small proportion of unliganded receptor can also be in the active conformation, as can be shown in cells transfected with FSHR. The basal cAMP production and steroidogenesis are higher in cells transfected with wild-type FSHR, in the absences of ligand, than in mock transfected cells, or those transfected with FSHR carrying an inactivating ligand (Fig. 10.6). This phenomenon may explain why FSHβ knockout mice

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FIGURE 10.5 Simplified schemes of association of the FSHR with transmembrane and intracellular effectors in (A) the resting stage, (B) upon acute signal transduction, and (C) upon attenuation of signal after the stimulatory phase. (A) In resting stage, with no FSH-receptor interaction, the Gs protein remains as an inactive α/β/γ trimer and the α-subunit binds GDP. Protein kinase A (PKA) is as inactive tetramer in which each catalytic subunit (C) is associated with one regulatory subunit (R). (B) Upon ligand binding, the α-subunit of Gs protein continued

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have a milder phenotype than FSHR knockout mice [51]. Conformational changes in the receptor protein through mutations may also alter its activity. Some mutations bring about constitutive activation of the unliganded receptor (i.e., activating or gain of function mutation), and constitutive inactivation (i.e., inactivating or loss of function mutation) [52]. In the former case, the unliganded receptor assumes a conformation that activates at least partly the signal transduction; in the latter, the conformational change hinders the activating conformation even in liganded state.

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C. Alternative Splicing of the FSHR Message

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FIGURE 10.6 Progesterone and cAMP production in a Leydig cell line (mLTC-1; to avoid confounding effect of endogenous FSHR expression) transfected with wild-type mouse FSHR (mFSHR-WT), mFSHR with an inactivating point mutation (mFSHR-C566T), or control vector. (A) Results for an experiment (mean ± SEM, quadruplicates) showing cAMP production by transfected cells under basal conditions and in the presence of increasing levels of hFSH. (B) Combined data for five experiments showing basal cAMP and progesterone (Prog) production in cells transfected with mFSHR-WT or vector alone, in the absence of FSH. (From [51], with permission.)

The FSHR message in Sertoli cells occurs in a number of splice variants, ranging in size from 1.3 to 6.5 bp [53–57]. A message of about 2.5 kb in size corresponds to the full-length FSHR protein. There seems to be no clear species specificity in alternate splicing, and neither have systematic differences been found between the ovary and testis, or between different functional or hormonal states of the gonads. The short messages apparently represent exon skipping, or usage of alternative splice donor or acceptor sites, with potentially unaltered reading frame, and translation of functional though truncated FSHR protein. The longer forms apparently represent usage of different polyadenylation sites, giving rise to longer 3’-untranslated regions [18]. Some of the alternatively spliced variants have been tested in connection with the full-length receptor in transfected cell lines, but they have not been found to modify the function of the full-length protein, which suggests that these forms have no major functional role [39, 58, 59]. In the case of LHR, one report detailed the enhancing effect of a truncated receptor on function of the full-length receptor [60]. The only truncated FSHR protein that has been shown to have biological function is that cloned from the sheep testis by the group of Sairam [61]. This receptor, termed FSH-R3, represents a growth factor type I–like receptor with a ligand binding domain and

dissociates from β/γ, binds GTP and activates adenylyl cyclase (AC), which catalyzes the conversion of ATP to the second messenger, cyclic adenosine monophosphate (cAMP). cAMP binds to the regulatory subunits of PKA, activating the dissociated catalytic subunits. They catalyze phosphorylation of target proteins, thereby bringing about the cellular responses to FSH stimulation. (C, a) Levels of the second messenger, cAMP, are decreased through activation of phosphodiesterase (PDE) through protein kinase A (PKA) stimulated phosphorylation. (C, b) G-protein–coupled receptor kinase (GRK) phosphorylate serine and threonine residues in the first and third intracellular loops of FSHR, which blocks coupling of Gs protein with the receptor, hence attenuating the signaling pathway. (C, c) The phosphorylated FSHR binds β-arrestin, which provides an additional hindrance to receptor-Gs coupling. The receptor–β-arrestin complex then binds to the clathrin-dynamin (D) endocytotic machinery and becomes internalized in endosomal vesicles. Consequently, the number of FSHRs on plasma membrane decreases, that is, becomes downregulated. The intracellular fate of the endocytosed receptor is either to become degraded or to enter the rapid recycling pathway.

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a single transmembrane domain, and it has arisen through splicing out of exons 9 and 10 of the wild-type FSHR, and inclusion of a novel C-terminal exon that encodes the single transmembrane domain. This receptor binds FSH, was found to have a dominant negative effect when cotransfected with wild-type receptor [62], and induced calcium-mediated signaling and MAPK activation [63, 64]. Recently, FSH-R3 has been found to be hormonally regulated in the developing mouse ovary, adding more credence to its physiological role [65].

D. Attenuation of FSHR Signaling through Receptor Desensitization and Downregulation The Sertoli cells have a mechanism to prevent FSH overstimulation that includes two parts: uncoupling of the second messenger signaling, or desensitization, and the loss of receptor sites, or downregulation (Fig. 10.5). In principle, similar mechanisms can be detected in the responses of all GPCRs. Multiple steps can be identified in this response, including (1) rapid internalization and sequestration of the ligand-bound receptor [66–68], (2) post-translational modification of the receptor [40, 66–69], (3) reduction of adenylyl cyclase activity [70], (4) increased phosphodiesterase activity [40, 66–69, 71, 72], (5) direct inhibition of PKA by specific inhibitors [73, 74], and (6) downregulation of the transcription, and consequent translation, of the receptor gene [75–77]. Because the levels of circulating FSH vary relatively little in the males of nonseasonally breeding species, the modulation of levels of FSHR and its signal transduction may be an important part in the physiological regulation of FSH action. The first response of Sertoli cells to prolonged FSH stimulation, independent of decreased receptor synthesis and internalization, is desensitization or uncoupling, which means a reduction in the ability of the receptor to activate second messenger systems. Time-wise, desensitization precedes receptor downregulation, and it is believed to occur through post-translational modifications of the receptor [78, 79]. A similar desensitization mechanism is employed by all GPCRs and it involves phosphorylation of serine or threonine residues in the intracellular loops or the cytoplasmic tail of the receptor, which disrupts the capability of the receptor to couple with Gs [79, 80]. In FSHR, the phosphorylation occurs in the first and third intracellular loops of the receptor [81]. At low agonist concentration, PKA catalyzes the phosphorylation, and at high concentration specific G-protein–coupled receptor kinases (GRKs) catalyze the reaction. Although not

studied with all receptor systems, the removal of phosphorylation sites results in a decrease in agonist-induced desensitization [82–84]. Receptor phosphorylation also promotes the interaction of the receptor with β-arrestin [85], which provides another hindrance for the receptor– Gs protein interaction. The gonadotropin-induced uncoupling thus entails functional changes in the receptor rather than in the Gs protein or adenylyl cyclase. From experiments using inhibition of PKA and PKC activities in cultured cells, Ascoli [79] concluded that the FSH-induced phosphorylation of rat FSHR is not mediated by PKA, and PKC appears to have minimal effect. However, there are also studies emphasizing the role of PKC in FSHR phosphorylation upon desensitization [86], but the physiological significance of these findings remains unclear. Recent findings have shown that the GRK/arrestin system has a pivotal role in these responses [87, 88]. Primary rat Sertoli cells express GRK2, -3, 5, -6α, and -6β and β-arrestins 1 and 2 [89]. The findings that inhibitors of GRK2 and GRK6 impair phosphorylation, but only the inhibitors of GRK2 impair internalization, suggest that different GRKs have differential effects on receptor desensitization and internalization [87], but details of these differences are not yet known. The second phase of the FSH target cell responses to prolonged hormonal stimulation is homologous downregulation. A high concentration of FSH results in a 90% decrease within 4 hr of the steady-state levels of FSHR mRNA in cultured Sertoli cells, with a parallel loss of FSHR binding [77]. These data suggest that the downregulation was due to cAMP-mediated RNA degradation. Subsequent data of Maguire et al. [75] suggested a role for transcription, rather than mRNA stability, in this phenomenon. A very different model for the FSHR downregulation was presented by the study of Monaco et al. [76], where a role of altered FSHR de novo transcription and translation was proposed. These authors observed that in cultured primary Sertoli cells, after FSH stimulation, a dramatic increase in the CREM isoform ICER occurred. The downregulation and desensitization effects are brought about by binding of ICER to a CRE sequence in the FSHR promoter. The latest data on this controversial topic demonstrate that the first phase in the response of cultured Sertoli cells to treatment with FSH is a reduction in the amount of heteronuclear (hn) RNA for FSHR, which reflects a decrease in the level of transcription of this gene [90]. However, no effects on FSHR promoter activity, or a role for ICER, were observed in these conditions, and it was concluded from experiments modulating histone deacetylation that the downregulation may be mediated by changes in chromatin structure. Hence, despite

Chapter 10 FSH Regulation at the Molecular and Cellular Levels

several clues the final mechanism of the homologous FSHR downregulation remains elusive. Another phase of FSHR downregulation is independent of alterations in FSHR transcription and it follows the responses initiated by PKA or GRK catalyzed FSHR phosphorylation (Fig. 10.5). A large body of evidence suggests that this phenomenon, common for GPCRs, occurs through coupling of the receptor to nonvisual arrestins (β-arrestin) [91]. The formation of this complex is facilitated by agonist-induced activation and/or by the GRK-catalyzed phosphorylation of the GPCRs. Ligand binding to FSHR promotes rapid internalization of the hormone–receptor complexes. The internalization occurs through clathrin-coated pits via a dynamin-dependent pathway [85, 92]. The internalized endocytotic vesicles containing FSHR are destined when inside the cell either to degrade or to be recycled back to the plasma membrane, which replenishes the functional state of the cell to respond to FSH stimulation. All details of the cascade described have not been explored with respect to FSHR and Sertoli cell function, but it is assumed that the principles of these responses are similar to those of other GPCRs.

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IV. MODIFICATION OF FSH ACTION IN SERTOLI CELLS BY SPERMATOGENIC CELLS FSH action in Sertoli cells is modified by the surrounding spermatogenic cells [93]. Spermatogenic cells develop synchronously in cell associations that follow each other in a wavelike fashion in the seminiferous epithelium. Sertoli cells form the structural and functional framework for these cell associations, also called stages, and each Sertoli cell harbors different stages in a cyclic manner. Several FSH-dependent functions are stage dependent.

A. FSHR Expression and FSH Binding Steady-state levels of FSHR mRNA vary in different stages of the rat seminiferous epithelial cycle [94]. The highest levels are found in stages XIII through II, and low levels are detected in stages VII and VIII (Fig. 10.7). FSH binding in the seminiferous tubules shows the same pattern [95]. The binding reflected receptor numbers rather than binding affinity [95]. FSH binding was altered by modification of cell

FIGURE 10.7 Stage-specific pattern of the expression of FSHR mRNA (■——■; on the basis of [94]), FSHstimulated cAMP production (■——■; on the basis of [96]) and the expression of FSH-stimulated inhibin α mRNA (•——•; on the basis of [103]) in the rat seminiferous epithelium. The curves are based on original arbitrary values that show relative amounts.

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associations in irradiated rat testes. Absence of differentiating spermatogonia and early primary spermatocytes was associated with decreased FSH binding, and the stage specificity disappeared when all primary spermatocytes were abolished [95]. These results suggested that spermatogenic cells stimulate FSH binding in Sertoli cells in a stage-specific manner.

B. FSH-Stimulated cAMP Production Sertoli cells from the adult animals respond poorly to FSH compared to the cells from young animals. However, the cAMP response is also stage dependent and follows the same pattern as FSHR content and FSH binding in the seminiferous tubules. Both basal and FSH-stimulated cAMP production are low in stages VII and VIII and subsequent stages IX through XI, whereas increasing response is found in stages XIII through I and the highest cAMP production occurs in stages II through V, that is, somewhat later than the highest FSH binding [96]. Modulation of cAMP responses by spermatogenic cells can also been seen in testes that are missing defined germ cells. The responses tend to increase particularly in stages VII through VIII in the absence of different spermatogenic cells [97]. In addition to cAMP production, CREB is also stage dependent in rat Sertoli cells [98]. CREB mRNA levels are high at stages III through VI and low in other stages, resembling the pattern of cyclic FSH-stimulated cAMP production. CREB protein expression seems to be low in Sertoli cells, whereas haploid spermatids show strong expression in stages VI through VIII, that is, after the high mRNA expression in Sertoli cells. Sertoli cell responses to FSH are dominated by cAMP-mediated activation of PKA and the downstream cascade. However, during Sertoli cell proliferation in young rats, FSH also activates the ERK/MAP kinase pathway through coupling of the FSHR both to Gs and Gi heterotrimeric proteins in a PKA- and Src-dependent manner [48]. Although this pathway is activated in proliferating Sertoli cells, FSH inhibits it in Sertoli cells that have stopped dividing and undergo differentiation.

C. FSH-Stimulated Protein Synthesis FSH stimulates protein synthesis in Sertoli cells by increasing transcription and mRNA stability. Although the majority of proteins are stimulated, some genes are also repressed by FSH [99]. An oligonucleotide microarray analysis of gene expression in FSH-stimulated rat Sertoli cells [99] identified

100 to 300 transcripts that were either upregulated or downregulated at least twofold. Among these were several proteins that were previously known to be regulated by FSH. Interestingly, many of these proteins are also expressed in a stage-dependent fashion in the seminiferous tubules. Some of the proteins are discussed briefly next. 1. Inhibin Inhibins are dimeric glycoproteins composed of two subunits, one of which is α and the other either βA (inhibin A) or βB (inhibin B) [100]. Both inhibins inhibit pituitary FSH secretion. Sertoli cells produce mainly α and βB, and therefore inhibin B is the main circulating inhibin hormone in serum [100]. Both α and βB mRNAs are cyclically expressed in the seminiferous epithelium (Fig. 10.7) [101, 102]. The highest mRNA levels are found in stages XIII through IV and the lowest in stages VII through VIII [101, 102]. FSH stimulates expression of inhibin α mRNA and secretion of inhibin B [100]. Interestingly, the accurate stage-specific expression pattern of inhibin α mRNA in the seminiferous epithelium follows closely the FSH-stimulated cAMP secretion [103, 104]. Furthermore, the absence of pachytene spermatocytes and round spermatids at stages VII and VIII is associated with a many-fold increase in inhibin α mRNA expression [103]. Inhibin secretion is also increased when pachytene spermatocytes are missing from seminiferous tubules [105]. In vitro studies with cocultures of spermatogenic cells and Sertoli cells have given mixed results. Pachytene spermatocytes were reported to inhibit the expression of βB mRNA and secretion of inhibin B, whereas expression inhibin α was unaffected and spermatids had no effects [106]. In another study spermatids had a stimulatory effect on inhibin α expression [107]. Isolation and culture of spermatogenic cells is complicated and may result in great differences in experimental results. Nevertheless, both in vivo and in vitro findings imply that spermatogenic cells strongly modulate the Sertoli cell responses to FSH. Inhibin may have a paracrine role in the regulation of spermatogenesis in addition to its endocrine function. 2. Testibumin and α2-Macroglobulin Several Sertoli cell products are differentially regulated by FSH, depending on the stage of the seminiferous epithelial cycle. FSH elevates testibumin levels in all stages by about 40%, whereas α2-macroglobulin secretion is stimulated by FSH in stages XIII through I more than in other stages [108]. Basal testibumin and α2-macroglobulin levels are highest in stages II

Chapter 10 FSH Regulation at the Molecular and Cellular Levels

through VI and II through VIII, respectively. α2-Macroglobulin is a nonspecific protease inhibitor that also binds several growth factors and thereby modify their action. Testibumin shares some features with albumin and α-fetoprotein [109]. The specific functions of these proteins in the seminiferous epithelium are not known, but their different regulation by FSH demonstrates the importance of local interactions. 3. Stem Cell Factor Stem cell factor (SCF) is the ligand of c-kit, a tyrosine kinase receptor that is expressed both on germ cells and Leydig cells. The SCF/c-kit system is involved in the development of testes and regulation of spermatogenesis as an important survival factor [110–113]. Sertoli cells produce SCF under FSH regulation [114]. FSH action is mediated through the cAMP/PKA signaling cascade, whereas testosterone and many growth factors have no effects on SCF expression [115, 116]. SCF expression is also stage dependent in the seminiferous epithelium, and the basal expression pattern is similar to that of inhibin α [117]. FSH stimulates transcription of SCF mRNA in all stages, but the stimulation is strongest in stages II through VI and weakest in stages VII and VIII, following closely the pattern of FSHR expression and binding [116]. SCF protects germ cells from apoptosis in vitro [113, 117]. One of the main effects of FSH in the seminiferous epithelium is the stimulation of Sertoli cell and spermatogonial proliferation and inhibition of cell death [113, 118]. When the SCF/c-kit system is blocked by receptor antibodies ACK-2, apoptosis of spermatogonia, spermatocytes, and spermatids is increased and spermatogonial proliferation is inhibited [113, 119, 120]. ACK-2 does not only block the cell survival effect of SCF, but also that of FSH to a large extent, suggesting that this effect of FSH is mainly mediated through SCF/c-kit [113]. FSH and SCF influence the apoptotic machinery by regulating the Bcl-2 family of proteins in the testis [121, 122]. SCF is produced in soluble and membranebound forms in Sertoli cells. The soluble form is involved in the proliferation of Leydig cells [123]. Therefore, FSH can also influence Leydig cell development via Sertoli cells. 4. Lactate Dehydrogenase A Meiotic and postmeiotic spermatogenic cells, that is, spermatocytes and spermatids, depend on Sertoli cells in their energy metabolism because they cannot use glucose as a substrate. They instead prefer lactate or pyruvate [124–126]. Sertoli cells provide lactate for

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spermatogenic cells by the action of lactate dehydrogenase A (LDHA), which is regulated by FSH [127, 128]. FSH upregulates LDHA mRNA by increasing its stability [129]. Lactate is transported to germ cells by specific proton/monocarboxylate transporters [130] that are expressed in spermatogenic cells [128], and lactate is converted into pyruvate in germ cells by lactate dehydrogenase C [131]. Lactate is important for the survival of the germ cells [124–126], and it inhibits apoptosis in the human testis [132]. Spermatogenic cells influence LDHA activity in the Sertoli cells by secreting growth factors and cytokines, such as TNFα, that stimulate LDHA mRNA transcription [129]. Therefore, the Sertoli cells and germ cells can cooperate effectively to guarantee the optimal energy balance for developing spermatogenic cells. Although great advancements in the elucidation of the molecular mechanisms of FSH action in Sertoli cells have been made during the last decade, more detailed analyses of the specific effects in the seminiferous tubules may provide tools to modulate FSH action to maintain fertility and control spermatogenesis in contraceptive manner.

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119. Yoshinaga, K., Nishikawa, S., Ogawa, M., Hayashi, S., Kunisada, T., and Fujimoto, T. (1991). Role of c-kit in mouse spermatogenesis: identification of spermatogonia as a specific site of c-kit expression and function. Development 113, 689–699. 120. Tajima, Y., Nishina, Y., Koshimizu, U., Jippo, T., Kitamura, Y., and Nishimune, Y. (1993). Effects of hormones, cyclic AMP analogues and growth factors on steel factor (SF) production in mouse Sertoli cell cultures. J. Reprod. Fertil. 99, 571–575. 121. Yan, W., Suominen, J., Samson, M., Jegou, B., and Toppari, J. (2000). Involvement of Bcl-2 family proteins in germ cell apoptosis during testicular development in the rat and prosurvival effect of stem cell factor on germ cells in vitro. Mol. Cell Endocrinol. 165, 115–129. 122. Yan, W., Samson, M., Jegou, B., and Toppari, J. (2000). Bcl-w forms complexes with Bax and Bak, and elevated ratios of Bax/Bcl-w and Bak/Bcl-w correspond to spermatogonial and spermatocyte apoptosis in the testis. Mol. Endocrinol. 14, 682–699. 123. Yan, W., Kero, J., Huhtaniemi, I., and Toppari, J. (2000). Stem cell factor functions as a survival factor for mature Leydig cells and a growth factor for precursor Leydig cells after ethylene dimethane sulfonate treatment: implication of a role of the stem cell factor/c-Kit system in Leydig cell development. Dev. Biol. 227, 169–182. 124. Jutte, N. H., Grootegoed, J. A., Rommerts, F. F., and van der Molen, H. J. (1981). Exogenous lactate is essential for metabolic activities in isolated rat spermatocytes and spermatids. J. Reprod. Fertil. 62, 399–405. 125. Jutte, N. H., Jansen, R., Grootegoed, J. A., Rommerts, F. F., Clausen, O. P., and van der Molen, H. J. (1982). Regulation of survival of rat pachytene spermatocytes by lactate supply from Sertoli cells. J. Reprod. Fertil. 65, 431–438. 126. Mita, M., and Hall, P. F. (1982). Metabolism of round spermatids from rats: lactate as the preferred substrate. Biol. Reprod. 26, 445–455. 127. Mita, M., Price, J. M., and Hall, P. F. (1982). Stimulation by follicle-stimulating hormone of synthesis of lactate by Sertoli cells from rat testis. Endocrinology 110, 1535–1541. 128. Goddard, I., Florin, A., Mauduit, C., Tabone, E., Contard, P., Bars, R., Chuzel, F., and Benahmed, M. (2003). Alteration of lactate production and transport in the adult rat testis exposed in utero to flutamide. Mol. Cell Endocrinol. 206, 137–146. 129. Boussouar, F., Grataroli, R., Ji, J., and Benahmed, M. (1999). Tumor necrosis factor-alpha stimulates lactate dehydrogenase A expression in porcine cultured Sertoli cells: mechanisms of action. Endocrinology 140, 3054–3062. 130. Halestrap, A. P., and Price, N. T. (1999). The proton-linked monocarboxylate transporter (MCT) family: structure, function and regulation. Biochem. J. 343 Pt 2, 281–299. 131. Li, S. S., O’Brien, D. A., Hou, E. W., Versola, J., Rockett, D. L., and Eddy, E. M. (1989). Differential activity and synthesis of lactate dehydrogenase isozymes A (muscle), B (heart), and C (testis) in mouse spermatogenic cells. Biol. Reprod. 40, 173–180. 132. Erkkila, K., Aito, H., Aalto, K., Pentikainen, V., and Dunkel, L. (2002). Lactate inhibits germ cell apoptosis in the human testis. Mol. Hum. Reprod. 8, 109–117.

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C H A P T E R

11 In Vivo FSH Actions CHARLES M. ALLAN AND DAVID J. HANDELSMAN ANZAC Research Institute, Sydney, NSW 2139 Australia

I. INTRODUCTION II. ANIMAL MODELS TO STUDY IN VIVO FSH ACTIONS: NONSEASONAL BREEDERS III. ANIMAL MODELS TO STUDY IN VIVO FSH ACTIONS: SEASONAL BREEDERS IV. NONHUMAN PRIMATES V. HUMANS VI. THE BALANCE BETWEEN FSH AND STEROIDS: THE E-T SWITCH References

in vivo models for FSH and Sertoli cell function remain indispensable. Historically, the role of FSH in regulating spermatogenesis via effects on Sertoli cell function has been increasingly appreciated during the last century. Following Sertoli’s morphological identification of the somatic support cells for the germinal epithelium, at least five major landmarks have occurred in the progressive acquisition of knowledge about the hormonal control of spermatogenesis. The first was the development in the late 19th and early 20th century of hypophysectomy, especially by Phillip Smith whose technique facilitated experimental research proving the centrality of pituitary control of testicular function including spermatogenesis [2]. A second was the recognition by Herbert Evans and Roy Greep, among others, that the pituitary produced two distinct gonadotrophic hormones to regulate gonadal function. The third landmark was the development of radioimmunoassay by Berson and Yalow allowing accurate measurement of circulating hormones. The fourth was the identification of the hypothalamic gonadotrophin-releasing hormone (GnRH) by Guillemin and Schally together with the recognition of its pulsatile secretion by Knobil. Finally, the molecular biology era has allowed researchers to experience profound new insights into the genes and proteins involved in the hormonal regulation of Sertoli function. During the past decade, the use of customized, genetically modified rodent models, with the targeted addition or deletion of selected hormones, receptors, or potential downstream mediators of hormonal responses, has created a novel window into this physiological complexity by allowing more specific dissection of the in vivo FSH response. These more sophisticated

I. INTRODUCTION Follicle-stimulating hormone (FSH) plays a major role in both the development and mature function of Sertoli cells. The testicular actions of this pituitaryderived heterodimeric glycoprotein hormone are mediated via a specific plasma membrane FSH receptor exclusively expressed on Sertoli cells (reviewed in [1]). Although the FSH response is initiated by a well-characterized receptor, any in vivo examination of FSH activity has to contend with a complex microenvironment of different interacting cell types (germ and somatic) and biological pathways that influence Sertoli cell function, including multilevel regulation by both local and extratesticular hormones comprising the hypothalamic–pituitary–testicular axis. A major challenge provided by this physiological complexity has been to isolate FSH activity from the other closely related pituitary gonadotrophin, luteinizing hormone (LH), which activates receptors on interstitial Leydig cells to stimulate testicular testosterone production. Because the accurate replication of these complex tissue and cellular interactions is not yet feasible, in vitro and SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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animal models have produced a fundamental shift in the prevailing view held until the mid-1990s that FSH was essential for the initiation of spermatogenesis [3]. Several mouse models clearly demonstrated that FSH is not required to initiate Sertoli cell differentiation or support the qualitative completion of spermatogenesis, nor is it essential for male fertility. The extension of this finding to primate and human physiology remains incomplete and controversial, largely due to limited opportunities for decisive investigation in primates. This has left controversy over an essential function for FSH in the human testis. However, in all mammalian models examined to date, it is clear that FSH plays a pivotal role in the establishment and support of full Sertoli cell function, a prerequisite to induce and maintain the optimal spermatogenic capacity of the testis. This chapter reviews the paradigms used to evaluate in vivo FSH actions in the mammalian testis and highlights key findings from the past decade.

II. ANIMAL MODELS TO STUDY IN VIVO FSH ACTIONS: NONSEASONAL BREEDERS A. Rat Most present knowledge of in vivo FSH actions on the testis is attributable to studies of the rat [4, 5]. During male rat development, circulating levels of FSH are detectable toward the end of fetal life ~19 days postcoitus (dpc) and rise until birth [6]. After birth, FSH levels first decline 50–70% and then rapidly rise two- to threefold during postnatal life, after which FSH levels steadily decline with age [6–8]. The rat FSH receptor appears earlier in male development than its cognate ligand. FSH receptor mRNA is detectable in the testis 16.5 dpc [9], consistent with the appearance of testicular FSH binding sites at 17.5 dpc [10]. Therefore, the manifestation of FSH activity occurs after the development of early Sertoli and germ (gonocyte) cells and formation of seminiferous chords (between 14.5 and 15.5 dpc) [11], but coincides with the crucial perinatal period of Sertoli cell proliferation. Early 3H-thymidine incorporation studies established that rat Sertoli cells only proliferate from ~16 dpc to about 15 days after birth, beyond which further mitotic activity is hard to detect [12, 13] although low levels of ongoing replication may still occur. The in utero injection of fetal rats (18 dpc) with anti-FSH serum halved the percentage of 3H-thymidine–labeled Sertoli cells the following day [14]. In contrast, elevated FSH levels after the hemicastration of neonatal or immature rats was associated with Sertoli cell proliferation (final

numbers increased above normal) and testicular hypertrophy [15–18], whereas in mature rats serum FSH concentrations also increase after hemicastration but testis size does not [15, 17]. The magnitude of perinatal FSH effects was shown by the direct injection of an antimitotic agent (cytosine arabinoside) to newborn rat testes, which ultimately halved the mature populations of both Sertoli cells and spermatids [19]. These findings demonstrated that FSH is a key Sertoli cell mitogen during the perinatal period, thereby making it a critical determinant for the final Sertoli cell number and presumably therefore the ultimate germ cell carrying capacity of the testis. Recent technical advances have led to improved investigational insights. For example, the specific testicular effects attributable to FSH in these early rat studies were complicated by impure FSH preparations, because contributing effects due to stimulation of Leydig cell testosterone production by LH contamination could not be ruled out [20, 21]. More incisive analysis of the FSH response was provided by the availability of LH-free recombinant human FSH (rhFSH) [22]. Furthermore, unbiased statistically valid quantitative techniques have been developed to enumerate Sertoli and germ cell populations using modern stereological techniques [23]. By providing more accurate quantitative estimates of the cellular populations of the testis, this has facilitated more insightful investigations of the hormonal regulation of Sertoli cell number and function, including the development and maintenance of germ cell populations that require the support of differentiated Sertoli cells. For instance, neonatal exposure of normal rats to supraphysiological levels of rhFSH caused testicular hyperplasia, with stereological analysis demonstrating that Sertoli and spermatid populations were proportionately elevated above normal levels [24]. A key finding from this study was that the extra Sertoli cell proliferation was stimulated by excessive FSH treatment without prolonging the normal duration of postnatal Sertoli cell division (i.e., up to 15 days of age). In more recent work, postnatal rhFSH treatment of 5- to 15-day-old rats doubled Sertoli and early spermatogonia cell numbers compared with controls [25]. Previous studies showed that neonatal estradiol treatment alone decreased circulating FSH during postnatal life [26–28], suggesting that perinatal effects of excessive estradiol may be due to inhibition of pituitary FSH secretion. Overall, these findings supported the view that (1) there is a limited time frame for the postnatal mitogenic activity of FSH and (2) the final germ cell capacity of the testis depends on the FSHinduced postnatal proliferation of Sertoli cells. Exogenous neonatal FSH treatment in the preceding studies was superimposed on normal endogenous

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pituitary FSH and LH secretion, which continue to act on the testis. For example, FSH- and LH-stimulated testosterone (or estradiol) can act simultaneously on FSH and androgen (or estradiol) receptors found in Sertoli cells, making it difficult to separate FSH and LH activities in the presence of continuing pituitary gonadotrophin secretion. To isolate the FSH response, investigators have studied FSH loss and/or replacement (LH-free recombinant or highly purified FSH) in rats following hypophysectomy [29–35], or the administration of GnRH antagonists [36–40] or immunoneutralizing antibodies [14, 41–44]. These classic approaches were performed primarily on immature rats after the perinatal Sertoli cell proliferation period, so the findings are most applicable to FSH actions in maintaining or restoring Sertoli cell function rather than initiating Sertoli cell proliferation. In mature rats, short-term (1-week) rhFSH treatment at 3 [32] or 6 days [35] after hypophysectomy reduced germ cell degeneration following complete pituitary ablation. FSH partially maintained mitotic, meiotic, and postmeiotic germ cell stages, without any effect on testicular androgen production [32]. Nevertheless, administration of flutamide (a nonsteroidal androgen receptor antagonist) further reduced germ cell degeneration, suggesting that residual androgen action after hypophysectomy also contributed to maintaining germ cell survival. A limitation on the use of hypophysectomy to study gonadotrophin effects on Sertoli cells is the nonselective removal of all other pituitary hormones, including growth hormone [45], prolactin [46], and thyroid-stimulating hormone [47–50], which also affect testicular development and Sertoli cell numbers. A more selective suppression of gonadotrophin activity can be achieved using pharmacological GnRH antagonists, although this may be less complete especially for FSH. Neonatal GnRH-antagonist (GnRH-A) treatment produced a permanent reduction (~50%) of Sertoli cell numbers in mature rats [37, 51]. In contrast, GnRH-A administration in mature rats resulted in germ cell regression similar to that after hypophysectomy, with no change in Sertoli cell number [39]. This supports the view that gonadotrophin-dependent Sertoli cell proliferation is restricted to the perinatal period. rhFSH also partially maintains mitotic (SgB) and meiotic (preleptotene) germ cells after 4 weeks of gonadotropin deprivation using a GnRH-A, whereas rhFSH did not prevent regression of later stage pachytene spermatocytes and spermatids, nor was Leydig cell steroidogenesis altered [39], confirming observations from classical ablate–replace models using hypophysectomized rats [29, 32]. Thus, mature rat Sertoli cells stimulated by FSH alone can partially support early germ cell development and meiosis, but not later stage postmeiotic germ cell populations.

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Comparable findings were obtained studying the restorative effects of rhFSH (1- to 3-week treatments) in rats immunized against GnRH [52]. In mature GnRH-immunized rats with severe spermatogenic regression, rhFSH partially restored spermatogonia, meiotic spermatocytes, and early spermatids, but not later elongated spermatid numbers. Overall, hormone replacement studies in rats show that FSH alone cannot fully maintain or restore postmeiotic spermatogenesis, indicating that mature Sertoli cell function requires additional factors, notably testosterone, for optimal function. The later study also highlighted inadvertent immunoneutralization as an important limitation of longer term rhFSH administration in rodents, because 66% of all animals treated for 3 or 4 weeks developed detectable antibodies to rhFSH [52]. This is particularly limiting given that the full rat spermatogenic cycle occupies more than 50 days. Although antibodies to exogenous FSH may confound interpretation by active immunization, anti-FSH antibodies have also been exploited in a complementary passive immunoneutralization approach to study Sertoli cell function. For example, the short-term administration (under 9 days) of antiserum raised to FSH increased apoptosis [43] and decreased the absolute numbers [44] of spermatogonia and spermatocytes in adult rats. Immunization of female rats with a peptide representing a unique, conserved extracellular domain of the FSH receptor during the last week of pregnancy and lactation resulted in male offspring with more aspermatogenic tubules, presumably due to passive transfer of maternal antibodies across the placenta and/or via colostrum [53]. In addition, the direct immunization of immature (18-day-old) or adult (80-day-old) male rats to this receptor peptide reduced sperm numbers per gram weight of testis by ~50 and 15%, respectively [53]. This suggests that immature Sertoli cells (after completion of replication but before the establishment of full spermatogenesis) are more vulnerable to lost FSH activity than are mature Sertoli cells already supporting complete spermatogenesis. The drawbacks of immunoneutralization include the assumption of specificity, absence of bystander autoimmune Sertoli cell damage, and uncertain degree of FSH blockade. For example, sera from immunized rats only blocked FSH binding to testis membranes by 5–40% [53]. In summary, several distinct rat models have shown that FSH plays a key role in the perinatal proliferation of Sertoli cells and early germ cells. In mature rats, FSH promoted spermatogonial and meiotic germ cell development, but in the absence of LH-stimulated testosterone production, FSH only stimulated a limited and incomplete level of postmeiotic spermiogenesis. Although these rat models have provided the foundation for understanding FSH regulation of

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Sertoli cell proliferation and function, several drawbacks in these approaches include the difficulty of obtaining complete and sustained FSH suppression by surgical, pharmacological, or antibody-mediated blockade of GnRH or FSH activity, as well as temporal limitations investigating the selective perinatal or longer term consequences of the FSH response. As described next, the development of mouse models with specific genetic modifications has circumvented many of these limitations.

B. Mouse Sertoli cell replication in the mouse follows a pattern similar to that of the rat, being restricted to the late fetal and early postnatal period. Early 3H-thymidine labeling studies showed that the percentage of proliferating Sertoli cells was highest during perinatal development (from 16 dpc to ~5 days old), then rapidly declined to undetectable levels 12–17 days after birth [54, 55]. These findings were consistent with direct stereological enumeration of total Sertoli cell numbers in normal mouse testes, which reached maximal levels in postnatal males ~18 days old [56, 57]. In comparison, the characterization of the developmental appearance of FSH activity remains incomplete for the mouse testis. For example, ontogeny of the mouse FSH receptor has primarily focused on the ovary with little known of the testis [58]. Furthermore, the application of classical approaches to study FSH in vivo activity has been more restricted in the mouse. Hypophysectomy was limited by the technical difficulty of smaller scale surgery, and pharmacological suppression of gonadotrophin secretion by GnRH analogues was less effective in mice compared to rats [59], requiring bulky sustained-release delivery devices [60]. However, during the last decade, investigation of FSH regulation of Sertoli cell biology in vivo was significantly advanced by mouse models with well-characterized or targeted genetic alterations. In particular, transgenic technology has expanded the use of mice as an alternative rodent model to the rat, primarily because transgenic procedures are easier and more economical for mice, a species with more extensive genome information available. Selective germline disruption or gene overexpression in mice has allowed a detailed definitive analysis of the selective loss or gain of in vivo FSH activity. 1. Loss of FSH Activity The genetic ablation of FSH in mice has provided an effective, albeit indirect, approach to the investigation of its role in Sertoli cell development. For example, the

gonadotrophin-deficient hypogonadal (hpg) mouse has provided a valuable model to selectively study postnatal FSH deficiency. The hpg mice carry a 33.5-kb deletion in the GnRH gene [61] making them functionally deficient in circulating FSH, LH, and androgens [62–64]. These mice retain all other pituitary hormones and remain responsive to gonadotrophin and androgen. Confirmation that the hpg phenotype was solely due to the GnRH gene defect was provided by the rescue of normal testicular development and fertility by hypothalamic transplantation [65] and transgenic GnRH [66]. The hpg testis remains immature with males continuing to exhibit Sertoli cells with undifferentiated immature-like morphological characteristics [67, 68], such as the presence of irregularly shaped nuclei lacking the basally located, tripartite nuclear structure found in mature Sertoli cells [55, 69]. The immature Sertoli cells in hpg mice maintain spermatogenesis arrested at the pachytene stage of meiosis [62, 63]. Because this replicates the spermatogenic development of tfm mice with nonfunctional androgen receptors, this indicates that the hpg represents a model of complete postnatal functional androgen deficiency [70] albeit with preserved androgen sensitivity. Consequently, the dormant immature hpg testes have provided a valuable experimental platform on which reproductive hormones at any level of the hypothalamic–pituitary–gonadal axis can be evaluated, such as GnRH [66], FSH or LH/hCG [67, 68, 71–73], or steroids [63, 74, 75]. The evaluation of FSH-deficient hpg mice receiving testosterone treatment alone revealed the presence of qualitatively complete spermatogenesis and restoration of male fertility [63, 67, 76], demonstrating that Sertoli cells functionally differentiate in the absence of FSH, contrary to prior prevailing consensus [3]. However, mature testes of hCG or testosterone-treated and FSHdeficient hpg males remain small (30–50%) and Sertoli cell numbers ∼ 50% of normal, highlighting the primary role of FSH in stimulating sufficient Sertoli cell replication to support the full germ cell capacity of the testis [63, 67, 68]. In hpg males, dihydrotestosterone (a potent nonaromatizable androgen) alone initiated qualitatively complete spermatogenesis on a background functionally deficient in FSH and LH [63]. Surprisingly, another study suggested that estradiol alone can stimulate spermatogenesis in hpg mice [75], perhaps due in part to the pharmacological stimulation of pituitary FSH secretion. Other gonadotrophin-deficient mouse models replicating the hpg phenotype have been produced by the targeted ablation of pituitary gonadotrophs. A toxic transgene approach was developed using glycoprotein alpha-subunit gene promoter directing expression of diphtheria toxin A chain to gonadotrope but not thyrotropes, thus eliminating gonadotrophins while

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TSH and thyroid function remained normal [77]. Transgenic males were infertile and exhibited immature gonads similar to hpg mice. An analogous inducible ablation strategy used a transgenic FSHβ subunit gene promoter to target thymidine kinase (TK) expression to gonadotrophs [78]. The cytotoxic ablation of transgenic TK+ gonadotrophs was induced by ganciclovir, which generated hypogonadal mice. However, fetal gonadal development was normal, suggesting that FSH and LH (and TSH) are not required for sexual differentiation and genital development. In this model, plasma FSH levels were not completely suppressed, possibly due to mosaic TK expression in gonadotrophs. Furthermore, transgenic TK was detected in the postnatal testis, which could confound interpretation with this model due to direct testicular toxicity. In comparison, complete germline deletion of the α-subunit by homologous recombination produced hypogonadal animals totally deficient in FSH and LH, as well as TSH, resulting in hypothyroid dwarfism as well as hpg-like testes [79]. These models have in common the concurrent loss of LH and/or TSH, which complicates the interpretation and isolation of FSH action alone. More selective “loss-of-function” mouse models disrupting the FSH pathway have been produced by homologous recombination in embryonic stem cells. Targeted deletion of either the FSH beta-subunit [80] or FSH receptor genes [81, 82] produced knockout models with qualitatively complete spermatogenesis, which confirmed the earlier proposal that FSH is not essential for mouse Sertoli cell function or male fertility [63]. The genetic loss of the FSH receptor or its circulating FSH ligand produced apparent differences in testicular phenotype. First, total Sertoli cell numbers were ~70% of normal in FSHβ null mice [83], but only ~50% of normal in FSHR null (FSHR–/–) mice [84]. Secondly, FSHR–/– models exhibited lower than normal intratesticular testosterone levels [82, 85], compared to normal circulating and intratesticular testosterone levels found in FSHβ knockout mice [80, 85]. Therefore, removal of the FSH receptor caused more disruption to Sertoli cell development and testicular steroidogenesis compared to the selective loss of FSH ligand. Leydig cell numbers are also reduced in FSHR–/– males, but normal in FSHβ null mice [85]. It was suggested that a diminished Leydig population and ensuing reduction in androgen production were due to the loss of constitutive FSH receptor activity in FSHR–/– testes [85]. Thus, concerted analysis of these loss-of-function mouse models suggested that the presence of the FSH receptor promotes Leydig cell function, which is in agreement with the abundant in vitro evidence for FSHmediated paracrine communication between Sertoli and Leydig cells (reviewed in [86] and Chapter 18).

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Conversely, it could be that lower intratesticular testosterone levels contribute to reduced Sertoli cell proliferation (by ~20%) in FSHR–/– compared to FSHβ–/– males. It is possible that higher (normal) postnatal testosterone levels in the FSHβ null testes, relative to FSHR–/– animals, stimulated the additional Sertoli cell proliferation [83]. The relative differences in Sertoli cell numbers suggest that normal postnatal androgen activity accounts for ~20% of the Sertoli cell population. Supporting this proposal, the increase of total Sertoli cell numbers following the administration of testosterone or hCG to FSH-deficient hpg mice represented ~13–17% of the normal population [67, 68]. Collectively, these models provide indirect evidence that androgen activity during the postnatal period may normally act in concert with FSH to orchestrate maximal Sertoli cell proliferation. The effect of absent FSH receptor signaling on male mouse fertility has been reported to differ in two independent FSHR–/– models. Both were generated by the targeted disruption of the 5’ region including exon I of the FSH receptor gene using the same mouse strain [81, 82]. Despite similar production strategies, the loss of the FSH receptor was reported to have either no effect [81] or reduce [84] male fertility. Although abnormal sperm were reported in FSHR–/– testes [87], the analysis of reduced fertility was limited to first litters produced by FSHR–/– or normal males [84], so any initial reduction in fertility may be transient and due to delayed sexual maturity of males in one FSHR–/– line [84]. Similar findings to this FSHR–/– line were obtained after the immunization of 3-week-old mice to peptides representing N-terminal extracellular regions of the FSH receptor. Fertility of immunized males was delayed up to 1 week and initial progeny size reduced up to 60% [88]. Further research is warranted to assess the longer term impact of absent Sertoli cell FSH receptor function on male mouse fertility. 2. Gain of FSH Activity Investigating the selective gain of FSH function in mice has provided a valuable complementary approach to studies examining the loss of FSH activity. These studies include the addition of excess FSH activity to normal animals, or the restoration of FSH to the gonadotrophin-deficient mouse models described earlier. For example, the effect of FSH on perinatal Sertoli and germ cell development in hpg mice has been examined in detail by stereology [67, 72, 73]. In the absence of FSH, Sertoli cell numbers in late fetal life (18 dpc) were normal in untreated hpg testes, but significantly lower at birth (19–21 dpc) and reduced to ~50% of normal throughout postnatal life [57].

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Therefore, FSH activity in mice may have little effect on Sertoli cell proliferation until the late fetal period. Administration of rhFSH during the first 2 weeks after birth, followed by sustained maximal testosterone treatment beyond 3 weeks of age (i.e., after most Sertoli proliferation), increased Sertoli cell numbers twofold and thereby the ultimate size and sperm production of mature hpg testis [72]. Although final Sertoli cell numbers remained ~40% below wild-type testes levels, the potential additive effects of higher rhFSH doses were not examined [72]. In contrast, the administration of rhFSH after 3 weeks of age had no significant effect on Sertoli cell numbers in hpg testes [72, 73]. Absent mitogenic FSH activity after the postnatal period was consistent with previous studies that showed FSH treatment of mature hpg males stimulated 3H-thymidine incorporation in germ but not Sertoli cells and had no effect on testicular testosterone levels [89], reinforcing that these cellular effects were specific for FSH. Therefore, these early observations suggested that a limited postnatal epoch for expansion of the Sertoli cell pool was retained in the FSH- and androgen-deficient background of hpg mice. However, there is also evidence that Sertoli cells continue to proliferate at a low rate even in gonadotrophindeficient hpg mice [57] and that they proliferate to a small extent beyond postnatal age in response to exogenous testosterone [67] or hCG alone [68]. Sertoli cells in untreated hpg males were reported to reach 65% of normal numbers in 70- to 120-day-old hpg mice, and it was proposed that that FSH may be required to stop Sertoli cell proliferation during postnatal development [57]. However, separate studies showed Sertoli cell numbers were only 30–40% of normal in 63- to 70-day-old hpg testes [67, 68], which was lower than the numbers (~50% of normal) reported for 20-day-old mice in the earlier study [57]. Therefore, further investigation is required to clarify the extent of Sertoli cell proliferation during and beyond the postnatal window in the absence of FSH. Further analysis of hpg mice treated with exogenous rhFSH was restricted by its inaccessible delivery prior to parturition as well as the generation of circulating anti-FSH antibodies [73], which may diminish longer term FSH effects via immunoneutralization [44]. To overcome these limitations, transgenic FSH expression was achieved by the simultaneous expression of human FSH β and α subunits driven by the rat insulin II gene promoter [71]. Transgenic FSH expressed in hpg mice independently of GnRH requirement and LH secretion allowed FSH actions to be studied in the absence of LH-mediated steroid production [67, 71]. This transgenic-hpg strategy permitted the investigation of perinatal and longer term FSH effects, and the

evaluation of dose-dependent FSH effects in isolation or combined with steroids [67, 71], without confounding steroidal feedback pathways inhibiting FSH secretion [90]. Transgenic human FSH dose-dependently increased testis weight in hpg mice, although there was a threshold required for transgenic serum FSH levels for testicular effects [71]. Stereological analysis of transgenic-hpg testes confirmed the mitogenic effects of FSH on Sertoli and spermatogonia cells, and further showed that this FSH-mediated proliferation was not significantly affected by maximal testosterone treatment after weaning age (3 weeks old) [67]. The later finding differed from the reported inhibitory effects of neonatal testosterone treatment on FSH stimulated Sertoli cell proliferation [72]. The contrasting inhibitory or neutral effects of neonatal versus postweaning testosterone treatment supported the view that FSH regulation of Sertoli cell replication is established during early postnatal development. Furthermore, transgenic FSH can restore Sertoli cell numbers in hpg testes to phenotypically normal mature levels in a dose-dependent fashion despite the absence of LH, and without change in a persistently low intratesticular testosterone concentration (C. Allan et al., submitted). Yet, in the same mice spermatogonial expansion and meiotic spermatocyte development reached ~50% of normal, but there was only a minimal level of postmeiotic spermatid formation (C. Allan et al., submitted). Therefore, despite stimulating the full Sertoli cell complement, isolated expression of FSH does not initiate complete spermatogenesis in gonadotropin-deficient males, highlighting the requirement for other factors such as testosterone for full Sertoli cell function. In contrast to a stimulatory effect in hpg testes, the pituitary-independent expression of transgenic FSH (at 1–7 IU/L) had no effect on testis weight, sperm production, or fertility of normal male mice (C. Allan et al., unpublished data). In a separate transgenic mouse model, ectopic expression of human FSH via the metallothionein gene promoter also had no phenotypic effect in males at moderate serum levels (48 IU/L), whereas massively supraphysiological FSH levels (151,000 IU/L) increased serum testosterone (19-fold) and epididymal sperm counts (75%) but had no effect on testis weight or histology [91]. The extreme elevation in serum FSH was associated with male infertility, enlarged seminal vesicles, and altered reproductive behavior, probably due to the cross-reactivity of transgenic FSH at the LH receptor [91], with the excessive FSH levels possibly sacrificing some of the specificity of the model. The inability of excessive FSH to overstimulate Sertoli cell proliferation and testis size in these transgenic mouse models contrasts with previous work in rats, which showed supraphysiological

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FSH elevated Sertoli cell numbers and testis weight above normal [24, 25]; this discrepancy remains unexplained. In comparison, pituitary expression of a human FSHβ subunit transgene generating a functional interspecies heterodimer (human-β/mouse-α) increased both serum testosterone (78%) and testicular weight (22%) relative to normal (presumably by increasing total circulating bioactive FSH and Sertoli and germ cell numbers) and had no effect on male fertility [92]. An underlying mechanism for the differences in these phenotypes remains to be explained. Gonadotrophin-deficient mice have also been used to investigate the physiological actions of an activated mutant human FSH receptor (denoted FSHR+), which was identified following the unexpected fertility of a hypophysectomized FSH/LH-deficient man who required androgen replacement therapy [93]. It was proposed that this mutation sustained FSH receptor signaling in the absence of ligand, because the receptor had a single amino acid substitution in the third intracytoplasmic loop that corresponded to ligand-independent activating mutations in the related LH [94] and TSH [95] receptors. To evaluate the in vivo activity of FSHR+ in the absence of its cognate ligand, transgenic FSHR+ expression was targeted to Sertoli cells of FSHdeficient hpg males. Transgenic FSHR+ doubled hpg testis size [96], and stereological analysis showed that FSHR+ enhanced Sertoli and spermatogonia proliferation in hpg testes, producing an FSH-like response that resembled the testicular phenotype in transgenic FSHhpg mice. Therefore, the in vivo actions of FSHR+ in FSH-deficient hpg mice supported the proposal that this mutation provided a constitutive “gain-of-function” FSH receptor, albeit biochemically subtle. Because the man with the activating mutation underwent development of complete spermatogenesis prior to the onset of gonadotropin deficiency, fuller replication of this clinical paradigm would require further studies. Alternative mouse models with selective preservation of FSH activity in the absence of LH were generated by homologous recombination to permanently disrupt the LH receptor gene [97, 98]. Infertile LHR–/– mice had small testes, yet Sertoli cell numbers were equivalent to normal adult testes. The normal complement of Sertoli cells in either LHR–/– or LH-deficient transgenic FSH-hpg testes demonstrated that the full mitogenic potential of FSH on Sertoli cells does not require the LH pathway. A role for LH during normal Sertoli cell proliferation cannot be entirely excluded because LHR–/– males exhibit higher than normal blood FSH concentrations, which may partly overcome loss of LH activity [98]. Yet, in the absence of LH activity, FSH receptor signaling was not sufficient to support the completion of spermatogenesis, which

was arrested during postmeiotic spermatid maturation. Recent work demonstrated a striking combined effect of FSH and testosterone on Sertoli cell function. Transgenic FSH had additive effects with testosterone on meiotic spermatocyte numbers, and a strong synergistic effect with testosterone on postmeiotic spermatid maturation in hpg testes [67]. Therefore, the complete Sertoli cell in vivo response requires simultaneous FSH and androgen activity. The future challenge in this research area will be to determine which biological pathways are activated by the FSH and androgen responses, and why excessive circulating FSH promotes testicular hypertrophy and Sertoli proliferation beyond normal levels [24, 25, 92], yet fails to promote complete postmeiotic male germ cell development when compared to the qualitatively complete spermatogenic actions of testosterone alone, which reach plateau levels well below normal values [63]. In summary, deliberately created genetic mouse models have provided definitive in vivo evidence that the FSH response provides the primary but not sole mitogenic stimulus for the postnatal replication of Sertoli cells. Complete in vivo Sertoli cell proliferation and function also require at least the addition of androgen activity, which is crucial for the ability of Sertoli cells to functionally support the full complement of postmeiotic germ cell development. Based on concerted observations from rat and mouse models (many outlined in Table 11.1), the contributions of FSH to Sertoli and germ cell development are summarized in Figure 11.1.

C. Pig The pig has provided a useful large mammal model to investigate the physiological actions of FSH. In conventional domestic pigs, pituitary FSH becomes immunochemically detectable at 60–75 dpc [99, 100] followed by a gradual rise in serum FSH from 80 dpc until 3 weeks after birth [101], well after gonadal FSH receptor expression at ~28 dpc as detected by reverse transcriptase–polymerase chain reaction (RT-PCR) [102, 103]. Administration of porcine FSH (nonrecombinant) to prepubertal boars from 8 to 40 days of age was reported to increase Sertoli cell proliferation, based on indirect morphometric measurements showing increased seminiferous tubule length [104]. During testis development in immature Brazilian Piau pigs, two predominant phases of Sertoli cell proliferation were identified by morphometric analysis. The Sertoli cell pool expanded sixfold in the first month and a further 80% between 3 and 4 months of age, prior to sexual maturation at 4–5 months [105]. Circulating levels of FSH were elevated during each proliferative phase,

TABLE 11.1 Models to Study FSH Activity In Vivo Serum Species and model

Testis weighta

Germ cell response (total number)a,b

Fertility

Referencesc

↑

Sp > N

?N

[15–18]

N

?

?

?N

[15, 17]

↑

↑

Sg, Sc, Sp > N

?N

[24, 25]

↓

↑

Sg, Sp survival ↑

Infertile

[30, 31, 33, 40]

↓

No change

Sg, Sc, Sp survival ↑

Infertile

[32, 35] [52, 241]

FSH

T

Neonatal hemicastration

↑

N

↑

Adult hemicastration

↑

N

Neonatal + FSH

↑

↓−↑ ?

Immature HPX + FSH

?

↓

Adult HPX + FSH

?

↓

Sertoli numbersa

Rat

?

↓

↓

?

Sg, Sc, Sp survival ↑

?

Fetal + FSH Ab

↓?

?

?

↓

?

?

[14]

Adult + FSH Ab

↓

N

N

?

Sg, Sc, Sp < N

?

[43, 44]

Immunized to FSHR

?

↑

N

?

Sp < N

?

[53]

Immunized to GnRH + FSH

Neonatal + GnRH-A

↓

↓

↓

↓

Sp < N

Reduced

[37, 159, 242, 243]

Adult + GnRH-A

↓

↓

↓

N

Sc, Sp < N

Infertile

[38]

Adult + GnRH-A + FSH

?

↓

↓

N

Sg, Sc, Sp survival ↑

?

[39]

[72]

Mouse Gain of FSH activity Neonatal hpg +FSH

?

–

↑

↑

Sg, Sc, Sp ↑

Infertile

Immature hpg +FSH

?

–

↑

No change

Sg, Sc, Sp ↑

Infertile

[73]

Pituitary tgFSH

↑

↑

↑

?

?

N

[92]

Ectopic tgFSH

↑

↑

N

?

?

N-infertile

[91]

hpg + tgFSH or tgFSHR

–

–

↑

↑

Sg, Sc, Sp ↑

Infertile

[67, 71, 96]

LHR–/–

↑

↓

↓

N

Sg, Sc, Sp ↓

Infertile

[97, 98]

Loss of FSH activity Immunized to FSHR

?

N

?

?

?

Reduced

[88, 244]

FSHβ–/–

–

N

↓

↓

Sg, Sc, Sp ↓ (Leydig N)

N

[80, 83]

FSHR–/–

↑

↓

↓

↓

Sg, Sc, Sp ↓ (Leydig ↓)

N-reduced?

[81, 82, 84]

Monkey Bonnet M. radiata Adult immunized to FSH

↓

N

?

?

Sg ↑-N; Sc, Sp ↓-N

Reduced-N

[165, 168, 169]

Adult immunized to FSHR

?

?

?

?

Sc ↓?, Sp ↓

Reduced

[169]

Adult hemicastration

↑

N

↑

?

Sp, Sc, Sp ↑

?

[163, 245]

Adult hemicast. + FSH Ab

↓?

N

N

?

Sc ↓?

?

[163]

Adult + GnRH-A + T

↓

N

?

?

Sg ↑, Sp ↓

?

[168]

Rhesus M. mulatta ?↑

N

↑

↑

Sg ↑?

?

[156]

Juvenile + GnRH

?

?

↑

↑

Sg ↑

?

[155]

Adult HPX

↓

↓

↓N

N

?

?Infertile

[246]

Juvenile + FSH

↑d

Adult HPX + T + FSH

?

↓

↓N

N

Sg

Adult hemicastration

↑

N

↑

N

Sg, Sc, Sp ↑

?

[246]

?

[162]

Adult immunized to FSH

?

N

↓

?

Sp ↓-n

?

[167]

Adult + FSH Abs

?

N

↓

?

Sp ↓-N

?

[166]

Adult + GnRH-A

↓

↓


?

Sg, Sc, Sp < N

?

[170]

Adult + GnRH-A + FSH

?

↓

N

?

Sg N, Sc, Sp ↑d (but
?

[170]

?↓

N

?

↓ then N

?

N

[159, 247]

Cynomolgus M. fascicularis

Marmoset Callithix jacchus Neonatal + GnRH-A

Abbreviations: Ab, antibody/antiserum; T, testosterone; GnRH-A, GnRH antagonist/analogue; hpg, hypogonadal; HPX, hypophysectomy; N, normal; tg, transgenic; –/–, homozygous deletion. a Changes in testis size or Sertoli or germ cell numbers refer to differences between untreated controls in the hpg, HPX, and GnRH-A treated animals. b For comparison, cell types were grouped into Sg, spermatogonia; Sc, Spermatocytes; Sp, spermatids. c Cited studies restricted to those using rhFSH or highly purified FSH with no detectable in vitro or in vivo LH activity. d Indicates germ cell changes relative to treatment preceding FSH.

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Sc

PS

PS

Sc

Sg

RS

Sg

RS

ES

ES

Sertoli

Sertoli

FSH

Sc

T

PS

Sg

RS ES

Sertoli = Additive = Synergistic

FSH + T FIGURE 11.1 FSH contribution to Sertoli and germ cell development (see color plate). Arrow size approximates the contributions of FSH or testosterone (T) alone or in combination on the development of the different testicular cell types illustrated (based on rat and mouse studies reviewed in the text). Germ cells grouped as follows: Sg, spermatogonia; Sc, early spermatocytes; PS, pachytene spermatocytes; RS, round spermatids; ES, elongated spermatids.

the latter coinciding with the first wave of early spermatocyte development. These findings supported an earlier proposal that gonadal development was coupled to FSH concentrations in boars [101], and the findings were consistent with the positive FSH–Sertoli cell relationships reported in rats [13, 14], mice [54, 73], and sheep [106]. In contrast, Meishan, White composite, and crossbreeds have plasma FSH levels in young or mature boars that correlate negatively with testicular size [107, 108]. FSH levels were determined after most Sertoli cell replication (from 4 months old), and it is difficult to determine if this negative correlation reflected a direct inhibitory effect of FSH on the testis or was due to other related factors. For instance, Meishan boars exhibit higher postnatal-pubertal serum FSH, LH and androgen levels and smaller adult testes compared to conventional European breeds [109, 110]. In addition, thyroid hormone was recently found to account for some interbreed Sertoli cell differences [111]. Therefore, a negative correlation between FSH and testis growth may be confounded by strain differences

for other endocrine or genetic characteristics that influence gonadal development. Recent studies further proposed that the early postnatal period was important for establishing final Sertoli cell numbers in mature boars, but was not tightly coupled to FSH secretion [112, 113]. These studies estimated Sertoli cell numbers and proliferation using morphometric extrapolation via immunohistochemical detection of GATA-4 (Sertoli cell marker) and the Ki-67 antigen (cycling cell marker) [114]. However, comparison of postnatal Sertoli cell numbers between breeds using this technique produced inconsistent results (e.g., [114] compared to [113]), possibly due to body weight differences [113], and greater numbers of Ki-67 positive cells did not correspond to higher Sertoli cell numbers in different breeds [113]. In addition, the proportion of Ki-67 positive cells peaked between 90 and 105 dpc, well before the approximately eightfold postnatal increase in Sertoli cell numbers [113]. A perinatal rise in FSH also preceded the postnatal increase in Sertoli cell numbers. Therefore, although it is possible the hormonal regulation of Sertoli cell proliferation

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differs in pigs compared to most other species examined to date, any proposed diminished role for FSH in pig Sertoli cell proliferation should be interpreted cautiously, especially if based only on observational data. Closer examination by more direct and selective manipulation of FSH activity (e.g., ablate-replace approaches) would be of interest.

III. ANIMAL MODELS TO STUDY IN VIVO FSH ACTIONS: SEASONAL BREEDERS A. Hamster Although rats and mice are the most commonly used laboratory models for studying the FSH response in vivo, they do not exhibit the seasonal pattern of reproduction or gonadotrophin levels found in most mammalian species, possibly due to selective captive breeding for maximal reproductive efficiency. Many mammals exposed to the annual changes in photoperiod length are reproductively competent for only a brief or defined period of the year. The hamster has provided a valuable seasonal rodent model to investigate photoperiod-dependent changes in FSH activity and testicular function. In the laboratory, artificial photoperiod exposure has been used to regulate the seasonal breeding and spermatogenic activity of mature hamsters. As detailed later, “long-day” photoperiod exposure stimulates testicular function in mature hamsters and is associated with increased levels of circulating FSH. In juvenile hamsters, the effects of perinatal photoperiod exposure on testicular maturation are also well documented. In immature Siberian (Djungarian) or Syrian (golden) hamsters, 3H-thymidine incorporation studies showed that the percentage of proliferating Sertoli cells is greatest during the perinatal stage and declines during postnatal maturation [115, 116]. Although information about the ontogeny of FSH activity during prenatal gonadal development in hamsters is scarce, several studies in juvenile Siberian hamsters showed that circulating FSH levels and the rate of testicular maturation is regulated by exposure to short (<12.5 hr light/day) or long (16 hr light/day) photoperiods during gestation and/or after birth. It is well established that testicular maturation in juvenile Siberian hamsters is influenced by early photoperiod information received during late gestation, which is regulated by the maternal pineal gland and melatonin secretion [117, 118]. Exposure to an intermediate photoperiod (14 hr light/day) during postnatal life stimulated rapid testicular maturation in young hamsters that experienced short photoperiod

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(10–12 hr light/day) in gestation, but inhibited testicular maturation in those gestated on long photoperiod (16 hr light/day) [117, 119, 120]. Photostimulated acceleration of testicular development in these models is associated with elevated serum FSH levels. Exposure of young (18-day-old) Siberian hamsters to long photoperiods increased serum FSH levels and testicular weight within 1 week, which preceded an elevation of LH and testosterone levels [121–123]. By comparison, males continuously exposed to short photoperiods (10 hr light/day) in gestation and after birth exhibited low serum FSH levels and no postnatal testicular growth [120]. Furthermore, long-day photostimulation of gonadal growth in juvenile (49-day-old) Siberian hamsters was suppressed by administration of porcine follicular fluid [121], which contains inhibin B, a negative regulator of FSH secretion (see Chapter 14). One stereological study of testis development in Siberian hamsters suggested that the durations of preand postnatal photoperiod had no effect on the ultimate Sertoli cell numbers by 29 days of age, even though a short prenatal photoperiod (4 hr light/day) delayed the onset of spermatogenesis and altered postnatal Sertoli cell proliferation rates to a small extent [115]. Collectively, these findings suggest that the photostimulated elevation of circulating FSH promotes the initiation of testicular growth in immature Siberian hamsters, which may reach a threshold during postnatal life to maintain normal Sertoli cell numbers. Note that in contrast to Siberian hamsters, gonadal development in immature Syrian (golden) hamsters (<5–6 weeks old) exhibits a photoinsensitive period (until 5–7 weeks of age), after which exposure to short-day photoperiods induces testicular atrophy [124, 125]. In contrast to the photoperiod sensitivity differences between immature Siberian and Syrian hamsters, mature males of both species exhibit testicular regression or recovery in response to short- or long-day photoperiods, respectively. Mature hamsters transferred from a short-day to a long-day photoperiod (as would naturally occur in spring/summer) exhibit increased gonadotrophin levels and testicular weight and restored reproductive function. Similarly, in mature Siberian hamsters, circulating FSH levels and total FSH binding sites per testes were increased between 10 and 47 days after transfer to long-day photoperiods [126]. The rise in hamster FSH production was primarily dependent on the rate of pulsatile gonadotrophinreleasing hormone (GnRH) secretion [123, 127, 128], which may initiate the seasonal cycles of reestablishment of testicular function and spermatogenesis in response to altered rhythms of melatonin secretion [120, 129]. This photodependent rise in FSH was not altered by exposure to prospective female mates,

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unlike the rapid rise in serum LH levels when photostimulation was combined with exposure to females [128]. Thus, the hamster provides a valuable model to investigate differential gonadotrophin regulation by the hypothalamus. Conversely, transferring Siberian or Syrian hamsters from long to short photoperiods significantly decreases serum FSH levels and total FSH receptor content of regressed testes [126, 130–132]. FSH treatment alone completely restored spermatogenesis in regressed testes of photoinhibited [133, 134] or hypophysectomized Siberian hamsters [134]. Yet, circulating FSH alone is not sufficient to complete spermatogenesis in conventional nonseasonal mouse and rat models. Therefore, the hamster provides a valuable model to study the pivotal role of FSH in seasonal testicular function, which is regulated by both FSH secretion and availability of the FSH receptor, but can only be cautiously extrapolated to nonseasonal mammals. Few studies have enumerated the absolute changes in Sertoli cell numbers associated with the FSH response in mature hamster testes. In one example, immunohistochemical detection of proliferating cell nuclear antigen (PCNA) suggested that no Sertoli (or Leydig) cell proliferation occurred during the photoperiod-induced reestablishment of testicular function in adult Syrian hamsters, which contrasted with detectable PCNA in early spermatogenic cells [135]. This finding supported previous observations that Sertoli cells decrease in volume but not number and become morphologically immature in the regressed testes of Syrian hamsters during short photoperiods [136, 137]. Sertoli cell dedifferentiation paralleled spermatogenic disruption with primarily spermatogonia and only occasional spermatocytes and round spermatids present. However, a recent study using more direct stereological analysis to enumerate cell populations reported total Sertoli cell numbers were decreased up to 66% in Siberian hamsters exposed to short photoperiods, which could be restored to normal levels by FSH treatment alone [138]. This surprising observation may indicate that FSH seasonally regulates the absolute numbers of Sertoli cells in the adult hamster, which challenges the previous conclusions of a constant mature Sertoli cell population. The need for more detailed quantitation of Sertoli cell numbers in these seasonal models of reproduction is clearly indicated.

B. Sheep The domestic sheep is a well-studied “short-day” seasonal breeder. In fetal sheep, the pharmacological inhibition of GnRH activity (by sustained delivery of the GnRH analogue buserelin) during the second half

of gestation suppressed gonadotrophin production, and at birth the testis size and total Sertoli cell numbers were reduced by 40 and 45%, respectively [139]. Likewise, decreased testicular development followed the postnatal immunization of ram lambs against ovine β-FSH from birth to 100–160 days old [106]. Treatment reduced testis size and total Sertoli cell numbers by 49–63% and 18–43%, respectively; blood concentrations of LH and testosterone remained normal. In addition, the production of type B2 spermatogonia and early meiotic spermatocytes was reduced ~45%. An earlier study reported that ram Sertoli cell numbers stop dividing around 40 days of age, although this work did not evaluate numbers in rams beyond 100 days of age [140]. On the other hand, testicular hypertrophy after hemicastration of neonatal lambs was associated with elevated serum FSH levels (versus unchanged serum LH and testosterone) and increased Sertoli cell replication [141]. Collectively, these findings showed that FSH regulates the perinatal proliferation of both Sertoli and early germ cell populations, and they also provided support for the reported significant linear relationship between the level of circulating FSH and testicular size [142], as well as the photoperiod-induced FSH-related endocrine changes that coincide with restoration of testicular size and function [143, 144]. There are limited but conflicting data regarding the seasonal effect on Sertoli cell populations in adult sheep. One study of adult Soay rams suggested that the size but not number of Sertoli cells changed between long or short photoperiod exposures [145]. More recent stereological examinations of Corriedale ram testes found a significant reduction (23%) in total Sertoli cell numbers from autumn (breeding season) to winter (nonbreeding), although it was speculated that this decline may be due to aging rather than season [146].

C. Deer and Horse An early study in red deer (Cervus elaphus) found that the total testicular Sertoli cell population declined 27% in the sexual season (autumn) compared to the quiescent period (spring) [147]. More recent stereological studies in stallions suggested total Sertoli cell numbers annually increase during the breeding season [148–150], which also coincided with increased circulating FSH concentrations [150, 151]. A limitation of investigating large outbred animals is highlighted by the large numbers of stallions necessary to overcome the inherently large variations in testicular size and sperm production among genetically unmatched animals (obtained from commercial abattoirs) [150]. However, these findings highlight the need for definitive enumeration studies of the Sertoli cell population

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in seasonal breeders exposed to annual photoperiodic variation, to precisely determine whether or not annual FSH-regulated changes in testicular function are coupled to fluctuations in the absolute Sertoli cell population. If the reported changes in Sertoli cell numbers in mature hamster, horse, deer, and sheep are substantiated, the photoperiod-dependent paradigm raises several intriguing possibilities for Sertoli cell control in seasonal breeding species, which presumably must involve Sertoli cell apoptosis and some degree of dedifferentiation, followed by proliferation and redifferentiation. Alternatively, Sertoli cell numbers may remain constant throughout the changing seasons, regressing to a dormant immature phenotype during the nonbreeding season, followed by reactivation. In either case, the Sertoli cell of seasonal breeders must undergo major functional transformations between active and quiescent reproductive periods, and these seasonal models provide a valuable opportunity to examine the role of FSH in the “reactivation” of mature Sertoli cell function.

IV. NONHUMAN PRIMATES Most research into the actions of FSH in primates has utilized two Old World monkey species, the rhesus (Macaca mulatta) or bonnet (Macaca radiata) monkey. During normal testicular development in the rhesus monkey, two predominant phases of Sertoli cell proliferation were detected, the first in prepubertal life (10–15 months old) and the second during pubertal (~3 years old) development (reviewed in [152]). A recent assessment of prepubertal Sertoli cell proliferation, by BrdU incorporation and estimation of total cell numbers, showed that most proliferation coincided with the prepubertal rise of serum gonadotrophins [153], around 25–29 weeks of age [154]. The pubertal phase of Sertoli cell proliferation was also associated with elevated gonadotrophin secretion and was prematurely induced in juvenile (15–17 months old) rhesus monkeys by administration of pulsatile GnRH [155] or rhFSH alone [156]. The precocious response to FSH also included a limited level of spermatogonial development. The selective administration of recombinant single-chain human LH (shLH) to juvenile males also initiated early Sertoli cell proliferation [156]. The effects of rhFSH and shLH on Sertoli cell proliferation were not additive, suggesting the pathways activated by each hormone in Sertoli cells share a common regulatory endpoint. However, the combined rhFSHshLH response stimulated more meiotic germ cell development than either treatment alone [156]. These findings supported earlier work that suggested highly purified FSH, hCG, or testosterone treatment each

183

independently enhanced Sertoli cell proliferation in juvenile rhesus males [157, 158], which further showed that hCG but not FSH also stimulated the morphological differentiation of Sertoli cells [158]. Therefore, FSH and androgen may have equally important contributions to Sertoli cell replication in immature rhesus monkeys. Investigation of the marmoset (Callithrix jacchus), a New World monkey species, showed that the total Sertoli cell numbers at the end of the neonatal period to beginning of infancy were equivalent to the numbers in adults [159]. Currently the absence of a homologous FSH immunoassay has limited direct assessment of the role of FSH activity in marmosets. However, the administration of GnRH-A during early marmoset development (1–14 weeks of age) initially decreased the total Sertoli cell population by ~35%, although Sertoli cell numbers were restored to normal by adulthood [159]. Therefore, Sertoli cell proliferation can occur beyond the “neonatal” stage of marmoset development. In contrast, GnRH-A treatment of adults had no effect on Sertoli cell numbers, suggesting that gonadotrophin-regulated proliferation of Sertoli cells is limited to immature marmosets. In the cebus monkey (Cebus apella), another New World species, Sertoli cells numbers increased markedly during neonatal and infantile development (birth to 12 months old) then remained stable during puberty, but there was no evaluation of total numbers in older mature males [160]. Other in vivo studies showed that adult monkey Sertoli cell numbers are resistant to changes in FSH activity. In the cynomolgus monkey (an Old World species), BrdU-labeling studies did not encounter proliferating Sertoli cells in adult testis [161]. Hemicastration of adult rhesus males increased FSH secretion and testis hypertrophy, but had no effect on Sertoli cell numbers, indicating that, like the mouse and rat, FSHstimulated proliferation of Sertoli cells is restricted to immature monkeys [162]. Although Sertoli cell numbers were unchanged in hemicastrated mature rhesus monkeys, the absolute numbers of spermatogonia and meiotic and postmeiotic germ cells were all increased above normal. Considering the prevailing view that mature Sertoli cells have a fixed germ cell carrying capacity, this finding suggests that the germ cell capacity of the rhesus Sertoli cell does not normally operate at its ceiling. Therefore, normal circulating FSH levels in rhesus males may provide insufficient stimulation to support the higher potential spermatogenic capacity of Sertoli cells and ultimately the whole testis (assuming Sertoli cell numbers remain constant). Hemicastration of the adult bonnet monkey also elevated serum FSH levels and produced hypertrophy of the remaining testis [163], although the absolute

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numbers of Sertoli cells were not determined. The passive administration of antiserum to ovine FSH reversed this testicular hypertrophy and decreased the primary spermatocyte population [163]. Mature bonnet and rhesus monkeys also experience seasonal variations in maximal testicular function, characterized by reduced meiotic and postmeiotic germ cell populations in the nonbreeding compared to breeding season, but no change in absolute Sertoli cell numbers [164]. Primate FSH action has been investigated by active immunization or passive immunoneutralization approaches (reviewed in [152]), similar to those described for the rat. For example, long-term immunization (up to 2 years) of mature bonnet monkeys to ovine FSH produced infertile males with transiently decreased spermatozoa production [165]. Serum testosterone levels remained normal, suggesting that the induced infertility was specific to disruption of FSH action. Total Sertoli cell numbers were not determined, although the gradual restoration of fertility in most (9 out of 10) immunized animals suggests no overt change to the total Sertoli population. In comparison, the use of active immunization against exogenous FSH or passively administered anti-FSH sera in adult rhesus monkeys produced more variable and transient reductions in sperm production [165–167]. Therefore, although antibody-driven suppression may selectively diminish FSH function without altering serum LH or testosterone levels in both rhesus and bonnet monkeys, longer term use is limited due to inconsistent immunogenicity or transient effects on spermatogenesis [165–169] and, by inference, variable disruption of Sertoli cell function. The active immunization of mature male bonnet monkeys with the extracellular portion of the sheep FSH receptor was reported to prolong the reduction in both sperm production and fertility compared to FSH immunization [169]. Serum testosterone remained normal in this model, which suggested the effects were specific to the Sertoli cell and did not involve the related LH receptor found on steroidogenic Leydig cells. Similar effects of FSH on spermatogenic regression have been obtained in GnRH-A treated monkeys. The suppression of gonadotrophin secretion by GnRH-A administration resulted in the progressive loss of more advanced germ cells [170]. Although GnRH-A treatment did not completely suppress circulating FSH levels in adult cynomolgus monkeys [161], the simultaneous administration of highly purified FSH delayed the testicular regression induced by GnRH-A, maintained a spermatogonial subpopulation (SgA), and reduced the loss of round spermatids [170]. A range of models used to investigate FSH in vivo actions in the monkey are summarized in Table 11. 1.

V. HUMANS Although ethical and practical constraints on human experimentation limit the ability to investigate experimentally the role of FSH in human spermatogenesis, considerable insight has been obtained from observational studies of the effects of FSH deficiency and its treatment. FSH deficiency is relatively uncommon and occurs either as part of combined gonadotrophin (FSH and LH) deficiency, with or without other pituitary hormone defects, or very rarely as isolated FSH deficiency. Although the latter is the most informative, few cases have been reported; note, however, that it may pass unrecognized. Combined gonadotrophin deficiency is due to either hypothalamic or pituitary disorders that have a congenital (prepubertal) or acquired (postpubertal) onset. The clinical manifestations of combined gonadotrophin deficiency are largely due to the LH deficiency, which reduces testicular testosterone output, leading to clinical features of androgen deficiency. By contrast, FSH receptors are expressed exclusively on Sertoli cells so that FSH deficiency produces no symptoms other than reduced spermatogenesis and infertility. Even this may only become apparent in the minority of men seeking fertility at any one time and in whom mild defects in sperm output and/or function are not overcome by fertile female partners. Hence, it is inherently likely that isolated FSH deficiency in men may be overlooked and underdiagnosed. Historically, it has long been axiomatic that, following completion of puberty, total hypophysectomy leads to full regression of spermatogenesis. Persistent spermatogenesis after hypophysectomy may reflect incomplete pituitary ablation [171] and/or effects of exogenous testosterone [172] maintaining residual foci of spermatogenesis. The major limitations of observational findings from men with hypopituitarism are that (1) both LH and FSH are deficient so that the relative roles of LH (and consequently testosterone) and FSH cannot be readily differentiated and (2) the confounding effects of concomitant deficits in other pituitary hormones, such as thyroid and adrenal hormones and growth hormone and IGF-I that also influence spermatogenesis. Thus the utility of observational studies of gonadotrophin withdrawal and treatment on human spermatogenesis is limited. Few well-controlled experimental studies of the hormonal control of human spermatogenesis have been reported. From these, however, good evidence exists to show that FSH has a role in induction and quantitative maintenance of human spermatogenesis when combined with LH/hCG, but not testosterone, presumably due to a threshold requirement for intratesticular testosterone that is unobtainable by administration of

Chapter 11

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conventional testosterone doses in humans [173]. In men with complete gonadotrophin deficiency, FSH combined with hCG effectively induces spermatogenesis, whereas when FSH is combined only with testosterone at conventional androgen replacement doses, no stimulation of spermatogenesis is observed [174]. This strongly suggests either that higher testosterone doses or intratesticular testosterone concentrations are required to induce spermatogenesis, as was found in nonhuman primates [173]. Conversely, during androgen-induced suppression of gonadotrophins in normal men, spermatogenesis is partially suppressed and can be restimulated by additional treatment with FSH or hCG [175–177]. The latter studies involved men with previously normal spermatogenesis suppressed by exogenous testosterone. Although gonadotrophin suppression is equivalent to that in untreated men with complete gonadotrophin deficiency due to hypophysectomy or congenital hypogonadotrophic hypogonadism, high circulating testosterone concentrations may partially maintain spermatogenesis even in the absence of gonadotrophin stimulation. Hence, these studies support a role for FSH in the induction and maintenance of spermatogenesis but only when coupled with sufficient testosterone. Selective gonadotrophin deficiency with otherwise normal pituitary function is uncommon but is regularly diagnosed as part of a spectrum of congenital disorders known collectively as idiopathic hypogonadotrophic hypogonadism (IHH), which includes Kallmann’s syndrome and its variants [178]. Increasingly, the genetic mutations underlying these conditions are being identified [179]. Although IHH disorders remove the confounding issue of nongonadotrophic pituitary hormones, the dissection of LH from FSH effects on human spermatogenesis remains intractable. Another insight into the role of FSH in human spermatogenesis is provided by interventional studies using FSH as part of gonadotrophin therapy for gonadotrophin-deficient infertile men. These cases constitute a small but important subset of infertile men because gonadotrophin therapy is effective at inducing spermatogenesis and facilitating paternity as one of the few specifically treatable causes of male infertility [180]. Effective gonadotrophin regimens are based on hCG purified from the urine of pregnant women together with FSH purified from human pituitaries [181], from the urine of menopausal women [182], or from recombinant FSH [183]. Pituitary extracts are no longer used due to the risk of transmitting Creutzfeldt-Jakob disease [184–186] and urinary extracts are increasingly supplanted by recombinant FSH [187]. Because gonadotrophin deficiency is an uncommon cause of male infertility, only few relatively small studies have

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reported the efficacy of gonadotrophin therapy to induce spermatogenesis and paternity. Among the 18 available studies [181, 182, 187–202], only 6 involve more than 20 men [182, 187, 199–202] and only 2 were single-center studies [182, 201]. The consistent positive prognostic significance of prior gonadotrophin treatment and larger pretreatment testis volume are consistent with stimulatory effects of endogenous FSH [201]. Conventionally, gonadotrophin treatment in infertile hypogonadotrophic men is based on treatment starting with hCG, a naturally long-acting analogue of LH. FSH is usually added to the treatment regimen if sperm production does not appear within 6 months of hCG treatment [201]. Due to this clinical convention, the real need for FSH is difficult to establish rigorously. For example, a retrospective review of a large single-center experience showed that 6/43 (14%) treatment cycles required only hCG alone without the need for FSH treatment to induce spermatogenesis within 6 months of treatment [201]. Consistent findings are reported from prospective therapeutic trials of gonadotrophin therapy in gonadotrophin-deficient men, wherein hCG treatment alone (prior to FSH treatment) produced sperm output in 1/10 men within 24 weeks [183], 9/49 men within 16 weeks [187], and up to 6/32 men within 24 weeks [202]. Due to the limited time (by study design) of the hCG-alone treatment, the pooled prevalence of 18% having spermatogenesis induced by hCG alone without FSH is an underestimate. Furthermore, prolonged treatment with hCG alone is sufficient to induce human spermatogenesis even in the absence of exogenous FSH in a large proportion of gonadotrophin-deficient men [196, 203]. It remains possible but unproven that addition of FSH may accelerate the appearance and increase output of sperm. Once spermatogenesis has been induced with hCG and FSH, it is also possible to maintain it with hCG alone without further FSH administration in some gonadotrophin-deficient men [204]. Whether such responsive men represent a subset with higher endogenous FSH secretion and/or whether hCG is more often sufficient but too slow for routine clinical practice remains unclear. A new perspective may be derived from human use of a novel genetically engineered recombinant single-chain long-acting FSH analogue [205] that is currently undergoing clinical trials [206]. The most informative form of FSH deficiency is isolated FSH deficiency. All published case reports are tabulated in Table 11.2, excluding the first two cases, which, in retrospect, have clear alternative diagnoses (Kallmann’s syndrome and craniopharyngioma) excluding isolated FSH deficiency [207]. The earliest case reports became available following the

TABLE 11.2 Isolated FSH Deficiency in Men Reference

Case

Genotypea

Blood FSHb (IU/L or μg/L)

Geneticsc

Pubertyd

Virilizede

Fertility

Testes (mL)

LH (IU/L)

T (nM)

Spermf (M/mL)

[248]

1

47XXY mosaic

<3.0/5.0/–

?

D

N

?

5/5

8–28

24.1

A

[249]

2

?

<3.0/<3.0/<3.0

?

N

N

?

?

8

36

low

[250]

3

?

1.4/4.0/–

?

N

N

Infertile

“Normal”

5.4

23.9

64

[250]

4

?

2.0/5.0/–

?

N

N

Fertile

“Normal”

6.5

18.6

66

[251]

5

?

2.4/4.8/–

?

?

N

Infertile

“Normal”

2.3

24.8

6–11 A

[252]

6

?

<0.4/1.4/–

?

N

?N

?

Impalpable

4.8

33.1

[253]

7

?

<0.5/<0.5/0.9

?

?N

N

Infertile

11–20/11–20

8.1

16.9

0.6 – 1.7

[253]

8

?

3.0/5.3/5.6

?

?N

N

Infertile

11–20/11–20

5.7

24.5

0.6 – 1.7

[254]

9

?

0/1.5/2.2

?

N

N

Infertile

25–30/25–30

2.2

21.8

0

[255]

10

FSHR (Ala189Val)

23.5/–/–

F

?N

N

Infertile

4/4

16.3

14. 5

<0.1

[255]

11

FSHR (Ala189Val)

12.5/–/–

F

?N

N

Fertile

15.0/13.8

5.6

8.8

5.6

[255]

12

FSHR (Ala189Val)

15.1/–/–

F

?N

N

Fertile

13.5/15.8

4.2

15.8

<0.1

[255]

13

FSHR (Ala189Val)

20.6/–/–

F

?N

N

?

8.0/8.0

16.2

26.2

<0.1

[255]

14

FSHR (Ala189Val)

39.6/–/–

F

?N

N

?

8.6/6.0

11.1

14.7

42

[256]

15

FSBβ (Val61X)

<0.5/<0.5/–

C – No

D

R

?

1.5/1.5

24.5

4.6

0

[257]

16

FSHβ (Cys82Arg)

0/0/–

?

N

N

Infertile

3/6

12

11.0

0

[258]

17

?

0.64/–/–

?

N

?N

?

?

2.8

14.1

A

[259]

18

FSHβ (Tyr76X)

0.5/1.1/–

C – Yes

N

?N

Infertile

12/12

36.2

26.0

0

[260]

19

?

0.15/0.15/–

C – No

N

N

?

12/15

4. 0

26.8

0

Note: Averaged values where multiple hormone or semen analyses were available. a FSHR, FSH receptor, FSHβ, FSHβ-subunit. b Blood FSH at baseline (B), after GnRH stimulation (G) and after clomiphene stimulation (C) indicated as B/G/C. c F, Finnish founder effect, C, Consanguinity (yes or no). d D, delayed, N, normal. e R, reduced, N, normal. f A, aspermia (no ejaculate) , 0, azoospermia (no sperm in ejaculate).

Chapter 11

In Vivo FSH Actions

development of FSH radioimmunoassay. However, the diagnosis of isolated FSH deficiency based on blood hormone measurements has significant limitations. Compared with modern two-site immunoassays, the early radioimmunoassays had limited sensitivity (~10-fold lower) and specificity (cross-reactivity with α and LHβ subunits and dimeric LH). Furthermore, FSH responses to GnRH may be low or absent even among men with normal reproductive function. Consequently, many early case reports are inconclusive although, in concert, the consistency of the picture suggests that milder forms of isolated FSH deficiency may exist among subfertile and even among fertile men. In the last decade, naturally occurring mutations in the human FSHβ subunit and FSH receptor genes have been identified, providing incisive evidence about FSH action. So far the most salient is an inactivating mutation of the FSH receptor observed in five men from four Finnish families [208]. Despite severe interruption of FSH action sufficient to cause ovarian failure in women, testicular development, spermatogenesis, and paternity could still occur although testis size was small and sperm output low [208–210]. The latter findings were most consistent with a diminished stock of Sertoli cells, which reflected a lack of proliferative effects of FSH in early life on the ultimate pool of Sertoli cells. Leydig cell function and androgen exposure appeared to be relatively preserved if not intact. This picture of congenital genetic deficiency in FSH action is supported by more isolated observations of genetic defects in the three cases reported of genetic mutations of the FSHβ subunit gene leading to lack of circulating dimeric FSH. The phenotypes of confirmed genetic cases of isolated FSH deficiency due to mutations in the FSHβ subunit or the FSH receptor are reasonably consistent. The single more striking case with a distinctively low blood testosterone in addition to the reduced testis size and sperm output was only 18 years old and had presented to a pediatric medical center with delayed puberty [211]. Whether this case represents the additional effects of incomplete sexual maturation (as noted in one genetic mouse model of an inactive FSH receptor [84]) or highlights lost paracrine effects of FSH on Leydig cell testosterone production [212] remains to be clarified. The prevalence of isolated FSH deficiency among subfertile or even fertile men with small testes and low sperm output is not known, but current evidence suggests that significant underdiagnosis may be occurring. If so, the possible effects of FSH in augmenting sperm output among oligozoospermic infertile men [213] may have a more physiological explanation and warrant further evaluation. In theory, human cases with autonomous ectopic secretion of FSH from tumors might—as an experiment

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of Nature—provide insight into the isolated biological effects of FSH on the human testis. Ectopic secretion of hCG, the placental member of the pituitary glycoprotein hormone family, is a well-known tumor marker for trophoblast-derived cancers and is rarely secreted by nontrophoblastic tumors as well. By contrast, ectopic secretion of dimeric pituitary glycoprotein hormones is very rare and reported only from benign pituitary adenomas. Among these the most frequently described has been FSH-secreting pituitary macroadenomas [214]. These occur mostly in hypogonadal men (and postmenopausal women) where chronic loss of gonadal negative feedback unleashes sustained gonadotrophin hypersecretion, leading ultimately to pituitary gonadotroph hyperplasia and macroadenoma. In this context, the underlying defective testicular function nullifies this as an informative paradigm for FSH action. However, in cases where FSH-secreting pituitary macroadenomas are reported in men without prior testicular dysfunction, testicular enlargement is reported in one study [215] but not in a previous literature review [214]. Further systematic studies are needed to determine whether such macroorchidism has been underreported as a consequence of postpubertal onset of FSH hypersecretion in eugonadal men. Incomplete but corroborative evidence exists for the role of FSH in mediating distinctive effects on testis development and spermatogenesis in various clinical conditions. The fact that 80–90% of testis volume is attributable to spermatogenesis makes the observation of human testis volume a particularly simple way to estimate the integrity of human spermatogenesis apart from the unusual circumstance where the ductular system is obstructed (e.g., cystic fibrosis [216]). Many clinical conditions that include small testes as a feature have either intrinsic testicular pathology (e.g., myotonic dystrophy [217]) or hypogonadotrophic hypogonadism (numerous genetic neurodegenerative conditions [218]) as part of the underlying medical condition. Conversely, macroorchidism is a highly unusual but distinctive physical feature that has been reported in several conditions including fragile X syndrome [219], juvenile (but not adult) hypothyroidism [220], McCuneAlbright syndrome due to inactivating mutation of the Gs α-subunit [221], congenital adrenal hyperplasia (with testicular adrenal rests) [222], amyloid [223], bilateral testicular tumors, aspartylyglycosuria [224], and proteus syndrome [225], as well as precocious puberty, unilateral compensatory hypertrophy, and misdiagnosis of splenogonadal fusion [226]. Studies of a genetic mouse model for fragile X syndrome reproduce the macroorchidism; however, the Sertoli cell hyperproliferation was not related to either circulating FSH or FSH receptor expression levels in the affected

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Charles M. Allan and David J. Handelsman

mice lacking the FMR1 gene [227]. The involvement of perinatal FSH secretion remains an attractive common denominator pathogenic mechanism to explain these diverse conditions but this speculation remains to be fully evaluated.

VI. THE BALANCE BETWEEN FSH AND STEROIDS: THE E-T SWITCH Numerous experimental models demonstrated the interrelationships between FSH and steroids in regulating Sertoli cell function as a prelude to development of spermatogenesis. Testosterone or estradiol can regulate FSH-induced Sertoli cell proliferation or function and, conversely, FSH can regulate the steroidal responsiveness of Sertoli cells. Molecular and biochemical changes observed during postnatal development suggest that, functionally, Sertoli cells undergo a temporal switch from estradiol- to androgen-based responsiveness or regulation, denoted the estradiol-to-testosterone (E-T) switch. It is possible that a major role for FSH is to coordinate this Sertoli cell E-T switch in order to promote the maximal spermatogenic capacity of the testis through timely maturation of Sertoli and germ cells (Fig. 11.2). In this model, early postnatal Sertoli cell development is orchestrated primarily by proliferative FSH effects, during which estradiol actions enhance the Sertoli cell carrying capacity of early germ cells. As postnatal development continues, estradiol synthesis by aromatase is switched off, and subsequently postnatal androgen synthesis is activated, which, acting in synchrony with FSH-regulated events, serves as a pivotal trigger for final Sertoli cell differentiation. Neonatal testosterone treatment inhibits FSH action in gonadotropin-deficient mouse testis [72], whereas in normal mice inappropriate neonatal androgen (or excessive estradiol) exposure may block FSH-stimulated Sertoli cell proliferation, leading to premature differentiation and thereby reducing final Sertoli and germ cell populations in adult testes. Thus, the postnatal E-T switch may function to prevent premature androgen activation of spermatogenesis at a time (perinatal) when FSH is being secreted actively. Aromatase expression is highest in early postnatal Sertoli cells and declines thereafter [228], consistent with studies showing FSH-stimulated aromatase activity (5–7 fold) in Sertoli cells from 7- and 10-dayold rats compared to FSH-unresponsive activity in cells from 35- and 50-day-old rats [229, 230]. Hence, immature Sertoli cells express high levels of aromatase, which would serve to inactivate testosterone and supply estradiol for developing Sertoli cells. This could

FIGURE 11.2

Role of FSH during postnatal E-T switch in Sertoli cells. Early postnatal: FSH initially stimulates aromatase activity and promotes estradiol effects. Neonatal testosterone (T) activity in Sertoli cells is suppressed by limited AR expression, which is proposed to allow optimal FSH-stimulated Sertoli cell (and spermatogonial) proliferation before the initiation of the whole spermatogenic cycle. Mid postnatal: Aromatase activity declines and FSH is proposed to induce AR expression and increase androgen activity in Sertoli cells, in order to initiate Sertoli cell differentiation, as well as meiotic and postmeiotic germ cell development. Late postnatal: Differentiation of Sertoli cell is complete, and final Sertoli cell population is established.

enhance proliferation of early germ cells while also preventing premature differentiation of Sertoli and mature germ cells at a time when pituitary FSH secretion is sufficient to promote Sertoli cell proliferation. In neonatal rats estradiol enhances FSH actions on Sertoli cell function but not their proliferation [25]. Normal Sertoli cell numbers are also observed in aromatase knockout (estrogen-deficient) mice [231]. A crucial role for combined neonatal FSH-estradiol activity in Sertoli cell remains elusive, because aromatase-deficient or estrogen receptor (ER) α-null mice are initially fertile, and ER β-null males are always fertile [231]. However, in vivo effects of estradiol require cautious interpretation, due to possible effects from dietary phytoestrogens [232]. In addition, FSH-mediated neonatal activities may combine with several factors, such as activin [233],

Chapter 11

In Vivo FSH Actions

thyroid hormone [47, 50], or TGFβ activity [234], to produce normal levels of Sertoli proliferation. It is therefore possible that neonatal aromatization acts to reduce local testosterone levels, or provide estradiol activity, to allow the FSH-mediated perinatal proliferation of Sertoli cells to reach its normal capacity. While estradiol levels decline, there is evidence that FSH regulates the temporal development of androgen receptor (AR) expression in Sertoli cells. In situ analysis revealed that very low AR expression at birth switches to markedly higher postnatal expression in both rat [235] and marmoset [236] Sertoli cells (see Chapter 12). Current knowledge of early postnatal AR regulation is limited to in vitro studies, which found that FSH increased the amount of AR mRNA or protein in Sertoli cells from 5- to 15-day-old [237, 238] or 20-day-old rats [239]. The temporal regulation of postnatal AR expression by FSH may provide a plausible mechanism for delaying the androgen receptiveness of developing Sertoli cells, thus avoiding premature differentiation as proposed in the E-T switch model. Low AR levels and higher aromatization of testosterone may both initially act to suppress the androgen sensitivity of perinatal Sertoli cells to allow completion of FSH mitogenic effects. For spermatogenesis, delayed androgen actions in Sertoli cells may be necessary to stall germ cell development, until Sertoli cells proliferate via the FSH response and gain the functional capacity to then support and nourish full germ cell differentiation. Androgen action may also initially promote FSH-induced Sertoli cell proliferation, as suggested by the enhanced Sertoli cell proliferation and differentiation by combined FSH and hCG treatment of juvenile rhesus monkeys [158]. The FSH-stimulated synthesis of estradiol by prepubertal Sertoli cells was also proposed to suppress Leydig cell androgen production, thereby stalling the initiation of spermatogenesis (reviewed in [240]). This model is not exclusive of the postnatal induction of Sertoli cell androgen sensitivity by FSH, and it is possible both activities merge to orchestrate the final FSHmediated effects on Sertoli cell development and ultimately adult function. It remains to be determined if the regulatory role of FSH in the proposed E-T switch involves independent or merging estradiol-mediated or AR-dependent pathways either early or later, respectively, in postnatal Sertoli cell development.

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220. Jannini, E. A., Ulisse, S., and D’Armiento, M. (1995). Thyroid hormone and male gonadal function. Endocr. Rev. 16(4), 443–459. 221. Coutant, R., Lumbroso, S., Rey, R., Lahlou, N., Venara, M., Rouleau, S., Sultan, C., and Limal, J. M. (2001). Macroorchidism due to autonomous hyperfunction of Sertoli cells and G(s)alpha gene mutation: An unusual expression of McCuneAlbright syndrome in a prepubertal boy. J. Clin. Endocrinol. Metab. 86(4), 1778–1781. 222. Benvenga, S., Smedile, G., Lo Giudice, F., and Trimarchi, F. (1999). Testicular adrenal rests: evidence for luteinizing hormone receptors and for distinct types of testicular nodules differing for their autonomization. Eur. J. Endocrinol. 141(3), 231–237. 223. Handelsman, D. J., Yue, D. K., and Turtle, J. R. (1983). Hypogonadism and massive testicular infiltration with amyloidosis. J. Urol. 129, 610–612. 224. Chitayat, D., Nakagawa, S., Marion, R. W., Sachs, G. S., Hahm, S. Y., and Goldman, H. S. (1988). Aspartylglycosaminuria in a Puerto Rican family: Additional features of a panethnic disorder. Am. J. Med. Genet. 31(3), 527–532. 225. Viljoen, D. L., Nelson, M. M., de Jong, G., and Beighton, P. (1987). Proteus syndrome in southern Africa: Natural history and clinical manifestations in six individuals. Am. J. Med. Genet. 27(1), 87–97. 226. Sripathi, V. (1999). Macro-orchidism may be an indicator of continuous splenogonadal fusion. BJU Int. 84(6), 733–734. 227. Slegtenhorst-Eegdeman, K. E., de Rooij, D. G., Verhoef-Post, M., van de Kant, H. J., Bakker, C. E., Oostra, B. A., Grootegoed, J. A., and Themmen, A. P. (1998). Macroorchidism in FMR1 knockout mice is caused by increased Sertoli cell proliferation during testicular development. Endocrinology 139(1), 156–162. 228. Carpino, A., Pezzi, V., Rago, V., Bilinska, B., and Ando, S. (2001). Immunolocalization of cytochrome P450 aromatase in rat testis during postnatal development. Tissue Cell 33(4), 349–353. 229. Rommerts, F. F., de Jong, F. H., Brinkmann, A. O., and van der Molen, H. J. (1982). Development and cellular localization of rat testicular aromatase activity. J. Reprod. Fertil. 65(2), 281–288. 230. Rosselli, M., and Skinner, M. K. (1992). Developmental regulation of Sertoli cell aromatase activity and plasminogen activator production by hormones, retinoids and the testicular paracrine factor, PModS. Biol. Reprod. 46(4), 586–594. 231. O’Donnell, L., Robertson, K. M., Jones, M. E., and Simpson, E. R. (2001). Estrogen and spermatogenesis. Endocr. Rev. 22(3), 289–318. 232. Robertson, K. M., O’Donnell, L., Simpson, E. R., and Jones, M. E. (2002). The phenotype of the aromatase knockout mouse reveals dietary phytoestrogens impact significantly on testis function. Endocrinology 143(8), 2913–2921. 233. Boitani, C., Stefanini, M., Fragale, A., and Morena, A. R. (1995). Activin stimulates Sertoli cell proliferation in a defined period of rat testis development. Endocrinology 136(12), 5438–5444. 234. Dorrington, J. H., Bendell, J. J., and Khan, S. A. (1993). Interactions between FSH, estradiol-17 beta and transforming growth factor-beta regulate growth and differentiation in the rat gonad. J. Steroid Biochem. Mol. Biol. 44(4–6), 441–447. 235. Bremner, W. J., Millar, M. R., Sharpe, R. M., and Saunders, P. T. (1994). Immunohistochemical localization of androgen receptors in the rat testis: Evidence for stage-dependent expression and regulation by androgens. Endocrinology 135(3), 1227–1234. 236. McKinnell, C., Saunders, P. T., Fraser, H. M., Kelnar, C. J., Kivlin, C., Morris, K. D., and Sharpe, R. M. (2001). Comparison of androgen receptor and oestrogen receptor beta immunoexpression in the testes of the common marmoset (Callithrix jacchus) from birth to adulthood: Low androgen receptor immunoexpression in Sertoli cells during the neonatal increase in testosterone concentrations. Reproduction 122(3), 419–429.

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237. Blok, L. J., Hoogerbrugge, J. W., Themmen, A. P., Baarends, W. M., Post, M., and Grootegoed, J. A. (1992). Transient down-regulation of androgen receptor messenger ribonucleic acid (mRNA) expression in Sertoli cells by follicle-stimulating hormone is followed by up-regulation of androgen receptor mRNA and protein. Endocrinology 131(3), 1343–1349. 238. Arambepola, N. K., Bunick, D., and Cooke, P. S. (1998). Thyroid hormone effects on androgen receptor messenger RNA expression in rat Sertoli and peritubular cells. J. Endocrinol. 156(1), 43–50. 239. Sanborn, B. M., Caston, L. A., Chang, C., Liao, S., Speller, R., Porter, L. D., and Ku, C. Y. (1991). Regulation of androgen receptor mRNA in rat Sertoli and peritubular cells. Biol. Reprod. 45(4), 634–641. 240. Abney, T. O. (1999). The potential roles of estrogens in regulating Leydig cell development and function: a review. Steroids 64(9), 610–617. 241. Meachem, S. J., Wreford, N. G., Stanton, P. G., Robertson, D. M., and McLachlan, R. I. (1998). Follicle-stimulating hormone is required for the initial phase of spermatogenic restoration in adult rats following gonadotropin suppression. J. Androl. 19(6), 725–735. 242. Sharpe, R. M., Atanassova, N., McKinnell, C., Parte, P., Turner, K. J., Fisher, J. S., Kerr, J. B., Groome, N. P., Macpherson, S., Millar, M. R., and Saunders, P. T. (1998). Abnormalities in functional development of the Sertoli cells in rats treated neonatally with diethylstilbestrol: A possible role for estrogens in Sertoli cell development. Biol. Reprod. 59(5), 1084–1094. 243. Atanassova, N., McKinnell, C., Walker, M., Turner, K. J., Fisher, J. S., Morley, M., Millar, M. R., Groome, N. P., and Sharpe, R. M. (1999). Permanent effects of neonatal estrogen exposure in rats on reproductive hormone levels, Sertoli cell number, and the efficiency of spermatogenesis in adulthood. Endocrinology 140(11), 5364–5373. 244. Remy, J. J., Couture, L., Rabesona, H., Haertle, T., and Salesse, R. (1996). Immunization against exon 1 decapeptides from the lutropin/choriogonadotropin receptor or the follitropin receptor as potential male contraceptive. J. Reprod. Immunol. 32(1), 37–54. 245. Medhamurthy, R., Aravindan, G. R., and Moudgal, N. R. (1993). Hemiorchidectomy leads to dramatic and immediate alterations in pituitary follicle-stimulating hormone secretion and the functional activity of the remaining testis in the adult male bonnet monkey (Macaca radiata). Biol. Reprod. 49(4), 743–749. 246. Marshall, G. R., Zorub, D. S., and Plant, T. M. (1995). Folliclestimulating hormone amplifies the population of differentiated spermatogonia in the hypophysectomized testosteronereplaced adult rhesus monkey (Macaca mulatta). Endocrinology 136(8), 3504–3511. 247. Lunn, S. F., Cowen, G. M., and Fraser, H. M. (1997). Blockade of the neonatal increase in testosterone by a GnRH antagonist: The free androgen index, reproductive capacity and postmortem findings in the male marmoset monkey. J. Endocrinol. 154(1), 125–131.

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248. Rabinowitz, D., Cohen, M. M., Rosenmann, E., Rosenmann, A., Segal, S., Bell, J., Rosler, A., and Spitz, I. (1975). Chromatinpositive Klinefelter’s syndrome with undetectable peripheral FSH levels. Am. J. Med. 59(4), 584–590. 249. Stewart-Bentley, M., and Wallack, M. (1975). Isolated FSH deficiency in a male. Clin. Res. 23:96A. 250. Maroulis, G. B., Parlow, A. F., and Marshall, J. R. (1977). Isolated follicle-stimulating hormone deficiency in man. Fertil. Steril. 28(8), 818–822. 251. McConnon, J., Killinger, D., Gracey, W., and Ghany, F. (1979). Clomiphene in treatment of male infertility due to isolated follicle-stimulating-hormone deficiency. Lancet 2(8141), 525–526. 252. Mozaffarian, G. A., Higley, M., and Paulsen, C. A. (1983). Clinical studies in an adult male patient with “isolated follicle stimulating hormone (FSH) deficiency.” J. Androl. 4(6), 393–398. 253. Al-Ansari, A. A., Khalil, T. H., Kelani, Y., and Mortimer, C. H. (1984). Isolated follicle-stimulating hormone deficiency in men: Successful long-term gonadotrophin therapy. Fertil. Steril. 42(4), 618–626. 254. Diez, J. J., Iglesias, P., Sastre, J., Salvador, J., Gomez-Pan, A., Otero, I., and Granizo, V. (1994). Isolated deficiency of folliclestimulating hormone in man: A case report and literature review. Int. J. Fertil. Menopausal Stud. 39(1), 26–31. 255. Tapanainen, J. S., Aittomaki, K., Min, J., Vaskivuo, T., and Huhtaniemi, I. T. (1997). Men homozygous for an inactivating mutation of the follicle-stimulating hormone (FSH) receptor gene present variable suppressions of spermatogenesis and fertility. Nat. Genet. 15(2), 205–206. 256. Phillip, M., Arbelle, J. E., Segev, Y., and Parvari, R. (1998). Male hypogonadism due to a mutation in the gene for the beta-subunit of follicle-stimulating hormone. N. Engl. J. Med. 338(24), 1729–1732. 257. Lindstedt, G., Nystrom, E., Matthews, C., Ernest, I., Janson, P. O., and Chatterjee, K. (1998). Follitropin (FSH) deficiency in an infertile male due to FSHbeta gene mutation. A syndrome of normal puberty and virilization but underdeveloped testicles with azoospermia, low FSH but high lutropin and normal serum testosterone concentrations. Clin. Chem. Lab. Med. 36(8), 663–665. 258. Singh, S. K. (2002). Isolated follicle-stimulating hormone (FSH) deficiency syndrome (IFDS). J. Assoc. Physicians India 50, 1458. 259. Layman, L. C., Porto, A.L., Xie, J., da Motta, L. A., da Motta, L. D., Weiser, W., and Sluss, P. M. (2002). FSHbeta gene mutations in a female with partial breast development and a male sibling with normal puberty and azoospermia. J. Clin. Endocrinol. Metab. 87(8), 3702–3707. 260. Mantovani, G., Borgato, S., Beck-Peccoz, P., Romil, R., Boretta, G., and Persani, L. (2003). Isolated follicle-stimulating hormone (FSH) deficiency in a young man with normal virilization who did not have mutations in the FSHbeta gene. Fertil. Steril. 79(2), 434–436.

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12 Sertoli Cell Endocrinology and Signal Transduction: Androgen Regulation RICHARD M. SHARPE MRC Human Reproductive Sciences Unit, Centre for Reproductive Biology, Edinburgh Scotland

Because it is the Sertoli cells that are in direct contact with the germ cells during their development into spermatozoa, it is logical to presume that the effects of androgens on spermatogenesis are mediated either by direct action on the germ cells and/or by effects on the supporting “nurse” Sertoli cells. Though some have argued that androgen receptors (ARs) are expressed in germ cells, it seems almost certain that this is an artifact and that no such expression occurs [1, 3, 4]. Perhaps the most definitive evidence for the lack of importance of AR-mediated androgen action within germ cells is the finding that mice that carry a defective copy of the AR in their germ cells are still able to produce offspring provided that the Sertoli cells that support them have a normal phenotype and are expressing a functional AR. This has been demonstrated in two ways: first, by using chimeric mice in which the germ cells, but not the Sertoli cells, have an X-chromosome bearing a nonfunctional copy of the AR-encoding gene [5] and, second, by transplanting germ cells with a defective AR-encoding gene into the seminiferous tubules of azoospermic mice that are expressing a functional AR [6]. However, although such observations have ruled out a role for direct AR-mediated androgen action on the germ cells, they do not resolve another long-lasting debate, and that is whether or not it is androgen action on the Sertoli cells or peritubular myoid cells that is most important in terms of androgen support for spermatogenesis [1]. Discerning the relative importance of these two cell types is extremely difficult because it is clear that they interact and coordinate their functions

I. INTRODUCTION II. ACQUISITION OF ANDROGEN RESPONSIVENESS (AR EXPRESSION) BY SERTOLI CELLS III. ANDROGENS AND SERTOLI CELL PROLIFERATION IV. EVIDENCE OF A ROLE FOR SERTOLI CELL–MEDIATED ANDROGEN ACTION ON SPERMATOGENESIS V. MECHANISMS INVOLVED IN MEDIATING ANDROGEN ACTION ON SPERMATOGENESIS VIA THE SERTOLI CELL VI. CONCLUSIONS AND FUTURE PROSPECTS References

I. INTRODUCTION Arguably, the most important unanswered question in andrology is “How do androgens regulate spermatogenesis?” It is remarkable that, despite the many advances in molecular and cellular understanding and technology, even today we still have almost no answer to this important question. This ignorance is frustrating because it has been well established for decades that androgens are all-important for the maintenance of normal spermatogenesis, and hence for male fertility [1]. It is also long established for both animals and humans that suppression of testosterone levels within the testis will result in suppression of spermatogenesis, loss of sperm production, and, hence, infertility. Indeed, this is being actively explored as a means via which to induce infertility in men for contraceptive purposes [2]. SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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in a fairly complex way [7–9]. It is perhaps most likely that the main effects of androgens in supporting spermatogenesis are mediated via the Sertoli cells because it is these cells that are in direct and intimate contact with all of the germ cells [9], whereas the peritubular cells only have close contact with the earliest germ cell types, the spermatogonia. It should be possible to more accurately define the relative importance of Sertoli and peritubular myoid cells in mediating androgen action on spermatogenesis now that a transgenic mouse has been developed in which lack of expression of AR specifically in the Sertoli cells has been engineered. Probably the most important role of peritubular myoid cells in spermatogenesis is their role in early development and formation of the seminiferous cords, because this is a prerequisite for normal spermatogenesis in adulthood. During fetal life, the AR is not expressed by Sertoli cells, whereas there is strong expression of the AR in the peritubular myoid cells. Therefore, although lack of normal androgen action during fetal development can result in major problems with spermatogenesis in adulthood, these deficits do not result from a lack of direct androgen action on the Sertoli cells at these times. Instead, they probably result from a cascade of changes that lead to abnormal development of one or more of the testicular cell types, including the Sertoli cells. As should be apparent from the preceding comments, many fundamental aspects relating to androgen action on the Sertoli cell and hence on spermatogenesis remain to be resolved. The incentive to work in this area and to obtain new insights is considerable because such information could potentially revolutionize our ability to manipulate androgen action on spermatogenesis. This chapter reviews our current understanding of how and when androgens act on Sertoli cells and how this may support spermatogenesis; it also identifies the important areas in which our ignorance hinders progress.

II. ACQUISITION OF ANDROGEN RESPONSIVENESS (AR EXPRESSION) BY SERTOLI CELLS As already indicated, AR expression by Sertoli cells is not evident in fetal life in any of the species that have been studied, as shown in Figure 12.1 for the rat, marmoset, and human. Indeed, from studies of animals in which there are naturally occurring mutations in the AR that prevent it from being active, it is clear that formation of the Sertoli cells and of the testis usually occurs grossly normally and that major problems only

begin to arise in postnatal life, particularly entering puberty. Because the AR responsiveness of Sertoli cells is clearly essential for the support of spermatogenesis, it might be anticipated that the first appearance of AR in Sertoli cells occurs at puberty. In practice, it appears that it is not this simple (Fig. 12.1). It is evident that when comparing rodents such as the rat with nonhuman primates and the human, there is agreement in each of these that fetal Sertoli cells do not express AR (Fig. 12.1). However, at what point each of these species switches on AR expression is markedly different. In laboratory rodents, AR expression is switched on in Sertoli cells in the early postnatal period, some days before the initiation of true puberty; thus, in the rat, weak AR expression is first detectable immunohistochemically at around days 4–6 postnatal (Fig. 12.1) [3]. In contrast, in the marmoset during the neonatal period, when testosterone levels are high (the so-called “neonatal testosterone surge”), AR expression in Sertoli cells is still absent or barely apparent, yet when the neonatal testosterone surge has finished, AR expression is then switched on [10, 11] (Fig. 12.1). In this case, onset of AR expression is thus initiated many months before puberty and at a time when the testis is quiescent—this would equate to the period of childhood infancy in the human [10, 12]. Sertoli cell AR expression in the human does not switch on at the same time as in the marmoset despite the fact that the human, like all primates, also shows a neonatal testosterone surge followed by a “childhood” period. Instead, AR expression by Sertoli cells in the human appears to occur as a relatively late event during puberty [13], though the precise timing of this appearance remains to be established. Though it is certain that androgen action on Sertoli cells in adulthood plays an important role in the support of spermatogenesis in the rat, marmoset, and human, it is difficult to discern a unifying mechanism or purpose for the different timing of appearance of AR expression in the Sertoli cells in these species. It seems reasonable to conclude, however, that onset of AR expression is not coincident with onset of spermatogenesis, at least in the rat and marmoset. This may indicate that there are functions of androgen action on the Sertoli cell other than those related to spermatogenesis, or that onset of AR expression is related to other stages of development of the Sertoli cell that differ between these different species. An important question is “What regulates the onset of AR expression in Sertoli cells?” No information is available for the human or the marmoset, but several factors that can modulate AR expression in Sertoli cells in the rat have been identified. For example, in vitro studies have shown that follicle-stimulating hormone (FSH) and thyroid hormone both induce AR

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FIGURE 12.1 Age of onset of androgen receptor (AR) immunoexpression in Sertoli cells of the rat, marmoset, and human. Note the absence of AR immunoexpression in Sertoli cells in seminiferous cords (asterisks) in the fetal/newborn testes of all three species. Sertoli cell AR expression then switches on in the rat just prior to puberty, during infancy (marmoset), or probably late in puberty (human; not shown, see text). Scale bars show 100 μm (see color plate).

expression in immature Sertoli cells [14]. In this situation, the effects of FSH and thyroid hormone are additive. Because both FSH and thyroid hormone levels are changing during the neonatal and prepubertal period in the rat, it can be envisaged that these hormones,

working together, are able to induce progressively greater expression of AR as adulthood approaches. This would fit with the evidence for an age-dependent increase in the expression level of AR protein in the rat Sertoli cell [3, 11].

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Other evidence suggests that there may also be altered sensitivity of Sertoli cells to thyroid hormone action during the neonatal period, as a result of change in expression of thyroid hormone receptors [15]. More recently it has been shown that thyroid hormone and testosterone play a role in suppressing proliferation and inducing terminal maturation of rat Sertoli cells in vitro [16]. These findings can be interpreted in various ways, but the overall indication is that the process of Sertoli cell maturation and AR acquisition are not coincidental events and that AR expression is simply one step in a cascade of changes that culminates in Sertoli cell maturation and, thus, ensures its ability to support spermatogenesis. In species such as the marmoset and human, in which the onset of spermatogenesis shows a considerable delay until after the period of childhood, it may be that differences in expression of factors such as thyroid hormone receptors and/or thyroid hormone levels may alter the timing of AR acquisition and final Sertoli cell maturation [11, 17]. These various findings imply that there is no consistent alignment of AR expression with either cessation of Sertoli cell proliferation or their terminal differentiation/maturation.

A. Stage-Dependent Expression of the AR in Sertoli Cells In the rat, various lines of evidence point to preferential action of androgens at around stages VII and VIII of the spermatogenic cycle [1, 18–22] and this coincides with maximum immunoexpression of the AR in Sertoli cells [3]. Similar stage-specific expression of the AR has also been documented for the human [23], although this is more difficult to demonstrate than in the rat. In contrast to this variation in Sertoli cell AR expression, immunoexpression of AR in peritubular myoid cells shows no obvious variation according to the stage of the spermatogenic cycle [3, 23]. This is one of the observations that indirectly suggests that androgen action on Sertoli cells, rather than on peritubular myoid cells, plays the most important role in supporting spermatogenesis [1]. An important question, then, is “What regulates the stage-dependent expression of AR in the Sertoli cells?” In most situations in which a stage-dependent change in Sertoli cell function has been described, the regulating factor has turned out to be the presence or absence of one or more germ cell types [9, 24]. Numerous such examples exist, and these have been elucidated either by undertaking Sertoli cell + germ cell recombination experiments in vitro or by selectively depleting particular germ cell types in vivo using chemicalinduced ablation [24, 25]. However, the latter approach has failed to reveal any major impact of the germ cell

complement in regulating stage-dependent immunoexpression of the AR in the rat [3]. Interestingly, a recent study [26] has shown that acute depletion of specific spermatocytes in the rat may result in altered mRNA for the AR in the testis, though such studies did not distinguish whether this altered expression was in Sertoli cells or in other AR-expressing cells such as the peritubular myoid and Leydig cells. Nor was it shown that this change in mRNA expression was matched by a change in AR protein expression, because there is growing evidence that post-translational regulation of the AR (and other steroid receptors) is perhaps the most important mechanism for regulating receptor bioavailability [27]. Two recent studies suggest that isoforms of the Wilms tumor gene protein (WT1) can bind to the promoter region of the AR and suppress its transcription [28, 29]. Because WT1 is expressed in Sertoli cells very early during differentiation of the testis, it may be that the expression of a particular isoform of WT1 plays a role in determining the onset of AR expression by Sertoli cells; however, this has not yet been addressed by published studies. Similarly, the possibility that expression of isoforms of WT1 might play a role in regulation of the stage-dependent expression of the AR in Sertoli cells in the adult testis has yet to be explored. Another recent study [30] has demonstrated that in testes from humans in which various disorders may be present, immunoexpression of the AR in Sertoli cells can be related to the degree of maturation of the Sertoli cells. In particular, in cryptorchid testes in which there was no spermatogenesis and in which the Sertoli cells appeared morphologically immature, no immunostaining of the AR was present or there was only very weak staining in certain areas. The authors of the study concluded that the focal absence of AR expression in Sertoli cells correlated with a lack of local spermatogenesis in the same tubules. At face value, this finding might suggest that no degree of spermatogenesis is possible in the absence of AR expression in Sertoli cells. However, studies of transgenic mice in which AR expression in the Sertoli cell has been selectively knocked out (SCARKO mice) have shown that, in the complete absence of AR expression, spermatogenesis up to meiosis occurs (see later discussion). Finally, although AR expression in Sertoli cells is clearly of fundamental importance in mediating the effects of androgens on spermatogenesis (see later discussion), the possibility of non-genomic effects of androgens (i.e., that occur independently of whether or not the AR is expressed and binds to DNA) have also to be entertained, because evidence for such effects is increasing (see later discussion).

Chapter 12 Sertoli Cell Endocrinology and Signal Transduction: Androgen Regulation

III. ANDROGENS AND SERTOLI CELL PROLIFERATION The generally accepted view is that hormonal regulation of Sertoli cell proliferation is mediated primarily via the actions of FSH [11]. However, recent evidence has raised the possibility of a role for androgens in this process. First, based on studies of three transgenic/ mutant mouse lines (FSH knockout, FSH receptor knockout, testicular feminized male mouse), only the AR-deficient testicular feminized male mouse was shown to have reduced Sertoli cell numbers at birth, when compared with wild-type mice, indicating that androgen action, rather than FSH action, is important for Sertoli cell proliferation in fetal life [31]. The same study showed that, postnatally, FSH was important in determining Sertoli cell number, consistent with numerous other studies [11], although a role for androgens was still suggested [31]. Evidence from juvenile rhesus monkeys also supports a role for luteinizing hormone (LH)/testosterone in stimulating Sertoli cell proliferation prepubertally [32]. Conversely, studies using cultured Sertoli cells from neonatal rats have indicated that testosterone may inhibit Sertoli cell proliferation, though this effect is probably attributable to accelerated Sertoli cell maturation [11, 16]. The finding that androgens may play a role in Sertoli cell proliferation in the fetal testis in mice is distinctly at odds with the clear lack of AR expression in Sertoli cells at this age (Fig. 12.1). Moreover, analysis of Sertoli cell volume/number per testis in adult SCARKO mice, in which specific depletion of AR from the Sertoli cells has been engineered, has shown no difference from wild-type controls, despite a considerable reduction in testis size (see later discussion and [33]). The implication of these observations is that any effect of androgens on Sertoli cell proliferation in fetal life may be indirect, most likely mediated via effects on the peritubular myoid cells.

IV. EVIDENCE OF A ROLE FOR SERTOLI CELL–MEDIATED ANDROGEN ACTION ON SPERMATOGENESIS Throughout the years, various approaches have been used to study androgen action and the regulation of spermatogenesis. In all but the most recent studies, these have been unable to distinguish whether or not the effects of androgen withdrawal or replacement are mediated by the Sertoli cells, the peritubular myoid cells, or by a combination of effects on these two cell types. This is because the majority of the relevant studies

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have been in vivo and it is therefore not possible to make a definitive distinction. Before considering the approaches that have been used, it is worthwhile to first of all consider some of the important factors that potentially confound the results obtained in such studies. Of these, the most important is the complement of germ cells that is present. Because germ cells make up the vast majority (~90%) of the adult seminiferous epithelium, when compared to the contribution made by the Sertoli cells or peritubular myoid cells, even a small change in the number of germ cells present is likely to have a big influence on any endpoints being studied, particularly if these studies are searching for changes in gene or protein expression. Not only is the absence or loss of germ cells important for this reason, but also because it is well established that alteration of the germ cell complement will alter feedback to the Sertoli cells, which can result in major changes in Sertoli cell gene and protein expression (9, 24, 25). Because loss of germ cells after complete testosterone withdrawal begins within 3–4 days, it is very important that studies take into account such changes and that studies are designed so as to minimize the confounding effects that any change in the germ cell complement will have [1]. Such changes are probably of most importance when searching for changes in gene or protein expression and are of less importance when studies have been directed at an overall evaluation of spermatogenesis (i.e., maintenance of a normal germ cell complement). Numerous approaches to the study of androgen action on spermatogenesis have been used by different researchers [1], but the three most commonly used (all in rats) have been as follows: 1. Use of hypophysectomized rats to deprive Leydig cells of endogenous LH stimulation and thus to suppress testosterone production. Treatment of such animals with exogenous testosterone can then be used to study its role. 2. Destruction of the endogenous Leydig cell population by administration of the toxicant EDS (ethane dimethane sulfonate). A single injection of EDS into adult rats is able to eliminate all Leydig cells within about 36 hr of treatment and represents the most effective way of inducing complete testosterone withdrawal [1]. Exogenous administration of testosterone to EDS-treated rats can then be used to study the role of androgen action. 3. Suppression of endogenous gonadotropin levels and thus endogenous testosterone production by the use of GnRH analogues. This treatment has been used most effectively in combination with implants of both testosterone and estradiol.

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Irrespective of the method of study that has been used, virtually all studies are in agreement over certain central features. Thus, after testosterone withdrawal, stage-dependent apoptosis of particular germ cell types (mainly pachytene spermatocytes and round spermatids) occurs, with stages VII and VIII being the only stages initially affected in the rat. The fact that these stages are those at which expression of the AR in Sertoli cells is at its highest [3] provides the most compelling piece of evidence that these changes indicate a selective response to withdrawal of androgen action on the Sertoli cells. Perhaps the biggest difference between results obtained using the three different approaches just mentioned is the earliest day at which abnormal apoptosis of germ cells first becomes evident. This difference is due primarily to the degree of testosterone withdrawal achieved by the different treatment methods [1]. For example, although both hypophysectomy and gonadotropin-releasing hormone (GnRH) analogue treatment deprive the Leydig cells of the LH stimulus that they need to maintain testosterone production, the degree of suppression of steroidogenesis is incomplete and gradually subsides over 1–2 wk after initial treatment, such that testosterone levels within the testis can remain at 5–10% of normal levels even 1–2 wk later [1, 22]. In contrast, use of EDS rapidly (~ 36 hr) reduces testosterone levels to undetectable levels because it eliminates all of the Leydig cells [34, 35]. However, EDS is an alkylating agent and may therefore have other effects within the testis than those that result in the elimination of the Leydig cells, although no such effects have yet been described. The main complication of using the EDS approach is that regeneration of Leydig cells commences some 10–14 days after initial treatment and testosterone levels then gradually return to normal over the next week or two [35]. The approach involving combined administration of GnRH analogues to rats plus testosterone and estradiol implants, although soundly based at the time, now appears to be somewhat compromised by recent developments that have shown that Sertoli cells and most germ cells are potentially direct estrogen targets. This has resulted from the discovery that the second estrogen receptor, ERβ, is expressed in these different cell types [10, 36–38]. Therefore, as well as reinforcing negative feedback suppression of gonadotropins and direct suppression of Leydig cell function, the administered estradiol may directly have affected the Sertoli and/or germ cells [38]. This might explain the slightly different effects obtained using this approach, when compared to others, especially with regard to the effects on round spermatid attachment to Sertoli cells [39]; recent studies suggest direct actions of estrogens on

round spermatids [38, 40, 41]. Results from studies addressing testosterone action on spermatogenesis in which estradiol implants have been used therefore need to be considered with this knowledge in mind. Only selective germ cell types are effected initially by androgen withdrawal, and this occurs in a stagespecific manner, but it is still debatable whether this indicates that androgen action on the Sertoli cell is directed at functions that involve just these particular cell types. It is equally likely that there is a generalized, stage-specific effect of androgens on Sertoli cells and that the germ cells initially effected are simply those that are most dependent on the Sertoli cell because of their energy or nutrient requirements. Several studies (e.g., [42, 43]) have suggested that testosterone may play a stage-specific role in regulating the attachment of round spermatids to the Sertoli cell at stages VII and VIII of the spermatogenic cycle in the rat; when testosterone action was withdrawn experimentally, premature sloughing of spermatids into the lumen of the seminiferous tubule was induced. Further studies went on to explore what cell adhesion molecules (CAMs) might be involved in mediating this apparent effect of testosterone. These demonstrated that one particular CAM, N-cadherin, was dose dependently regulated in isolated immature Sertoli cells by testosterone, although only in the presence of FSH [44, 45]. These studies also showed that upregulation of N-cadherin was associated with increased round spermatid attachment to the Sertoli cells. However, when the same experiments were undertaken using isolated Sertoli cells from mature rats, no effect of testosterone ± FSH on N-cadherin expression was detectable [45]. Subsequent studies have gone on to demonstrate that mRNAs for various other CAMs are all significantly upregulated by testosterone replacement in vivo using the rat model that employs the T + E implants referred to earlier [39]. However, in this situation it is impossible to distinguish whether the increase in CAMs expression is a primary response to testosterone or merely reflects the increased numbers of spermatids attached to the Sertoli cells due to other (supportive) effects of testosterone (e.g., [46]). Despite these doubts, it is interesting that expression of N-cadherin in motoneurons is clearly androgen regulated [47]. For many researchers unfamiliar with the testis, the problem just highlighted, namely, the distinction between cause and consequence when studying the effects of testosterone withdrawal and replacement, is a major confounding issue. Unless extreme care is taken, very misleading conclusions can be drawn, which then result in wasted research time and effort. The literature

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is littered with such examples. In essence, the problem centers on the fact that germ cells have very specialized cell–cell contacts with Sertoli cells that are essential for two-way signaling and for germ cell survival [9, 48, 49]. Clearly, a germ cell may detach from the Sertoli cell if the latter ceases to express a CAM necessary for germ cell adhesion, and this could result from withdrawal of testosterone action; this would be a primary effect of testosterone. On the other hand, any effect on the testis that results in germ cell detachment from the Sertoli cell—whether this results from an effect on the Sertoli cell or on the germ cell itself—may secondarily result in loss of expression of CAM molecules, by either the germ cell, Sertoli cell, or both cell types, because of the loss of cell–cell signaling [9, 24]. In this instance, the effect on the CAM in question is secondary, not primary. It can be appreciated that distinguishing between primary and secondary effects can be extremely difficult (e.g., [46]), and this undoubtedly represents one of the major obstacles facing researchers who are trying to identify the mechanisms via which androgens support spermatogenesis.

A. Effects of AR Mutations/Knockouts or GnRH Mutations (Hypogonadal Mouse) Animal models, in which naturally occurring mutations of the AR or of upstream regulatory pathways (e.g., hypogonadal mouse, with an inactivating GnRH mutation) have been described in some detail, have provided insights into the importance of androgen action in spermatogenesis. In these situations, little or no spermatogenesis is evident in adulthood, although there is normal formation and development of the testis up to the period around birth [50]. The naturally occurring AR mutation in rats and mice gives rise to a testicular feminized male (tfm) phenotype in which AR-mediated androgen action is prevented [51, 52]. Although spermatogenesis fails in tfm males [51, 53, 54], other important changes also occur that confound the straightforward interpretation of such findings. Primarily this involves the failure of masculinization of other component parts of the reproductive system; in particular, impairment of scrotum development and the inguinal phase of testicular descent. Because the latter two events are both impaired in the tfm rat, the result is that the testis remains incompletely descended (cryptorchid) in 85% of animals [31, 55] with consequent impairment of spermatogenesis because of the elevated testicular temperature. This means that it is impossible in this situation to discern the contribution of the inactive AR versus the position of the testis, although use of “cryptorchid controls” can provide some means of distinction [31].

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Another naturally occurring model, in which the compromise of cryptorchidism also occurs, is the hypogonadal (hpg) mouse. In this model, the deficit is in the production of GnRH and, as a consequence, there is failure of normal gonadotropin secretion and consequent absence of normal testosterone production by the postnatal testis. However, because production of testosterone by the fetal Leydig cells is gonadotropin independent in rodents, the absence of LH secretion does not result in apparent deficit in testosterone production in fetal life [56]. As a consequence, normal development of the reproductive tract, including the scrotum, occurs. However, the testes do not descend normally due to the lack of postnatal production of testosterone. However, if hpg mice are treated with gonadotropins or testosterone, their testes will descend normally into the scrotum [57]. Therefore in adult hpg males, spermatogenesis does not occur, though it can be induced by manipulating hormone levels exogenously by various hormonal treatments. These have included detailed studies in which the relative importance and interactions of FSH and testosterone have been evaluated (e.g., [58, 59]). The conclusions from these studies are generally in line with those obtained in the rat models described earlier, namely, that testosterone action is essential for completion of meiosis and spermiogenesis and that interactive effects with FSH may occur. Use of the hpg mouse provides an alternative and, in many respects, a better model for studying hormonal control of spermatogenesis than the models described above that are based on experimental manipulations in the rat. Nevertheless, the hpg mouse still suffers from the same problems as these other models in that it is impossible to distinguish between androgen affects on Sertoli cells or on peritubular myoid cells. More recently, mice in which the AR has been inactivated by transgenesis have been generated [33, 60]. The phenotype of these animals is identical to that for the animal models in which there is a naturally occurring mutation in the AR (i.e., the tfm rat and mouse). Thus, there is failure of masculinization, variable impairment of scrotum formation, and failure of normal testicular descent and gross impairment of spermatogenesis in adulthood. So again, although such models are of considerable usefulness for particular studies, they are not of much use for studying the specific actions of androgens on Sertoli cells. These problems have now been overcome by the development of a selective knockout of the AR in Sertoli cells, referred to as the SCARKO mouse. The SCARKO mouse was generated using Cre-lox technology, namely, by mating female mice, in which the AR had been floxed, to male mice that were

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heterozygous for an AMH-Cre transgene [33]. In the resulting SCARKO males, normal masculinization occurs, including development of a normal scrotum and normal descent of the testis into the scrotum. However, appearance of the AR in Sertoli cells in the neonatal period fails to occur in SCARKO mice and remains permanently absent throughout adult life. In contrast, normal expression of the AR is apparent in both peritubular myoid cells and Leydig cells and in arterioles in the testis (Fig. 12.2). Based on standard cell-counting methods, apparently normal numbers of Sertoli cells and Leydig cells develop in SCARKO mice and they also exhibit normal onset of spermatogenesis. However, in adulthood testis size is reduced by ~70% and spermatogenesis does not proceed beyond the spermatocyte stage; there is a 36% decrease in number of spermatocytes and a 97% decrease in numbers of round spermatids per testis, whereas the numbers of spermatogonia remain normal [33] (Figs. 12.2 and 12.3). In addition, the SCARKO mice show a massive decrease in volume of the seminiferous tubule lumen, indicative of dysfunction of fluid secretion by the Sertoli cells (Fig. 12.3). Because the production

of seminiferous tubule fluid (STF) by the Sertoli cell is an androgen-dependent process, this change is not surprising and further affirms the functional loss of the AR. In some respects it is perhaps surprising that spermatogenesis proceeds up to the early spermatocyte stage in a relatively ordinary manner in SCARKO mice, at least as far as this has been studied [33]. This may indicate that these earlier steps in spermatogenesis are normally driven by the effects of FSH on the Sertoli cell, because levels of FSH are slightly elevated in the SCARKO mice [33]. Alternatively, this observation may be an indication of androgen effects on peritubular myoid cells, which then influence Sertoli cell function to aid the maintenance of the earlier steps in spermatogenesis. In comparing what happens to spermatogenesis in the SCARKO mice with what has been observed previously in animal models in which androgen levels have been manipulated by Leydig cell destruction, it is apparent that there is some general agreement between these observations. For example, all of these studies have identified that midpachytene primary spermatocytes are negatively affected by the

FIGURE 12.2 Testicular histology and androgen receptor (AR) immunoexpression in adulthood in wildtype mice, animals in which AR expression has been completed ablated (ARKO) and mice in which Sertoli cell–specific ablation of the AR (SCARKO) has been engineered. Large arrows, Sertolic cell nuclei; small arrows, peritubular myoid cell nuclei; LC, Leydig cells. Scale bar shows 100 μm (see color plate). (Photomicrographs courtesy of collaborative studies with de Gendt et al. [33].)

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particular cell types; spermatids appear to be the most important of germ cells in this respect [9, 24].

V. MECHANISMS INVOLVED IN MEDIATING ANDROGEN ACTION ON SPERMATOGENESIS VIA THE SERTOLI CELL

FIGURE 12.3 Testicular cell composition and seminiferous tubule lumen volume in adulthood in wild-type mice (WT) and mice in which Sertoli cell–specific ablation of the AR (SCARKO) has been engineered. Values are means ± SEM for N = 5 (wild-type) or N = 3 (SCARKO). * p < .05, in comparison with appropriate value for WT. (Adapted from de Gendt et al. [33].)

absence of testosterone, as also are round spermatids (see earlier discussion) [1, 39]. It remains to be seen whether or not the changes in germ cell complement observed in adult SCARKO males show any evidence of stage dependence in the absence of a functional AR; absence of Sertoli cell AR expression could mean that the normal stage-dependent differences in the germ cell complement are blurred, although preliminary evidence does not support this possibility [33]. Future studies in SCARKO males will also aid considerably in defining whether or not androgen action via the AR is important for normal Sertoli cell maturation, although the finding that spermatogenesis proceeds into mid- to late meiosis most likely indicates that Sertoli cell maturation has occurred normally [11]. Another potentially important role for SCARKO males will be in studying the relative importance of nongenomic effects of androgens on Sertoli cells (see later discussion) versus the importance of those that are mediated via the AR. Of most potential importance would be utilization of SCARKO males to determine the cellular mechanisms that are normally driven by androgens, because it is these for which our ignorance is most complete. However, this may not be a straightforward process. Because SCARKO males show absence of particular germ cell types, in particular of virtually all spermatids, it is to be expected that there will be secondary changes in Sertoli cell function that result from the absence of signals from these

Despite the compelling evidence for androgen effects on Sertoli cells being responsible for regulation of spermatogenesis, it is remarkable that examples of androgen-regulated processes or gene/protein expression are almost nonexistent. In our own experience, we have applied each new technological development in our attempts to identify such changes, that is, subtraction hybridization, differential display, and use of microarray technology ([61, 62] and K. Tan, K. Turner, R. M. Sharpe, and P. T. K. Saunders, unpublished data). Though these studies identified some potential leads, we have been unable to confirm the absolute androgen dependence of any of the initially identified genes (K. Tan, K. J. Turner, R. M. Sharpe, and P. T. K. Saunders, unpublished data). The fact that no other studies have come forward in the past 5 years to identify androgen-regulated gene expression in Sertoli cells using similar approaches is a further demonstration of the frustrating nature of this area. The one notable exception to this situation is the identification of androgen-dependent expression of the Pem homeobox gene [63]. The Pem gene is expressed in adult Sertoli cells in both the mouse and rat in a stage-specific manner with expression encompassing the androgen-dependent phases of stages VII and VIII. Initial studies showed that Pem gene expression was dependent on both androgen and gonadotropin stimulation, as determined by studies using hypophysectomized mice, hpg mice and tfm mice [63, 64], and, most recently, SCARKO mice [33]. These encouraging findings prompted the generation of transgenic mice in which the Pem gene was inactivated [65]. However, these mice showed normal development and grossly normal spermatogenesis and fertility, suggesting that whatever the role and importance of Pem, it can be substituted by other factors. Just recently, generation of another transgenic line in which Sertoli cell expression of Pem occurs at all stages of the spermatogenic cycle has revealed a potential role for Pem in regulating pre-meiotic DNA regulation and/or chromatin remodeling in adjacent germ cells [66]. This perhaps suggests that the role of Pem is connected to the entry of germ cells into meiosis, an event that occurs during the androgen-dependent phase of the spermatogenic cycle [1]. In this particular

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transgenic line, increased DNA strand breaks were detected in preleptotene spermatocytes, but despite these changes there was no overall change in spermatogenesis or in fertility of the animals. These new observations perhaps suggest that the role of Pem is more directed toward the initiation of meiosis in germ cells than to more generalized aspects of spermatogenesis, and if this is the case then it suggests it may not play a critical role in mediating the more widespread effects of androgens on the Sertoli cell to support spermatogenesis. Apart from the identification of Pem, only bits and pieces of information are available in the literature that indicate an effect of testosterone on Sertoli cell expression of specific genes, or in which gene expression in germ cells has been altered via a presumed effect on Sertoli cells. An example of the latter is the studies on germ cell adherence to Sertoli cells and the role of cell adhesion molecules referred to earlier in this chapter [39]. Similarly, a series of studies have indicated that the expression of claudins, in particular claudin-1 and claudin-11, may be regulated in Sertoli cells by testosterone [67, 68], though studies in tfm mice and FSHβ knockout mice indicate no role for androgens, but a role for FSH, in regulation of claudin-1 [31]. Claudins play a role in formation and maintenance of the blood– testis barrier (i.e., the inter-Sertoli cell tight junctions) and it is of interest that morphological studies have shown that withdrawal of testosterone leads to aberrant function of these junctions [1, 69]. Other studies have used cultures of isolated Sertoli cells, predominantly cultures of immature Sertoli cells from either rats or from pigs, and have demonstrated the effects of adding testosterone on the expression of selected genes. Examples are TGFβ-1 [70], glutathione S-transferase alpha [71], and the proto-oncogene c-myc [72]. A number of earlier studies have also identified other factors using the same approaches and these have been reviewed elsewhere [1]. One of these was androgen-binding protein (ABP), which in vitro and in vivo evidence have suggested was regulated by FSH and testosterone [73]. Recent studies of mice overexpressing ABP in Sertoli cells have also noted impairment of spermatogenesis with features similar to those of androgen deprivation [74, 75]. However, secretion of ABP by Sertoli cells is also profoundly influenced by germ cells, especially by spermatids [9, 24], which again means that changes in ABP expression induced by germ cell changes have to be very clearly separated from direct hormonal regulation of ABP expression in Sertoli cells, and this is extremely difficult to do for reasons already discussed. Disappointingly, for none of the potential “androgenregulated” factors referred to earlier has a convincing case been made for them playing a fundamental role in

mediating Sertoli cell support for spermatogenesis as a primary effect of androgens. The most persuasive case can be made for Sertoli-spermatid adhesion molecules, but even in this instance there are doubts as to whether these are primary or secondary effects and/or whether estrogen effects might also be involved, as has already been discussed. It is remarkable that so few androgenregulated genes/proteins have been identified, although it is perhaps understandable that few have been identified using cultures of immature Sertoli cells, which are primarily FSH regulated rather than androgen regulated (reviewed in [1]). As discussed earlier, studies in vivo in which androgen levels are manipulated and in which changes in gene/protein expression are then studied are plagued by the problem of altered germ cell complement; wherever significant loss of germ cells occurs, the loss of mRNA/protein from these cell types will have a confounding effect. Because Sertoli cells make up a minor cellular component of the seminiferous epithelium of the normal adult testis, searching for changes in gene/protein expression in isolated seminiferous tubules or in whole testes is very much like looking for a “needle in a haystack.” Yet it is equally clear that as soon as the germ cells are removed to facilitate relative enrichment of Sertoli cells, then the function of the Sertoli cells changes so dramatically that their response to hormones, including to androgens, appears to be severely compromised [1, 24].

A. Production of Seminiferous Tubule Fluid Sertoli cells secrete the fluid termed STF, the production of which is hormonally regulated. During development and puberty, production of STF is driven primarily by the effects of FSH on the Sertoli cell [1, 76–78], but starting during puberty and very much evident during adulthood, the production of STF becomes predominantly controlled by testosterone [78–80]. STF has at least two functions, the most obvious being to transport the released spermatozoa out of the testis and into the epididymis [76]. Perhaps the less understood function of STF is to deliver general nutrients to the germ cells of the seminiferous epithelium [1, 76]. STF production can only be measured directly by using unilateral efferent duct ligation, which is a rather invasive method, although simpler approaches appear to give valid results [81, 82]. However, judging by the size of the lumen of seminiferous tubules in the adult testis, it appears that STF production is probably also stage dependent, with production being greatest at the androgen-dependent stages VII and VIII [78, 82]. Thus the size of the lumen at stages VII to VIII increases by about 50% compared to stage VI and then

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returns to the smaller level during stage IX after sperm release [82, 83]. Within 5–6 days of androgen withdrawal, this stage-dependent difference in lumen size is abolished, but can be maintained or restored by the administration of appropriate levels of testosterone [82]. It is also noteworthy that the normal androgendriven increase in seminiferous tubule lumen size fails to occur after depletion of elongate spermatids [83]. This provides a very clear demonstration of the complexity of germ cell modulation of androgen action on Sertoli cells. Though there are likely mechanisms via which Sertoli cell production of STF may occur, based on physiological studies of the testis and of other fluid-secreting cell types in the body [76], exactly what components are regulated by androgens (or by germ cells) remain unknown. When androgens are withdrawn, effects on STF production are not immediate but are delayed for several days (beyond 4 days) [82]. This probably rules out the possibility that STF production is regulated by the rapid (non-genomic) effects of androgens on the Sertoli cell, such as those involved in regulating calcium flux (see later discussion). The mechanisms via which STF production is regulated by androgens is an obvious target for study in SCARKO mice because these clearly show a deficit in STF production based on seminiferous tubule lumen size (Fig. 12.3).

B. Protein Synthesis and Secretion by Seminiferous Tubules/Sertoli Cells There is good evidence in the literature to demonstrate that one of the key effects of androgens at stages VII and VIII of the spermatogenic cycle is to regulate overall protein secretion. Thus, if seminiferous tubules are isolated from adult rats according to stage groupings and are cultured in vitro with 35S-methionine, it can be shown that overall protein secretion approximately doubles during the androgen-dependent stages of the spermatogenic cycle when compared with stages immediately preceding and following these stages [1, 20]. If androgen withdrawal is induced by EDS treatment prior to isolation of the seminiferous tubules, the stage-dependent increase in protein secretion can be completely abolished within 3–4 days of androgen withdrawal [1, 20]. Similarly, androgen replacement in EDS-treated rats is able to maintain the normal stagedependent pattern of protein secretion. The effect of androgen withdrawal on protein secretion under these circumstances appears to be unrelated to any change in germ cell complement because the maximal effects are observed after 4 days of androgen withdrawal, when the germ cell complement is virtually normal [20]. However, it is equally evident that if androgen levels

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are maintained at normal and the germ cell complement at stages VII and VIII is experimentally manipulated by other means, then the normal stage-dependent increase in protein secretion can be attenuated [84, 85]. In these studies, there was no major change in overall protein synthesis, which also shows little in the way of stage dependence or of androgen modulation [20, 85, 86]. Proteomics were used to establish whether or not the secretion of specific proteins was altered under these experimental conditions, and several candidate “androgen-regulated proteins” were identified [1, 20, 85]. Although some of these proteins were identified, it remains unclear as yet whether or not the altered secretion of these proteins is as a result of transcriptional modulation or reflects some other factor such as altered packaging and secretion of proteins in general [86]. Though the approach of using isolated seminiferous tubules and monitoring protein secretion using proteomics has proved rewarding and has given significant insights into how androgens may regulate seminiferous tubule function, it has been far from an ideal experimental situation. For example, there is still the problem that any changes that are observed may reflect effects both on Sertoli cells (which are a minor cellular component of the tubules) or on the germ cells (via nongenomic mechanisms or via conversion to estrogens). Moreover, even though only a short-term (24 hr) culture was used for such studies, significant degeneration of some of the germ cells occurs during this culture period, which introduces potential confounding effects, as has been referred to repeatedly above. Similarly, effects mediated via the peritubular myoid cells cannot be excluded in these studies. Nevertheless, these findings built on earlier studies which had indicated via other means that androgens might regulate protein synthesis and/or secretion by Sertoli cells [87–89], and the notion that androgens might have an overall effect at stages VII to VIII on Sertoli cell protein synthesis and/or secretion is still an attractive idea and worthy of further investigation [86].

C. Role of Androgens in Spermatogonial Development: Germ Cell Transplantation and Recovery of Spermatogenesis Potentially new insights into the effects of androgens on the Sertoli cell have emerged from studies in the last few years involving the recovery of spermatogenesis after induction of its regression by a variety of mechanisms, such as those used for cancer therapy (i.e., irradiation or chemotherapy). What these studies have revealed is that failure of spermatogenesis to recover in some situations is not due to the complete ablation of all germ cells but instead reflects the failure

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of early A-type spermatogonia to differentiate and survive and thus to restart the process of spermatogenesis [90–92]. Unexpectedly, recovery of spermatogenesis in these situations requires suppression of intratesticular levels of testosterone via one of several routes, such as the administration of GnRH analogues [93]. When testosterone levels are suppressed, differentiation of A-type spermatogonia occurs and gradual expansion and recovery of spermatogenesis ensues, sometimes with complete recovery to normal. These are quite remarkable effects and appear to have widespread relevance to a number of clinical situations. For example, the most recent finding has been that induction of azoospermia by administration of the nematocide dibromochloropropane (DBCP) to rats results not in complete germ cell aplasia but in regression (and arrest) of spermatogenesis back to A-type spermatogonia [94, 95]. Recovery of full spermatogenesis in DBCP-treated rats can then be triggered by suppression of intratesticular testosterone levels [95]. The experiments with DBCP in rats are of direct relevance to humans because numerous studies from the 1970s and 1980s had shown that men involved in the manufacture or application of DBCP were rendered azoospermic or oligospermic [96–98]. In a large proportion of DBCP-exposed men who became azoospermic, no recovery of spermatogenesis occurred, though in some there was recovery over many years [97, 98]. This pattern of effect and recovery is remarkably like that observed in rats treated with various gonadotoxic agents, as described earlier. This has raised the intriguing possibility of whether some men who have received gonadotoxic chemotherapy or radiotherapy, or who have been exposed to gonadotoxic chemicals such as DBCP, and are azoospermic as a consequence, are in fact suffering from arrest of spermatogenesis at an early spermatogonial stage [92]. There are so far only two published studies that have addressed this possibility directly, one supporting this possibility, the other not. In the first study, men with nephrotic syndrome were being treated with cyclophosphamide as immunosuppressive therapy. Of 15 such men, 5 were treated with exogenous low-dose testosterone to suppress intratesticular testosterone levels before and during an 8-month cycle of chemotherapy [99]. All 15 men were rendered azoospermic or severely oligozoospermic within 6 months of commencing cyclophosphamide therapy, and 9 out of 10 men who received this treatment alone remained azoospermic within 6 months after the end of chemotherapy. In contrast, in the 5 men in whom intratesticular testosterone levels had been suppressed before/during cyclophosphamide therapy, sperm counts had returned to

normal within 6 months of treatment cessation [99]; confirmation of this potentially important finding is eagerly awaited. In contrast, in a second study of seven men who had been rendered azoospermic by treatment for cancer during childhood, suppression of intratesticular testosterone levels by administration of a “male contraceptive steroid regimen” failed to initiate recovery of spermatogenesis [100]. This study also revealed the absence of any spermatogonia in testis biopsies from the seven subjects taken both before and after the steroid treatment. Although these two studies do not agree they may not be irreconcilable because the second study was undertaken 6–10 yr after cancer therapy (in childhood), whereas in the first study there was no such delay. It is quite likely that if arrest of spermatogenesis at the level of A-spermatogonia is induced, then recovery of spermatogenesis is only possible within a defined period before the remaining A-spermatogonia and stem cells degenerate in the presumed “hostile” testicular environment. The mechanisms via which suppression of intratesticular androgen levels leads to recovery of spermatogenesis in the preceding situation is still unclear although, curiously, one study has shown that irradiation of the testis results in selective inhibition of the expression of Pem [101], the androgen-dependent homeobox gene that is expressed in Sertoli cells and which was described earlier in this chapter. However, it remains unknown whether this change has any bearing on the role of androgens in regulating the recovery of spermatogenesis in these experimental situations. Indeed, it is still unknown which cell type is primarily affected by the changes in androgen levels in these situations, though effects on Sertoli cells are obviously at the top of the list. Some of the studies that have addressed this particular problem have established that conversion of testosterone to estradiol is probably not part of the mechanism involved [102] and, because germ cells do not themselves express AR, it is presumed that the effect of androgens in this situation is indirect and must be mediated via either the Sertoli cells or the peritubular cells. In this regard, the peritubular myoid cells are perhaps a valid candidate for the site of whatever androgen actions are involved in affecting spermatogonial development, because they lie in proximity to the spermatogonia. Whatever the mechanisms involved, this is clearly the very opposite situation to that in which androgens normally maintain full spermatogenesis. In the latter instance, testosterone levels within the testis have to be maintained at very high levels to ensure that complete, quantitatively normal spermatogenesis occurs, whereas in the instances of

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recovery of spermatogenesis after its experimental suppression, androgen levels have to be suppressed to low levels to enable the recovery to occur. An obvious interpretation of this difference is that the “recovery of spermatogenesis” is akin to normal puberty in which intratesticular levels of testosterone are normally low, rather than very high as in the adult testis. Therefore, when spermatogenesis is experimentally suppressed but intratesticular testosterone levels remain high, this environment is not favorable to the onward differentiation and development of spermatogonia in the absence of later germ cell types. Whether or not this particular situation has anything to tell us about the role of androgens in regulating full spermatogenesis, remains to be established.

domains on the Src protein, and antagonists of either the AR or ER can block activation of this pathway by either androgens or estrogens [107]. Whether or not the component factors of this pathway are all present in Sertoli cells and whether their expression may vary in a stage-dependent fashion remains to be shown, but because many of the components are expressed fairly ubiquitously, it is likely that this is a realistic pathway for androgen action on the Sertoli cells that does involve the AR but does not involve gene transcription as the regulatory mechanism.

D. Genomic versus Nongenomic Effects of Androgens on the Sertoli Cell

Since the last edition of The Sertoli Cell, significant progress has been made in our understanding about androgen action on the Sertoli cell, although this still remains largely descriptive with minimal information available on the underlying mechanisms via which androgen action is translated into biological support for developing germ cells. Due to numerous studies of androgen depletion and replacement and the generation of new transgenic animals in which the AR has been ablated, it is possible to at least make some summary statements:

In the numerous studies outlined earlier, the predominant presumption has been that any effects of androgens on the Sertoli cell that supports spermatogenesis are mediated via the classical AR and that this then binds to DNA and activates or represses transcription of specific genes in the standard way. However, interest is growing in the possibility that steroids such as testosterone can exert significant nongenomic effects on Sertoli cells in which a biological response is extremely rapid (a matter of seconds or minutes), effects that cannot be accounted for via the classical genomic pathways [103]. Various studies have been conducted using isolated Sertoli cells, mainly from immature rats, to study such nongenomic effects. These have shown that testosterone can cause rapid elevations in intracellular levels of calcium in Sertoli cells (e.g., [103–105]. Other studies have shown that these effects may be mediated by inhibition of K+ATP channels [103, 106]. Although it seems quite certain that the nongenomic effects occur via specific receptor or nonreceptor mechanisms in the plasma membrane of the Sertoli cells, based on the use of protein-bound ligands and other experimental approaches, it is also evident that other nongenomic pathways may exist in which the effects are mediated by intracellular mechanisms. One such pathway via which androgens may exert their effect is via activation of Src-Raf1/Shc-Erk2, in which androgens and estrogens may induce assembly of a novel ternary complex comprising the AR, ER (either ERα or ERβ) and Src [107, 108]. This complex triggers activation of the protein kinase domain of Src, and downstream effects such as cell proliferation or inhibition of apoptosis may then be induced. The androgen– AR and estrogen–ER complexes bind to separate

VI. CONCLUSIONS AND FUTURE PROSPECTS

1. Generation of the SCARKO mouse has proven definitively that androgen action on Sertoli cells is not required for normal development of the testis and reproductive system or for testicular descent into the scrotum. This understanding was more or less in place beforehand because the temporal delay in expression of the AR in Sertoli cells already clearly suggested the lack of involvement of androgen action on the Sertoli cell in these processes, as did studies in tfm animals. 2. Androgen action directly on Sertoli cells probably does not play any role in Sertoli cell proliferation, based on findings in the SCARKO mouse. Indirect actions, presumably via peritubular myoid cells, may be important in fetal life at least. 3. Androgen action on Sertoli cells probably does not play any role in their terminal differentiation/ maturation. This conclusion is tentative because it is based mainly on the species differences in timing of AR expression and its lack of coincidence with terminal maturation of the Sertoli cells [11]. Nevertheless, this conclusion is reinforced by the demonstration that spermatogenesis in SCARKO mice is initiated and progresses reasonably normally up to early stages of meiosis [33]. The available evidence suggests that if

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a Sertoli cell fails to mature then it will be incapable of supporting spermatogenesis [11]. The finding of spermatogenesis in SCARKO mice therefore implies that Sertoli cell maturation has occurred, although this has yet to be confirmed by the use of specific maturational markers [11]. 4. Androgen action on Sertoli cells is not required for the initiation of spermatogenesis or for spermatogonial replication, based on observations in SCARKO mice [33]. This represents a substantial improvement in our understanding, in that opinion was very much divided as to what the expected phenotype would be in SCARKO males; the expectation for many was that there would be inadequate or incomplete initiation of spermatogenesis and its progression. Nevertheless, there remains one caveat to this conclusion and that relates to the role and mechanism of testosterone in suppressing recovery of spermatogenesis following its regression in the adult testis [92]. It should be possible to definitively address this question by experimental studies in SCARKO males, which we hope will serve to prove whether or not it is androgen action on Sertoli cells or on peritubular myoid cells that lies at the heart of this particular regulatory puzzle. 5. Androgen action on Sertoli cells is required for the progression of germ cells through meiosis. Whether androgen action on Sertoli cells is required for spermiogenesis is more debatable. As has been indicated earlier in this chapter, several pieces of information from studies that have involved androgen depletion/repletion indicate that spermatid attachment to Sertoli cells may be dependent on androgen action on the latter. However, whether or not this is the case cannot be confirmed or denied in SCARKO mice due to the failure of virtually all germ cells to complete meiosis [33]. The same androgen depletion/ repletion studies had also indicated that spermatocytes were particularly vulnerable to loss of androgen action on Sertoli cells, and this occurred in a stage-dependent manner, so these older findings are very much in keeping with the new observations from SCARKO mice. 6. The molecular and biochemical mechanisms via which androgen action on the Sertoli cell supports spermatogenesis remain unknown. There seems little doubt that these effects occur in a stagespecific manner and that they involve quite major stage-specific changes in the overall level of protein and fluid secretion by the Sertoli cell [1], but it remains unknown how these changes are regulated. The only unequivocal androgen-regulated

gene that has been identified in Sertoli cells is the Pem homeobox gene, which has been shown to be expressed stage specifically [63]. However, the latest data on Pem suggest that it may play a role in regulating DNA repair and/or in initiating meiosis in germ cells (mechanisms unknown) rather than having a more generalized effect on Sertoli cells and thus a role in the support of germ cell development through to spermatozoa. 7. Despite the enormous technical advances in genomics and proteomics during the past 15 years, these advances have failed to identify the mechanisms via which androgen action on Sertoli cells supports spermatogenesis. From a logistical point of view, this is perhaps understandable because this task is a daunting one. Not only do normal cell–cell interactions in the seminiferous epithelium have to be maintained for such studies to be undertaken, but the numbers and types of the different germ cells have also to be maintained [1]. This is true for two reasons. First, these germ cells signal to the Sertoli cell and affect many of its functions [9, 24]. Second, these germ cells have very high and dynamic levels of gene and protein expression, and so their depletion will change the gene/protein profile [46] in a fashion that will not reflect the primary effects of androgens on the Sertoli cell. The Sertoli cells themselves occupy 7–8% of the cellular volume of the seminiferous epithelium in the normal adult testis and identification of stage-specific effects of androgens in this minor cellular component, against the high background of dynamic changes in gene and protein expression in germ cells, thus represents a Herculean task for researchers. Because the germ cell complement cannot be altered without introducing confounding effects, this problem cannot be reduced and there are no other obvious solutions. The development of laser capture microscopy promised the possibility of being able to sample individual cells in situ and then to examine their gene expression profile. Unfortunately, in practice, it is impossible to sample individual cells such as Sertoli cells (see, e.g., [109]), which lie between other potentially contaminating cells types (in this case the germ cells), which means that this technique cannot be applied to resolving the question of what genes are regulated in Sertoli cells by androgens in a stage-specific way. It is also not possible to isolate pure preparations of Sertoli cells in a stagespecific manner unless researchers are prepared for an extremely laborious task with the high

Chapter 12 Sertoli Cell Endocrinology and Signal Transduction: Androgen Regulation

chance of an inadequate return at the end. Some hope resides in further studies in SCARKO mice, though it remains to be seen whether or not the lack of particular germ cell types proves to be a confounding influence even in this situation. For the moment, it seems likely that findings from SCARKO males and ad hoc findings from other transgenic mice are the most likely sources to provide the insight that is still so urgently needed to enable researchers to trace the pathways of androgen action on Sertoli cells. In conclusion, despite the significant advances that have been made in this area, it remains clear that our lack of understanding of androgen action on Sertoli cells in the regulation of spermatogenesis remains a huge obstacle to our overall understanding of spermatogenesis and our ability to manipulate this for contraceptive purposes [2] or to try and rescue it when it does not work properly in patients. As indicated earlier, there are no straightforward paths to resolving this ignorance and no expectation that such resolution will come quickly. Indeed, insights are perhaps more likely to arise by accident. The complexity of spermatogenesis and its dynamics, in particular the dynamic interaction between germ cells and Sertoli cells, acts as a considerable deterrent to new researchers looking for an area in which to work. This is particularly true for those with molecular skills who wish to dissect signaling pathways and who recognize that this can be achieved more quickly using isolated, purified cell preparations or cell lines that have been transfected with various constructs. In many respects the challenge posed by the need to resolve the pathways of androgen action on spermatogenesis crystallizes one of the biggest dilemmas facing the biomedical world at present, namely, how to translate what we know about the structure and function of individual genes into the dynamic real world in which complex cell–cell interactions are the stuff of life itself.

Acknowledgments I thank Laura McGeer for her help in putting together this chapter. I am grateful to Karel de Gendt, Philippa Saunders, Karen Tan, Nina Atanassova, and Guido Verhoeven for collaborative studies on SCARKO mice and for allowing me to report some of the data in this chapter.

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93. Shetty, G., et al. (2000). Gonadotropin-releasing hormone analogs stimulate and testosterone inhibits the recovery of spermatogenesis in irradiated rats. Endocrinology 141, 1735–1745. 94. Meistrich, M. L., Wilson, G., Shuttlesworth, G. A., and Porter, K. L. (2003). Dibromochloropropane inhibits spermatogonial development in rats. Reprod. Toxicol. 17, 263–271. 95. Meistrich, M. L., et al. (2003). Restoration of spermatogenesis in dibromochloropropane (DBCP)-treated rats by hormone suppression. Toxicol. Sci. 76, 418–426. 96. Eaton, M., Schenker, M., Whorton, M. D., Samuels, S., Perkins, C., and Overstreet, J. (1986). Seven-year follow-up of workers exposed to 1,2-dibromo-3-chloropropane. J. Occup. Med. 28, 1145–1150. 97. Potashnik, G., and Porath, A. (1995). Dibromochloropropane (DBCP): A 17-year reassessment of testicular function and reproductive performance. J. Occup. Environ. Med. 37, 1287–1292. 98. Goldsmith, J. R. (1997). Dibromochloropropane: Epidemiological findings and current questions. Ann. NY Acad. Sci. 837, 300–306. 99. Masala, A., et al. (1997). Use of testosterone to prevent cyclophosphamide-induced azoospermia. Ann. Intern. Med. 126, 292–295. 100. Thomson, A. B., et al. (2002). Investigation of suppression of the hypothalamic–pituitary–gonadal axis to restore spermatogenesis in azoospermic men treated for childhood cancer. Hum. Reprod 17, 1715–1723. 101. Maiti, S., et al. (2001). Irradiation selectively inhibits expression from the androgen-dependent Pem homeobox gene promoter in Sertoli cells. Endocrinology 142, 1567–1577.

102. Shetty G., et al. (2002). Inhibition of recovery of spermatogenesis in irradiated rats by different androgens. Endocrinology 143, 3385–3396. 103. Silva, F. R. M., Leite, L. D., and Wasserman, G. F. (2002). Rapid signal transduction in Sertoli cells. Eur. J. Endocrinol. 147, 425–433. 104. Gorczynska E., and Handelsman, D. J. (1995). Androgens rapidly increase the cytosolic calcium concentration in Sertoli cells. Endocrinology 136, 2052–2059. 105. Lyng, F. M., Jones, G. R., and Rommerts, F. F. (2000). Rapid androgen actions on calcium signalling in rat Sertoli cells and two human prostatic cell lines: Similar biphasic responses between 1 picomolar and 100 nanomolar concentrations. Biol. Reprod. 63, 736–747. 106. Von Ledebur, E. I., Almeida, J. P., Loss, E. S., and Wassermann G. F. (2002). Rapid effect of testosterone on rat Sertoli cell membrane potential. Relationship with K+ATP channels. Horm. Metab. Res. 34, 550–555. 107. Migliaccio A., et al. (2000). Steroid-induced androgen receptoroestradiol receptor b-Src complex triggers prostate cancer cell proliferation. EMBO J. 19, 5406–5417. 108. Kousteni S., et al. (2001). Nongenotropic, sex-nonspecific signaling through the estrogen or androgen receptors: Dissociation from transcriptional activity. Cell 104, 719–730. 109. Sluka, P., O’Donnell, L., and Stanton, P. G. (2002). Stage-specific expression of genes associated with rat spermatogenesis: Characterization by laser-capture microdissection and real-time polymerase chain reaction. Biol. Reprod. 67, 820–828.

C H A P T E R

13 Thyroid Hormone Regulation of Sertoli Cell Development PAUL S. COOKE

DENISE R. HOLSBERGER

Department of Veterinary Biosciences and Division of Nutritional Sciences, University of Illinois, Urbana, Illinois

Department of Veterinary Biosciences and Division of Nutritional Sciences, University of Illinois, Urbana, Illinois

LUIZ R. FRANÇA Laboratory of Cellular Biology, Department of Morphology, Institute of Biological Sciences, Federal University of Minas Gerais, Belo Horizonte, Brazil

after birth, depending on the species. For example, in rats and mice, Sertoli cell mitogenesis is high at birth but concludes before weaning [1, 2]. Therefore, the adult Sertoli cell population is established by this time. In contrast, in larger animals such as primates, pigs, and cattle, Sertoli cell proliferation continues for several months to several years postnatally and larger animals have a biphasic pattern of Sertoli cell proliferation that differs markedly from that seen in rodents, with an initial perinatal period of proliferation and a second burst of proliferation around puberty [3–5]. Each Sertoli cell supports a relatively fixed number of germ cells, though the specific number of germ cells supported by each Sertoli cell varies with species. Therefore, factors regulating Sertoli cell proliferation and their adult numbers are critical in view of the fact that this is the major factor that establishes the magnitude of adult sperm production in that particular animal. Understanding hormonal control of Sertoli cell development is also crucial for other reasons. Maximizing sperm production is an important priority in species such as dairy cattle, where artificial insemination is used extensively, and increasing sperm production from bulls with particularly desirable genetics can have significant economic implications for semen producers. Although a number of approaches are possible to increase sperm production, the centrality of the Sertoli cell in regulating the magnitude of spermatogenesis

I. INTRODUCTION II. SERTOLI CELL DEVELOPMENT IS REGULATED BY THYROID HORMONE III. THYROID HORMONE EFFECTS ON OTHER ENDOCRINE PATHWAYS THAT AFFECT TESTICULAR DEVELOPMENT IV. THYROID HORMONE EFFECTS ON SERTOLI CELL PROLIFERATION IN SPECIES OTHER THAN RATS AND MICE V. ROLE OF THYROID HORMONE IN SERTOLI CELL DEVELOPMENT IN HUMANS VI. ARE EFFECTS OF THYROID HORMONE ON SERTOLI CELL PROLIFERATION MEDIATED BY p27Kip1 AND p21Cip1? VII. TRα1 AND TRβ1 ARE EXPRESSED IN SERTOLI CELLS VIII. CRITICAL QUESTIONS RELATED TO THYROID HORMONE EFFECTS ON SERTOLI CELLS AND TESTIS References

I. INTRODUCTION Initial Sertoli cell differentiation is induced by the Sry gene on the male Y chromosome and does not appear to involve hormones. However, hormones play critical roles in the developmental proliferation and adult function of Sertoli cells. Sertoli cell proliferation is initiated fetally and continues for variable periods SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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emphasizes that strategies to increase Sertoli cell proliferation during development, and ultimately adult Sertoli cell populations, are among the most promising potential approaches for increasing sperm production. Spermatogenesis can be quantitatively and qualitatively disrupted by a wide variety of reproductive toxicants. A number of these agents act exclusively or at least partially by affecting Sertoli cell development and/or function. For example, recent work by Sharpe and colleagues [6] showing that diethylhexaphthalate (DEHP), which has well-known adverse effects on spermatogenesis, inhibits Sertoli cell maturation and consequently their ability to support germ cell development illustrates this mode of action. Clearly, continued advances in understanding hormonal regulation of Sertoli cell development are critical for reproductive toxicology, in that this process is a frequent target of toxicants affecting testicular development and sperm production. Although toxicological exposure to various agents has well-documented deleterious effects on Sertoli cells in humans and this can be a cause of infertility, the majority of male infertility does not involve toxicological agents but instead results from a broad array of genetic, hormonal, structural, and other abnormalities. Here again, advancing our understanding of the normal regulation of Sertoli cell development will facilitate efforts to understand, diagnose, and treat a broad array of infertility cases where the etiology involves alterations in the Sertoli cell. During the past 15 years, it has become increasingly clear that thyroid hormone plays a major role in regulating Sertoli cell development, ultimate testis size, and sperm production in rodents. This chapter reviews the literature that led to the present concept that thyroid hormone is a critical regulator of the developing Sertoli cell and discusses recent developments related to the mechanism of this effect. We also present information related to thyroid hormone effects on Sertoli cells of the human and animals of economic interest, and finally discuss some of the critical gaps that remain in our understanding of the mechanism by which thyroid hormone regulates Sertoli cell development.

II. SERTOLI CELL DEVELOPMENT IS REGULATED BY THYROID HORMONE Sertoli cell proliferation begins fetally in rodents and is rapid at birth. This proliferation declines rapidly during the neonatal period, and Sertoli cell proliferation has essentially ceased by approximately day 16 postnatal

in the rat [2] and mouse [1]. Follicle-stimulating hormone (FSH) from the pituitary is a major regulator of neonatal Sertoli cell proliferation, as described elsewhere in this volume (Chapters 10 and 11). However, neonatal Sertoli cells slow and finally stop their proliferation despite continual FSH exposure neonatally [7]. Similarly, cultured neonatal Sertoli cells respond mitogenically to FSH, but juvenile or adult Sertoli cells do not [8, 9]. These findings indicate that some factor other than FSH is responsible for developmental changes that result in the transition of the neonatal Sertoli cell from a proliferative to a nonproliferative state. Thyroid hormone appears to play a key role in Sertoli cell maturation, and in this capacity promotes both cessation of Sertoli cell proliferation and the key developmental changes that allow Sertoli cells to support germ cell development. Older work had indicated that thyroid hormone was not a major factor in regulation of the adult testis [10, 11]. However, pioneering work by Palmero et al. [12] and Jannini et al. [13] indicated that thyroid hormone receptors (TRs) were present in high quantities in neonatal Sertoli cells, then the TR expression declined to low levels by adulthood. Thyroid hormone treatment was shown to induce changes in various secretory proteins [14, 15], indicating that thyroid hormone could have significant functional effects on young Sertoli cells. Early hypothyroidism inhibits testicular growth, germ cell maturation, seminiferous tubule lumen formation, and other developmental events [16–18]. However, neonatal hypothyroidism in rats, followed by a subsequent recovery to euthyroidism, produced unexpected and unprecedented increases in testis size and daily sperm production (DSP) during adulthood [19, 20]. Testis weight and DSP were 80 and 140% greater than normal, respectively, in adult rats that had been transiently hypothyroid neonatally. This raised important questions related to the role of thyroid hormone in the neonatal testis and to how a short period of neonatal hypothyroidism could produce the pronounced stimulatory effects on adult testis size and DSP despite the known inhibitory effects on the testis during the period of hypothyroidism. A critical finding that helped define the mechanism of this effect was that hypothyroidism lengthened and hyperthyroidism shortened the period of Sertoli cell proliferation in rats and mice ([1, 21, 22]; Fig. 13.1). These results indicated that neonatal hypothyroidism allowed extended Sertoli cell proliferation, leading to an increased adult population of Sertoli cells [23] that was responsible for the observed increases in adult testis size and sperm production in the adult following neonatal hypothyroidism. Subsequent work

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III. THYROID HORMONE EFFECTS ON OTHER ENDOCRINE PATHWAYS THAT AFFECT TESTICULAR DEVELOPMENT

FIGURE 13.1 Effect of hypothyroidism and hyperthyroidism on neonatal Sertoli cell proliferation in the rat. (Data adapted from van Haaster et al. [21, 22].)

indicated that the effects of the biologically active thyroid hormone, 3,5,3’-triiodo-L-thyronine (T3), on Sertoli cell proliferation and a variety of other developmental markers was predominately direct [24–28], though the ability of T3 to alter levels of other hormones and receptors in the Sertoli cell indicated that T3 could also have indirect effects through its modulation of other endocrine signaling systems. The picture that has emerged from extensive in vivo and in vitro studies from laboratories all across the world is that T3 normally stimulates maturation of the neonatal Sertoli cell, which entails a cessation of proliferation and a concomitant functional maturation to support germ cells. This normal developmental sequence can be experimentally perturbed by the induction of hypothyroidism, which allows extended Sertoli cell proliferation and produces an increased pool of Sertoli cells despite its immediate inhibitory effects on germ cell development and testis growth [16, 19]. When euthyroidism is restored, Sertoli cell differentiation occurs and these cells become capable of supporting full spermatogenesis. The increased Sertoli cell population [23] then results in increased numbers of germ cells, probably due to the increased capacity of the larger Sertoli cell population to support germ cell development [29]. These processes ultimately result in large increases in testis size and sperm production [19, 20].

T3 can directly decrease proliferation and increase expression of protein markers characteristic of the mature Sertoli cell. However, T3 has effects on the production of other hormones and levels of expression for hormone receptors that could be involved in Sertoli cell mitogenesis, so it is presently unclear what role secondary changes in expression of other receptors and/ or hormones could play in the observed inhibitory T3 effect on Sertoli cell proliferation. Androgen receptor (AR) is expressed at minimal levels in the rodent Sertoli cell at birth, but AR expression increases markedly during the postnatal period, and is present at high levels in adult Sertoli cells (reviewed in [5]). The period of increasing Sertoli cell AR expression corresponds with declining Sertoli cell proliferation, leading some authors to suggest that androgens may play an inhibitory role in Sertoli cell proliferation [5, 30] and be involved in the decreasing Sertoli cell proliferation seen in neonatal rodents. T3 has strong stimulatory effects on AR expression [26], probably as a result of stimulatory effects on Sertoli cell maturation. This increase in Sertoli cell AR could result in increased androgen/AR signaling. Thus, the decreased Sertoli cell proliferation induced by T3 may partially involve secondary effects on the androgen/AR signaling pathway. During early development, Sertoli cells express aromatase, and thus can produce estrogens with potential autocrine/paracrine effects on the Sertoli cell. T3 decreases aromatase mRNA, protein, and activity and consequently estrogen production by cultured Sertoli cells [31, 32]. This potentially may have effects on Sertoli cell proliferation and/or differentiation, though the role of endogenous estrogen in normal Sertoli cell development remains to be established. Müllerian inhibiting substance (MIS) is produced by Sertoli cells at high levels during fetal life, and is essential for regression of Müllerian ducts in developing males. During postnatal life in rodents, MIS production by the Sertoli cell declines, again a process that correlates temporally with declining Sertoli cell proliferation. T3 at physiological concentrations (1 nM) decreases MIS mRNA by more than 80% in cultured neonatal Sertoli cells [25]. Sertoli cells express the type II MIS receptor [33], indicating that effects on MIS production could have autocrine effects on Sertoli cell development. In addition, MIS regulates Leydig cell

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maturation/proliferation [34], so changes in Sertoli cell MIS production could have effects on Leydig cells. Because Leydig and Sertoli cells communicate extensively, effects on Leydig cell proliferation/differentiation could have secondary effects on Sertoli cell development. T3 increases expression of mRNA for the α subunit of inhibin (inhibin-α) in cultured neonatal Sertoli cells [25]. Transgenic male mice lacking inhibin-α are unable to make functional inhibins and develop Sertoli cell tumors, suggesting that inhibin negatively regulates Sertoli cell proliferation [35]. Inhibitory effects of T3 on inhibin-α mRNA expression by Sertoli cells suggest that T3 effects on Sertoli cell proliferation in vivo could also involve secondary effects resulting from T3 effects on inhibins. In summary, T3 affects the production of various hormones and receptors potentially involved in Sertoli cell proliferation. Elucidating the overall effects of T3 on Sertoli cell proliferation must take into account effects on these other endocrine pathways as a factor in overall T3 effects on Sertoli cells to inhibit their proliferation.

IV. THYROID HORMONE EFFECTS ON SERTOLI CELL PROLIFERATION IN SPECIES OTHER THAN RATS AND MICE Similar to rats and mice, neonatal treatment with the goitrogen propylthiouracil (PTU) significantly increased adult testis weight and sperm production in hamsters, though it reduced serum concentrations of FSH and LH ([36]; Table 13.1). However, the effective dose of PTU in hamsters was much higher than that required to produce testis increases in the rat, suggesting that each species might differ in its response to PTU. In the chicken (Table 13.1), transient hypothyroidism in roosters induced by PTU treatment from 6 to 12 weeks after hatching doubled adult testis weight and sperm production [37]. Treatment with PTU during later time periods (either 8–14 or 10–16 weeks of age) resulted in smaller (35%) increases in testis mass at adulthood relative to controls. Thus, PTU treatment during critical developmental windows can cause permanent increases in testis size and sperm production in the rooster. However, Knowlton et al. [38] showed that early hypothyroidism during a variety of postnatal windows in turkeys caused precocial testis development and a transient increases in testis size, but this was not maintained into adulthood, emphasizing that results obtained in chickens might not be generally applicable to other avian species. Hypothyroidism induced in young Nile tilapia (Oreochromis niloticus; body weight of ~1 g) by addition

of either of two different concentrations (100 or 150 ppm) of PTU to their water for 40 days resulted in increases of approximately 100% in gonadosomatic index (testis mass/body weight) and testis weight after the end of treatment (Table 13.1; [39]). Similar to rodents, during PTU treatment, the fish showed delayed spermatogenesis and decreased body weight, but both of these parameters recovered to normal in adult fish after PTU treatment. Morphometric analysis showed that seminiferous tubule area, number of Sertoli and germ cells per spermatogenic cyst, and total Leydig cell number per testis significantly increased in treated fish. Taken together, these data indicate that increased Sertoli cell proliferation was probably the primary factor that caused increased testis weight in this fish following PTU treatment. Remarkably, results seen in rats treated with PTU were reproduced in fish, suggesting strongly that the mechanisms by which thyroid hormone regulates Sertoli cell proliferation are conserved during evolution. Although more studies are necessary, these results show that early hypothyroidism may potentially provide a new method for improving sperm production and reproductive efficiency in fish. The ability of early hypothyroidism to increase adult testis size and sperm production in species as disparate as rodents, birds, and fish suggests that this methodology could potentially be used to increase sperm production in all species. The possibility of applying this methodology to mammalian species of economic interest has attracted significant interest during the past decade, especially in those species in which artificial insemination is used extensively and there would be an obvious economic benefit from increasing sperm production by the best males. Results to date have been disappointing, although it remains unclear whether it is possible to increase sperm production by neonatal hypothyroidism in certain species or if the correct conditions to do this successfully have just not yet been identified. Studies with pigs have indicated that PTU treatment at different time periods after birth, and before puberty, was not able to increase testis size ([40, 41]; Table 13.1) or sperm production [41]. Silva Jr. [42] showed that PTU treatment for about 3 months after birth actually decreased adult testis size, seminiferous tubule volume density and length, total number of Sertoli cells per testis, and sperm production, although the spermatogenic process and testis structure were not altered. The absence of a stimulatory effect of early hypothyroidism in this species does not appear to reflect a lack of responsiveness to thyroid hormone during this period. Palmero et al. [43] reported

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Chapter 13 Thyroid Hormone Regulation of Sertoli Cell Development TABLE 13.1 Effects of Transient Postnatal Treatment with 6-n-Propyl-2-thiouracil (PTU) on the Number of Sertoli Cells per Testis, Testis Size, and Sperm Production in Different Species

Species

Length of treatment (days)a

PTU concentration

Testis size

Number of Sertoli cells per testis

Sperm production

Reference

Rat Mouse

0–25 0–25

0.1% 0.1%

↑ 82% ↑ 29%

↑ 157% —

↑ 136% ↑ 52%

[19, 20, 23] [1]

Hamster

0–25

0.1% + 0.4%

↑ 30%

—

↑ 73%

[36]

Pig: Varied

0.01–0.03%

↓ 7–12% (n.s.)

—

—

[40]

7–102

8 mg kg/BW

↓ 30%

↓ 36%

↓ 50%

[42]

Varied

0.1%

n.s.

—

—

[41]

Chicken

Varied

0.1%

↑ 35–96%

—

↑ 15–115%

[37]

Turkey

Varied

0.1–0.5%

n.s.

—

n.s.

[38]

Fish

40b

100–150 ppm

↑ 95–117%

↑ 73–104%c

↑ 29–63%c

[39]

Abbreviation: n.s. = Not significant statistically. Duration of treatment after birth. b After fish reached 1 g of body weight. c Related to the number of Sertoli and germ cells per spermatocyte and spermatid cysts. a

TR expression in neonatal boar testes, and T3 had effects on protein synthesis in these cells [44]. In addition, treatment of boars with T3 for several weeks after birth [42] accelerated the onset of spermatogenesis, although it decreased Sertoli cell number. Thus, young boar testes express TR and respond to thyroid hormone, although transient neonatal hypothyroidism does not increase adult testis weight and sperm production as reported for other species; the reason for this is unclear. Majdic et al. [45] reported an inverse relationship between neonatal T4 levels and postpubertal testis size in bulls. Though this suggests thyroid hormone might have the same effects on testicular development in cattle as in rodents, there have been no reports of successfully increasing adult testis size and sperm production using neonatal hypothyroidism in bulls. We have observed that a marked hypothyroidism can be induced in neonatal rams or bulls by feeding PTU. This treatment causes body and testis weight decreases that parallel those seen in rats made neonatally hypothyroid. Following cessation of PTU treatment, compensatory body weight growth is observed in both species, and testis weight increases rapidly, similar to the rat. However, testis weight and sperm production in treated rams and bulls plateau at approximately normal levels, and adult testis weight and sperm production were never increased by various regimens of neonatal PTU treatment (Cooke, unpublished data). Although successful use of the neonatal hypothyroidism technique to increase adult Sertoli cell number

and sperm production in large mammalian species cannot be ruled out, these species have a longer and biphasic period of Sertoli cell proliferation [3, 5, 46]. Differences in Sertoli cell development and/or hormonal responsiveness as well that present challenges in adapting this technique to these animals will also need to be considered.

V. ROLE OF THYROID HORMONE IN SERTOLI CELL DEVELOPMENT IN HUMANS A role for T3 in human Sertoli cell development has not been definitively established, but available data are consistent with the concept that developing human Sertoli cells may be regulated by T3. An extensive clinical literature stretching back over a half century indicates that hypothyroidism in prepubertal boys is associated with testicular enlargement, but without virilization (reviewed in [47]). The etiology of the testicular enlargement is not definitively established, but could be due to increased Sertoli cell proliferation resulting from decreased circulating thyroid hormone. In support of this possibility, expression of TRα1 mRNA in the fetal, neonatal, and adult human testis was recently reported [48]. Expression of TRα1 mRNA was relatively high in young testes then decreased throughout development, becoming minimal in the adult. As in rodents, this TRα1 mRNA expression

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appeared to be predominately or exclusively in the Sertoli cells. Thus, the basic pattern of TRα1 expression in humans corresponds with that originally reported for the rat [49] and suggests that in humans, as in other animals, TRα1 mRNA is expressed early in Sertoli cell development and may play a role in proliferation and maturation of these cells.

VI. ARE EFFECTS OF THYROID HORMONE ON SERTOLI CELL PROLIFERATION MEDIATED BY p27Kip1 AND p21Cip1? Since the demonstration a decade ago that hypo- or hyperthyroidism altered Sertoli cell proliferation, one of the most critical questions in this field has been “What is the molecular mechanism by which T3 inhibits Sertoli cell proliferation?” Recent data showing that the cell cycle regulators p27Kip1 and p21Cip1 are involved in Sertoli cell proliferation and are regulated by T3 have provided new mechanistic insights into this long-standing question. During recent years, remarkable progress has been made in our understanding of the cell cycle and factors that control progression of cells through, or their exit from, the cell cycle into a differentiation pathway. A family of proteins termed cyclins regulates progression of mammalian cells through the cell cycle. These cyclins work by associating with and activating cyclin-dependent kinases (Cdks). Activation of the cyclin-dependent kinase complexes (cyclin + Cdk) results in progression of cells through the various phases of the cell cycle. The cyclin-dependent kinase complexes themselves in turn are regulated by cyclin-dependent kinase inhibitors (CDKIs), which bind cyclin-dependent kinases and exert negative control over progression through the critical G1 checkpoint of the cell cycle. CDKIs consist of two families of at least seven total individual proteins. The Cip/Kip family of CDKIs consists of three members (p21Cip1, p27Kip1, and p57Kip2), whereas the Ink4 family consists of four members. Testis expresses high levels of p27Kip1, which is predominately, though not entirely, in Sertoli cells [50]. Expression of p27Kip1 is inversely correlated with Sertoli cell proliferation. Levels of p27Kip1 are minimal in rapidly proliferating neonatal Sertoli cells [50]. Conversely, p27Kip1 expression is maximal in adulthood when Sertoli cell proliferation has ceased [50]. The importance of p27Kip1 in Sertoli cell proliferation is further illustrated by the report that p27Kip1 expression is sharply decreased in Sertoli cell tumors [51], which are proliferating rapidly. These findings, along with the extensive literature indicating that p27Kip1 is a major regulator of the cell’s decision to remain in or exit the cell cycle, suggest that the low neonatal p27Kip1 levels allow rapid Sertoli

cell proliferation, whereas high p27Kip1 levels seen later result in a cessation of Sertoli cell proliferation. Mice lacking either p27Kip1 [52, 53] or p21Cip1 (Holsberger and Cooke, unpublished) have increases in testis size and weight (Fig. 13.2). In addition, these effects are additive, and adult p21Cip1/p27Kip1 double knockouts have increased testicular organomegaly that is greater than with either single knockout alone (Fig. 13.2). These animals had increases in daily sperm production compared to wild-type controls that paralleled the increases in testis weight (Holsberger and Cooke, unpublished), and these increases were accompanied by similar increases in Sertoli cell numbers compared to wild-type controls (Holsberger and Cooke, unpublished). These results, indicating that both p27Kip1 and p21Cip1 regulate Sertoli cell proliferation and the establishment of adult Sertoli cell numbers, are of added significance in light of recent work from two laboratories showing that T3 regulates p27Kip1 and p21Cip1. Expression of p27Kip1 was higher in Sertoli cells from hyperthyroid mice when compared to euthyroid controls, whereas decreases in p27Kip1 expression were observed in Sertoli cells from hypothyroid mice ([54]; Fig. 13.3). Furthermore, T3, as well as retinoic acid and testosterone, was capable of inducing p27Kip1 and p21Cip1 protein expression in cultured neonatal rat Sertoli cells [30]. These results suggest that the ability of T3 to terminate postnatal Sertoli cell mitogenesis may involve stimulation of both of these cell cycle inhibitors. Neonatal hypothyroidism would therefore inhibit levels of both p27Kip1 and p21Cip1, and this may allow extended Sertoli cell proliferation and result in increased Sertoli cell populations and increased adult testis size and sperm production. Although these findings provide a mechanistic framework for further study of molecular changes in Sertoli cells induced by T3, it should be emphasized that the demonstrated link between T3 and p27Kip1 and p21Cip1 does not yet establish causality.

FIGURE 13.2 Loss of p21Cip1 and/or p27Kip1 results in testicular organomegaly. Testes taken from 4-month-old (A) wild-type, (B) p21Cip1 knockout, (C) p27Kip1 knockout, and (D) p21Cip1/p27Kip1 double-knockout mice after vascular perfusion. Scale bar = 5 mm.

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FIGURE 13.3 Immunohistochemical detection of p27Kip1 protein in 10-day-old testes taken from (A) euthyroid, (B) hypothyroid, and (C) hyperthyroid mice (see color plate). Positive staining for p27Kip1 is denoted by brown coloration. Scale bar = 25 μm.

Additional experiments are necessary to demonstrate whether changes in p27Kip1 and/or p21Cip1 are necessary and/or sufficient for the T3-induced decrease in Sertoli cell proliferation.

VII. TRα1 AND TRβ1 ARE EXPRESSED IN SERTOLI CELLS Nuclear TR, which are ligand-modulated transcription factors, regulate the effects of T3 on target organs. There are three functional TRs: TRα1, TRβ1, and TRβ2. TRβ2 has a restricted distribution in the nervous system and is unlikely to be involved in testicular development. A number of laboratories have shown that

TRα1 mRNA and protein are expressed at high levels in the developing testis, predominately in Sertoli cells ([12, 13, 49, 55, 56]; Fig. 13.4). Sertoli cells in culture express high levels of TRα1 mRNA, and these cells respond to T3 in vitro [24, 26]. Other groups [56, 57] have reported that TRβ1 mRNA could be detected by PCR in Sertoli cells from both immature piglets and rats. However, TRβ1 mRNA was not expressed in developing human testis [48]. The preponderance of TRα1 in the developing Sertoli cell and the lack of TRβ1 in the human Sertoli cell suggests that TRα1 may be the normal mediator of T3 effects on Sertoli cell development, although this has not been directly established.

VIII. CRITICAL QUESTIONS RELATED TO THYROID HORMONE EFFECTS ON SERTOLI CELLS AND TESTIS

FIGURE 13.4 Expression of TR in 5-day-old rat testes (see color plate). Immunohistochemical detection of TR was performed using an anti-pan TR primary antibody. Brown color indicates positive staining for TR. A negative control section is shown as an inset. Arrows: Sertoli cell nuclei, G: gonocyte. (From Buzzard et al. [56]. Copyright © 2000, The Society for the Study of Reproduction, Inc.)

The past 15 years have witnessed a dramatic paradigm shift in terms of our view of thyroid hormone’s role in the testis. The original concept derived from adult studies that thyroid hormone was not a major factor in the testis has been supplanted by the concept that thyroid hormone is a critical developmental regulator of the testis due in large part to its effects on Sertoli cell development. A number of critical questions still must be answered to fully elucidate how T3 regulates Sertoli and testicular development. The reason for the inconsistency of the effects of neonatal hypothyroidism on Sertoli cell proliferation between species is unclear. The explanation may involve differences in patterns of Sertoli cell proliferation (reviewed in [5]). Although it appears that in most large mammalian species the neonatal proliferative

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period is the more critical determinant of ultimate Sertoli cell number, it is unknown whether this period of Sertoli cell proliferation, or the peripubertal period of proliferation, would be the most amenable to modulation by hypothyroidism or even whether it is possible to use hypothyroidism to increase Sertoli cell numbers in some species. These questions need to be answered if this technique is ever to be used to increase sperm production in animals of economic interest. TRα1 is expressed at high levels in human Sertoli cells during the fetal/neonatal period, suggesting that T3 could be involved in Sertoli cell development. However, the expression of TR has not been examined during the second burst of mitogenesis that occurs in peripubertal testis, so it is unclear whether TR expression is also present during this time. A definitive determination of whether T3 regulates Sertoli cell development in humans is needed. If there are T3 effects, it is also critical to establish whether T3 has effects on Sertoli cell proliferation during the neonatal and/or peripubertal periods of Sertoli cell mitogenesis. Present literature suggests that TRα1 is more highly expressed than TRβ1 in developing Sertoli cells, but it is important to directly determine if T3 acts through TRα1 and/or TRβ1 to inhibit Sertoli cell proliferation. Steroid hormones such as estrogen act by binding to their receptor and turning on target genes. T3 also acts on TRs to activate gene transcription, but in contrast to estrogen, unliganded TRα1 and TRβ1 also have repressive effects on expression of certain genes [58]. This accounts for the initially paradoxical observation that the phenotype of even the TRα/TRβ double knockout is less severe than in hypothyroid mice [58–60]; genes normally repressed by unliganded TRs are transcribed in knockouts (just as they would be if both T3 and TR were present), leading to a less severe phenotype in TR knockout mice than hypothyroid mice. It is also critical to establish whether T3 effects through TR in Sertoli cells are due to T3 acting on these receptors to stimulate transcription of target genes or to release inhibition of genes normally repressed by unliganded TR. Mice lacking either p21Cip1 and/or p27Kip1 have increases in testis size, Sertoli cell numbers and DSP, and both p21Cip1 and p27Kip1 appear to be regulated by T3 in Sertoli cells [30, 54]. A clear priority for completely elucidating the mechanism of the T3 effect on Sertoli cells is to establish how T3 regulates these CDKIs, and the precise role that p21Cip1 and p27Kip1, alone and in combination, play in the T3 response in the developing Sertoli cell. Another critical question to answer is whether T3 effects on p21Cip1 and p27Kip1 can totally account for

the T3 effects on Sertoli cell proliferation/differentiation, or whether T3 causes additional effects not involving p21Cip1 and p27Kip1 but critical for the T3 effect on Sertoli cell proliferation. For example, in oligodendrocyte precursor cells, T3 directly inhibits the E2F transcription factor that is critical for progression of cells through the G1 restriction point [61]. It is unknown whether T3 has effects on E2F in Sertoli cells and, if so, whether this could play a role in the effects of T3 on Sertoli cell proliferation. T3 could also have effects on other targets that could directly or indirectly affect p27Kip1 expression and Sertoli cell proliferation. For example, connexins are a family of approximately 20 gap junction proteins, and connexin 43 (Cx43) has been implicated in regulation of cell growth and differentiation, both in normal and tumor cells [62]. Cx43 in seminiferous epithelium is confined to Sertoli cells [63], where it appears to have a critical function because Cx43 knockout mice lack germ cells [64, 65]. In rats, Cx43 was localized predominately in the cytoplasm of Sertoli cells during early postnatal life [63], but by day 30, Cx43 was entirely along the Sertoli cell plasma membrane in control rats. Neonatal hypothyroidism inhibited the relocalization of Cx43 from the cytoplasm to the lateral plasma membrane that normally occurred during postnatal Sertoli cell development. The cytoplasmic form of Cx43 is nonfunctional, whereas Cx43 in the plasma membrane is believed to be biologically active. Could the delayed Cx43 localization into the plasma membrane of hypothyroid mice be involved in the prolonged Sertoli cell mitogenesis in PTU-treated mice? Is this potentially a separate pathway through which T3 could modulate growth, or is this somehow involved with the T3 effects on p27Kip1 expression, as suggested by the report that forced Cx43 expression in lung and liver carcinoma cells in vitro [66] results in increased p27Kip1 expression and a concomitant decrease in proliferation? In conclusion, T3 is now well established as a critical regulator of Sertoli cell proliferation and maturation. Although significant progress has been made in this area, critical questions must still be addressed to establish the mechanism of this effect and understand how T3 effects on Sertoli cell development integrate into the overall developmental sequence of this cell in a variety of different species.

Acknowledgments The authors gratefully acknowledge the support from the National Institutes of Health, USDA, and the University of Illinois that has made our work possible.

Chapter 13 Thyroid Hormone Regulation of Sertoli Cell Development

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C H A P T E R

14 The Transforming Growth Factor β Superfamily in Sertoli Cell Biology KATE L. LOVELAND

DAVID M. ROBERTSON

Monash Institute of Reproduction and Development & The Australian Research Council Centre for Excellence in Biotechnology and Development, Monash University, Clayton, Victoria, Australia

Prince Henry’s Institute for Medical Research, Clayton, Victoria, Australia

trate what is known about TGFβ superfamily action at each stage of Sertoli cell development. Potential areas of functional overlap between family members and areas requiring further study will be highlighted. The current clinical applications of inhibin and activin assays relative to testicular function are addressed.

I. INTRODUCTION II. GENERAL STRUCTURE AND SIGNALING PATHWAYS III. INHIBINS AND ACTIVINS IV. MÜLLERIAN INHIBITORY SUBSTANCE V. BONE MORPHOGENETIC PROTEINS VI. TRANSFORMING GROWTH FACTOR β PROTEINS VII. GLIAL-DERIVED NEUROTROPIC FACTOR VIII. ANTAGONISTS OF TGFβ SUPERFAMILY SIGNALING IN THE TESTIS IX. CLINICAL RELEVANCE OF INHIBIN AND ACTIVIN X. IMPLICATIONS OF INTEGRATED TGFβ SUPERFAMILY FUNCTION IN THE TESTIS: WHAT WE DO AND DON’T KNOW References

II. GENERAL STRUCTURE AND SIGNALING PATHWAYS The TGFβ superfamily of cytokines/growth factors consists of more than 40 members, many of which are important in gonadal development and function [1, 2]. The members of this superfamily include: TGFβ, inhibins/activins, bone morphogenetic proteins (BMPs), Müllerian inhibitory substance (MIS), growth and differentiation factor 9 (GDF9), and glial-derived neurotropic factor (GDNF). Proteins in this family are characterized by amino acid homology centered on the location of conserved disulfide bonds that form a double cysteine knot structure. They exist extracellularly as dimeric forms, many of which are disulfide linked and are found as both latent precursor and biologically active carboxy-terminus mature forms. Binding of these ligands as homodimers or heterodimers causes receptor dimerization to form heterotetrameric receptor complexes that activate intracellular signaling via serine/threonine phosphorylation. For the general structure of these proteins, the reader is

I. INTRODUCTION The pleiotropic biological effects of the transforming growth factor β (TGFβ) superfamily in mammals are mediated by a set of functionally overlapping ligands, membrane receptor subunits, intracellular signaling molecules, and soluble and membrane bound signaling antagonists. In this chapter, the diverse components involved in TGFβ superfamily signaling in the testis will first be reviewed independently. Their roles in testis development and function will be presented to illusSERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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Copyright 2005, Elsevier Science (USA). All rights reserved.

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Kate L. Loveland and David M. Robertson TABLE 14.1 TGFβ Superfamily Ligands and Antagonists Identified in the Testis/Sertoli Cells

Ligands

Sites of detection

Activin A

References

Sertoli cells, Leydig cells, gonocytes

[19, 24, 30, 32]

Activin B

Sertoli cells, spermatocytes, round spermatids

[17, 27, 192]

MIS

Sertoli cells

[73]

BMP4

Sertoli cells

[92]

BMP7

Spermatogonia, early pachytene spermatocytes, spermatids

[96]

BMPs 8A and 8B

Pachytene spermatocytes, round spermatids

[94, 95]

TGFβ1

Sertoli cells

[101, 103, 105, 106]

TGFβ2

Sertoli cells

[101, 106]

TGFβ3

Sertoli cells, fetal Leydig cells, gonocytes, spermatogonia

[101, 107]

GDNF

Sertoli cells, peritubular cells, endothelial cells

[127, 127, 135]

NRTN

Sertoli cells

[126, 131]

Antagonists

Ligands

Betaglycana

Activin, BMP, TGFβ

Leydig cells, germ cells

[30, 102, 143, 144]

Inhibin

Activin, TGFβ

Sertoli cells, Leydig cells

[24, 30]

Follistatin

Activin, BMP4

Sertoli cells, spermatogonia, spermatocytes

[24, 156]

Bambi

Activin, BMP, TGFβs

Sertoli cells, spermatogonia, spermatocytes, round spermatids

[43]

CRIM1

BMP

Fetal Sertoli cells

[166]

α2-Macroglobulin

Activin

Sertoli cells

[167]

a Betaglycan

Sites of detection

References

(TGFRIII) is an agonist of TGFβ binding but an antagonist of activin and BMP signaling.

referred to other references [1, 3]. A list of proteins important in Sertoli cell function is presented in Table 14.1. The current model for TGFβ and activin signaling is based on the initial high-affinity binding of dimeric ligand to a constitutively active type II receptor, which then forms a heterodimer with a second type I receptor (Fig. 14.1) (for recent reviews, see [4, 5]). In contrast, the BMPs bind to both type I and II receptor subunits simultaneously before receptor dimerization. The different type I and II receptors and their ligands are presented in Table 14.2. It is important to note that a ligand can interact through several combinations of type I and II receptors to generate an intracellular signal. For example, BMPs can signal through either BMP receptor type II (BMPRII) or activin receptor type II (ActRII) to interact with either BMP or activin type I receptors (BMPRI or ActRI). A host of signaling antagonists, both soluble and membrane bound, have been identified (reviewed in [6]). Those shown to be relevant to

Sertoli cell biology are presented in Table 14.1 and will also be discussed further in Section VIII. Following receptor dimerization, phosphorylation of the type I receptor by the type II receptor subunit activates its kinase domain. This active serine/threonine kinase then phosphorylates and mediates release from the plasma membrane region of intracellular signaling factors, called receptor Smads (R-Smads). R-Smads are specific for each activated type I receptor, with TGFβ and activin activating Smad 2 and 3, and BMP and MIS activating Smads 1, 5, or 8 (Fig. 14.1). Activated R-Smads interact with a coreceptor Smad, Smad 4, which enables the R-Smad to enter the nucleus. Inhibitory Smads, Smads 6 and 7, have also been identified, and these compete for Smad 4 binding in the cytoplasm. In the nucleus, the translocated Smads interact with DNA in conjunction with a range of transcription factors (e.g., FAST-1, c-Jun, c-Fos). Smad 4, for example, stabilizes the interaction between Smad 2 and FAST-1, a transcription factor that mediates activin-induced regulation.

Chapter 14 The Transforming Growth Factor β Superfamily in Sertoli Cell Biology

A. BMPs ACTIVINs TGF β s

C Y T O P L A S M

B.

C.

Smad 4

Smad 1, 5, 8

TYPE 1 RECEPTOR

LIGAND

TYPE 2 RECEPTOR

229

Smad 2,3

N U C L E U S

Smad 6, 7 FIGURE 14.1 TGFβ superfamily members signal as dimers (stippled bars) using distinct and overlapping signaling components (filled and striped bars). (A) BMPs and activins use common Type 1 and Type 2 receptor molecules, while TGFβs use a distinct receptor set. (B) Inside the cell, signaling by activins and TGFβs activates the common R-Smads, Smad 2 and 3, while BMPs can activate R-Smads 1, 5, and 8. (C) All activated R-Smads bind to a common Smad 4 in the cytoplasm and this complex translocates into the nucleus to effect transcription. Inhibitory Smads 6 and 7 can prevent binding of the Smad 4 to any R-Smad, thereby preventing nuclear translocation and ligand-activated signaling.

III. INHIBINS AND ACTIVINS The inhibin proteins exist as a dimer of two subunits: one α inhibin subunit and one of either activin βA or βB subunits; these are termed inhibin A and inhibin B, respectively. Activin isoforms are homodimers and heterodimers of βA and βB subunits, termed activin A (βAβA), activin AB (βAβB), and activin B (βBβB). Genes encoding additional activin β subunits (C, D, and E) have been identified but have not been detected in the Sertoli cell. Because activin and inhibin share a common subunit, aspects of their structure and biological functions are intimately linked and are discussed as such within this section. Distinct roles for inhibin are addressed separately.

A. Inhibin Structure and Synthesis Inhibin, produced both by the Sertoli and Leydig cells in the testis [7, 8], was identified by its in vivo and in vitro follicle-stimulating hormone (FSH) suppressing activity [9, 10]. Inhibin, in conjunction with testosterone (their respective roles differ between species)

regulates FSH secretion by a feedback mechanism (reviewed in [11–13]). Inhibin is produced as precursor α and β subunit forms by the Sertoli cell, which are believed to dimerize intracellularly (120 kDa) and are then processed either intracellularly or extracellularly to form partially processed (~60 kDa) and mature forms (30 kDa). Inhibin B is the primary inhibin form in the adult male human and rat [14–18], whereas inhibin A has been identified in ram serum [19] and early in postnatal development in the rat [18]. It is unclear whether inhibin A and B exhibit qualitative or quantitative differences in biological activities. Up to 50% of inhibin B in male human serum is partially processed, while the remainder is fully processed. The free α subunit is also produced by Sertoli and Leydig cells; however, no biological activity has been ascribed to it in the male.

B. Activin Structure and Sites of Synthesis Activin was originally identified and purified based on its ability to stimulate FSH secretion by rat

TABLE 14.2 TGFb Superfamily Ligands, Receptors, and Smad Intracellular Effectors Implicated in Testis Functiona Ligand

Type II receptor

Type 1 receptor

R-Smads

Co-Smads

I-Smads

Activin A, AB, B

ActRIIA (ACVR2); ActRIIB (ACVR2B)

ActRIA (ALK2); ActR1B (ALK4)

2 or 3

4

7

MIS

ActRIIB; (MISRII)

ActRIA BMPRIB

1, 5 or 8

4

6

BMP 4, 7, 8A, 8B

BMPRII (BMPR2); ActRIIA ActRIIB

ActRIA BMPRIA (ALK3); BMPRIB (ALK6)

1, 5 or 8

4

6

TGFβ 1, 2, 3

TβRII (TGFBR2)

TβRI (ALK5)

2 or 3

4

7

aFrom

[2, 74, 99].

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Kate L. Loveland and David M. Robertson

pituitary cells [20, 21]. Since then it has been demonstrated to play major roles in tissue differentiation, including erythroid differentiation, and in reproductive and immune system function (reviewed in [22, 23]). Similar to inhibin, activin is produced from precursor β subunit forms by many testicular cell types, including the Sertoli cell. These dimerize intracellularly and are processed from mature forms of about 25 kDa. The α and βB subunits have been identified by immunocytochemistry and in situ hybridization techniques in the rat [24, 25, 192], human [17, 26–28], and ram [19] Sertoli cells. The localization of the βA subunit is less certain with immunocytochemical activity absent [25] or present although at low levels in the rat at most ages [24, 26, 28]. In contrast, βA subunit is readily detected by these methods in the ovine Sertoli cell and within gonocytes in both ovine and murine testes [19, 24]. Activin receptors (ActRII) have been located by in situ hybridization in rat [29] and human (ActRIIA and B [30]) Sertoli cells, whereas Smad2 and Smad3 mRNAs have been detected in rat Sertoli cells [31]. The mature 25-kDa dimer of activin A has been measured in Sertoli cell culture media [32]. No activin B immunoassay is currently available to assess the presence of activin B.

C. Regulation of Inhibin and Activin Synthesis 1. Synthesis Patterns: Clues to Function Inhibin synthesis and secretion vary across the cycle of the seminiferous epithelium of the adult testis. In the rat, the levels of inhibin α subunit mRNA are highest at stages XIII–I and lowest at stages VII–VIII [28, 33–35]. The protein secretion pattern for inhibin in stage-dissected tubule fragments shows a similar, cyclic pattern [35], which is juxtaposed with the maximum synthesis of the βA subunit mRNA and protein between stages VII and XII [36, 192]. The variation in relative proportions of inhibin and activin at different stages of the seminiferous cycle indicate the importance of regulated TGFβ superfamily signaling in the dynamic processes required for spermatogenesis. Inhibin α subunit expression has been detected in Sertoli and Leydig cells; therefore these are the only potential sites of intracellular inhibin synthesis. In addition to observations of the βA and βB subunits in Sertoli, Leydig, and peritubular myoid cells, activin subunit synthesis has been demonstrated in germ cells. Messenger RNA or protein has been detected in gonocytes (βA), spermatocytes (βB and βC), and round

spermatids (βB, βC) [19, 24, 30, 37, 192]. Activin β protein can form dimers with other activin subunits but not with inhibin α subunits in prostate cell lines [38]. Because activin βC subunit dimerization with the activin βA subunit appears to downregulate activin signaling [39] in a prostate cell line, synthesis of this subunit has been postulated to downregulate the amount of bioavailable activin. This is supported by studies of a liver cell line model, in which cotransfection of either βC or βE was shown to antagonize the function of activin βA [40]. The function of these molecules in the testis remains to be explored. The adult Sertoli cell produces predominantly activin B (βB dimers), whereas peritubular myoid cells synthesize activin A (βA dimers). During postnatal maturation of Sertoli cells, the α subunit mRNA level increases as the cells cease proliferation and undergo terminal differentiation. 2. Regulation by FSH FSH and db-cAMP stimulate inhibin α subunit mRNA expression and protein production in day 20 rat Sertoli cells [41, 42], and FSH treatment elevates α subunit mRNA in fragment cultures of 3-day-old rat testis [43]. FSH stimulates inhibin B production to a limited degree without elevating βB subunit mRNA production [42]. This suggests that FSH stimulation of excess α subunit synthesis promotes increased dimerization of the α subunit protein with free βB subunit to form inhibin B at the expense of activin synthesis [44], with the potential for release of free α subunit. Treatment of Sertoli cells in vitro with db-cAMP (but not FSH) stimulates βB subunit mRNA synthesis, whereas phorbol esters and Ca2+ ionophores inhibit its synthesis [45]. The βB subunit mRNA is produced as two transcripts (4.8 and 3.7 kb) that are independently regulated [46]. Their difference in size is due to an extension of the 5′ region. When the 5′-upstream flanking regions of the two transcripts containing the promoter regions are truncated, expression of a cotransfected reporter gene is markedly increased, suggesting the presence of negative regulatory elements in this region. Surprisingly, these promoter regions are unresponsive to both cAMP and phorbol esters. GATA binding sites have been identified within the promoter region of the α subunit and 4.8-kb (but not 3.7-kb) βB subunit genes. These DNA binding sites for GATA protein transcription factors are important in gonadal and erythroid cell development. GATA-1, -4, -6 proteins have been identified in Sertoli cells and GATA-1 and -4 have been shown to both positively and negatively regulate transcription from the 4.8-kb βB subunit promoter [47].

Chapter 14 The Transforming Growth Factor β Superfamily in Sertoli Cell Biology

These findings indicate that inhibin α subunit is predominantly regulated by FSH/cAMP, whereas the more complex regulation of βB subunit expression includes negative and positive regulatory elements in which cAMP and phorbol esters have actions through unidentified transcription sites. GATA proteins may be part of the regulatory process. In studies of both rats and monkeys, the number of Sertoli cells per adult testis has been positively correlated with the levels of inhibin B in serum [18, 48]. This reinforces the concept that FSH action on Sertoli cells drives the negative feedback loop in which pituitary FSH secretion is essentially self-regulating. 3. Other Regulators Local factors have also been implicated in the regulation of inhibin and activin synthesis, including insulin-like growth factor 1 (IGF-1; [41]) and interleukin-1 (IL1; Okuma et al., unpublished). In vitro studies of Sertoli cells from immature rats shows that both IL1α and IL1β promote activin A secretion and decreases inhibin B secretion, whereas FSH opposes these actions. The stage-dependent synthesis of IL1α indicates that it, too, is part of the feedback loop required to support continuous spermatogenesis by driving the cyclic modulation of Sertoli cell protein production.

D. Activin and Inhibin Function in the Testis 1. Impact on Sertoli Cell Proliferation The extent of Sertoli cell proliferation in the fetus and juvenile testis is a primary determinant of adult spermatogenesis. This is due to the observation that each Sertoli cell supports the maturation of a finite number of germ cells [49]. Hence, factors that increase or decrease Sertoli cell proliferation can impact on fertility as it relates to final sperm output. Our understanding of the role of TGFβ superfamily members at the onset of spermatogenesis derives predominantly from studies on rodents, with the rat and mouse being the most frequently studied. At birth, the Sertoli cells are proliferating, whereas the gonocytes remain quiescent in the center of the cord [50, 51]. Within 24 hr in the mouse and 3 days in the rat, the gonocytes reenter the cell cycle and migrate to the cord perimeter [52]. The germ cells begin their differentiation on the background of proliferating Sertoli cells within a rapidly growing seminiferous cord. However, within 15–16 days after birth, Sertoli cells cease mitosis [53, 54], establish basal tight junctions,

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and promote seminiferous tubule lumen formation by initiation of apical fluid secretion. The duration and extent of Sertoli cell proliferation during this prepubertal period is a crucial determinant of fertility in the adult. In the immature testis, FSH stimulates Sertoli cell proliferation and the onset of Sertoli cell maturation, and this is stimulated by activin in a developmentally regulated fashion. The work of Boitani et al. [55, 56] first identified this change in rat Sertoli cell proliferation using in vitro cultures of testis fragments from day 3, day 9, and day 18 postpartum animals. Although Sertoli cell proliferation at day 3 was clearly dependent on the presence of FSH and unaffected by exogenous activin, maintenance of proliferation on day 9 required the addition of both activin A plus FSH. At day 18, there was no measurable proliferation of these cells, in vivo [53, 54]. Boitani and colleagues have extended their studies using additional time points and analysis of activin receptor subunit expression levels to identify the elevation of the 6-kb ActRII mRNA between days 7 and 9 postpartum, in comparison to other ages [29]. This has been confirmed in studies using highly purified Sertoli cell preparations [57], and these data demonstrate an important role for activin A in driving events at the end phase of Sertoli cell proliferation. Additional data [57, 58] indicate that peritubular cells may be the source of activin A at this time. In a normal testis, the balance of signals between FSH and inhibin appears be important for cessation of Sertoli cell proliferation. The expression of the cell cycle inhibitor p27Kip1 commences at the time when the Sertoli cells undergo terminal differentiation [59]. In normal tissue, inhibin acts as a “tumor suppressor” by promoting association of p27Kip1 with cylinD2/E1 and Ck4/2, which keeps these cyclin complexes inactive. Mice lacking inhibin α subunit develop Sertoli cell and granulosa cell tumors starting around 4 wk postpartum [60]. In the absence of inhibin, the absence of negative feedback to the pituitary results in elevated serum FSH and elevated testicular cyclin D2 levels. Double knockout mice lacking both p27Kip1 and inhibin α subunit genes have elevated cyclin D2 and Cdk4 mRNA levels. These animals exhibit sustained proliferation and more rapid tumor growth, a finding interpreted as reflecting the increased formation of active cyclin D2/Cdk4 complexes that would in turn promote active cyclin E1/Cdk2 complex formation and sustained cell cycle progression [61]. In addition to the temporally discrete stimulatory effect of activin βA on immature rat Sertoli cell proliferation in vitro, activin A also regulates androgen receptor expression and FSH-induced aromatase activity [62].

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The interactions between activin and androgens was explored in vivo by crossing inhibin α −/− mice with testicular feminized male (tfm) mice that have a nonfunctional androgen receptor [63]. The compound mutant mice share several characteristics with the inhibin α −/− mice, including elevated activin secretion from gonadal tumors and cachexia-like wasting syndrome. However, the life span of the mice is somewhat prolonged (by ≥5 wk) in the absence of the functional androgen receptor, and the frequency of hemorrhagic phenotype in the testicular tumors is reduced. These data indicate that androgens play a modest role in regulating the development of tumors caused by the absence of inhibin. 2. Postmitotic Sertoli Cells Our understanding of how inhibin and activin act and are regulated within the adult testis is clearly incomplete. In a genetically engineered mouse lacking the inhibin α subunit gene, no effect on testicular development or adult fertility was observed [60], suggesting that the physiological role of inhibin is either limited or is at least partially met by another protein. However, the subsequent development of stromal cell tumors indicates that inhibin can act as a tumor suppressor. Activin may function to maintain the immunosuppressed nature of the interstitium in the adult testis. Activin has been shown to suppress proliferation of peripheral T lymphocytes [64], an activity that would limit the influence of lymphocytes when presented with novel antigens such as haploid germ cell products, as might occur during testicular damage.

cell function. In addition, the absence of the βB subunit gene does not affect male fertility [70]. When the βA subunit gene is absent, mice die immediately after birth due to respiratory complications [69], so these animals are not suitable for investigating βA protein function in the postnatal testis. To circumvent this problem, Brown et al. [71] inserted the activin βB subunit gene coding sequence into the βA subunit locus. Relying on the estimate that activin β has an approximately 10-fold lower affinity than does activin α for its receptor [72], this strategy enabled derivation of a mouse line with reduced activin bioactivity at sites where activin A is normally expressed. This would occur on the background of normal activin βB subunit production. In contrast to the activin βA subunit–/– mice, most of these animals survive to adulthood. In these BK/BK mice, the onset of male fertility is delayed due to a delay in completing the first wave of spermatogenesis, and there appears to be an early accumulation of spermatogonia. This study supports the concept that activin α is important in the first wave of spermatogenesis. It is important to note that the activin β subunit mRNA is downregulated during the first week of postnatal life in mice and rats [71, 192]. Detailed analyses of fetal and newborn rats revealed that the quiescent gonocytes synthesize activin βA subunit mRNA and protein in the fetal testis [24]. They appear to store the protein until the time of their transformation into spermatogonia and then begin to synthesize the activin antagonists, follistatin and Bambi [43]. These findings have been proposed to indicate that the commencement of spermatogonia differentiation requires a downregulation of signaling by activin or by other TGFβ superfamily members.

3. Impact on Germ Cells Addition of activin increases the rate of apoptosis by isolated mouse primordial germ cells in culture [65]. In the early postnatal rat testis, activin stimulates the number of gonocytes when present during cultures (1 or 3 days) of day 3 rat testis fragments [24], though whether this observation reflects increased survival or proliferation rates has not been determined. Activin has been reported to enhance proliferation of rat spermatogonia in cocultures with day 20 Sertoli cells [66], whereas inhibin enhanced DNA synthesis of Chinese hamster spermatogonia [67]. The use of transgenic and knockout mice to pinpoint the roles of activin and inhibin in biology has reinforced our understanding that they are key determinants of testicular growth and function (see Table 14.3; reviewed in [68, 69]). As mentioned earlier, deletion of the inhibin α subunit gene does not affect germ

IV. MÜLLERIAN INHIBITORY SUBSTANCE A. MIS Structure and Signaling MIS (also known as anti-Müllerian hormone, AMH) is a key factor in sexual differentiation produced by Sertoli cells [73]. It is a disulfide-linked homodimer of 140 kDa. The gene is located on the short arm of chromosome 19 in humans. MIS signals through a specific receptor (MISR1) and either ActRIA or BMPRIB as type II receptors [74]. Its signal is transduced through R-Smad 1, 5, or 8. Several sites for transcription factor binding within the MIS promoter regions have been identified that bind SF-1, Sox9, and GATA-4 [75]. MIS expression is restricted to Sertoli cells, and levels of synthesis are much higher in fetal and neonatal testes than in differentiated Sertoli cells.

Chapter 14 The Transforming Growth Factor β Superfamily in Sertoli Cell Biology

B. MIS Function in the Testis The key event in transformation of the indifferent gonad into a testis is the expression in somatic cells of the Sry gene [76, 77]. This signal is essential and sufficient for their differentiation into Sertoli cells, and it triggers a cascade of changes in Sertoli cell gene expression (reviewed in [78]). One downstream target of Sry is the gene encoding MIS. Sertoli cell expression of MIS is vital for regression of the Müllerian ducts that contain the precursors of the female uterus, fallopian tube, and upper vagina, acting through receptors (BMPR1a [74]) on mesenchymal cells of the Müllerian ducts. Male mice lacking the MIS gene are sterile, though not due to any apparent defect in spermatogenesis or behavior; the efferent ducts are blocked through retention of the female ducts in mature males, and sperm are thus not ejaculated during coitus [73]. Leydig cell hyperplasia was observed in foci in 27% of these MIS–/– mice, which is indicative of a potential role for MIS in governing Leydig cell proliferation. MIS has been reported to reduce proliferation of progenitor, prepubertal Leydig cells in day 21 postpartum rat [79] and to inhibit testosterone production in fetal and adult Leydig cells [80, 81]. MIS was also shown to block cAMP-stimulated aromatase activity in Sertoli cells by reducing levels of the aromatase mRNA [80]. Expression of MIS is a hallmark feature of the Sertoli cell lineage in the fetal gonad. In the search for factors that direct gonadal development into either a testis or an ovary, the enzyme prostaglandin D2 (PGD2) synthase was identified as being more highly expressed in the testis relative to the ovary at embryonic day 12.5. Exposure of cultured mouse urogenital ridges to PGD2 induced MIS expression in somatic cells of a female gonad at the onset of sex determination [82]. This indicates that local synthesis of PGD2 by somatic and germ cells in the male may provide a key positive feedback mechanism driving differentiation of the gonad into a testis at the time of gonad gender specification.

V. BONE MORPHOGENETIC PROTEINS A. BMP Structure and Signaling Within the BMP family of ligands, members of two well-characterized classes are distinguished by high sequence homology in the carboxy terminus of the mature protein. The 60A class includes BMP5, BMP6, BMP7, BMP8A, and BMP8B. The DPP class includes BMP2 and BMP4 (reviewed in [83, 84]). Similar to other members of the TGFβ superfamily of ligands, the

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BMPs are synthesized as large dimeric proteins that are cleaved by proteases around the time of secretion. The 21- to 25-kDa mature protein appears to bind both type I and type II receptor units simultaneously, in contrast with the activins and TGFβs that first bind the type II receptor to recruit the type I receptor subunit. Regulation of BMP signaling by many extracellular proteins has been reported (reviewed in [85]).

B. BMP Function in the Testis Even though a testis can form without germ cells, germ cells do not form in the absence of BMPs. Around 6.0 to 6.5 days postcoitus (dpc), cells of the mouse embryo epiblast require both BMP4 and BMP8b signals to enter the germ cell lineage, after which they proliferate and migrate to the gonadal ridge. This commitment to the germ cell lineage precedes the formation of Sertoli cells, and it has recently been used to drive germ cell formation from embryonic stem cells [86, 87]. Deletion of the components of the BMP signaling cascade leads to the absence of primordial germ cells, as observed in mice lacking BMP4, BMP8b, Smad1, and Smad5 (Table 14.3) [88–91]. Sertoli cell-derived BMP4 appears to be an important driver of germ cell differentiation at the onset of spermatogenesis in the postnatal testis. BMP4 mRNA production in isolated preparations of Sertoli cells was measured to be highest at 4 days postpartum (dpp), declining rapidly within 1 wk. In vitro studies of mouse germ cells revealed that incubation with BMP4 can increase the level of activated c-kit on spermatogonia entering the committed phase of differentiation, a step required ultimately for their progression into meiosis [92]. It is possible that other TGFβ superfamily ligands act through a similar mechanism to govern germ cell differentiation in the postnatal and adult testis, with the rate of germ cell progression regulated by local production of antagonists. Bambi and follistatin (Section VIII) are two potential candidates. An analogous function for BMPs in directing differentiation of murine stem cells has been uncovered through a germ cell transplantation assay, which is the only available method to assess their stem cell potential [93]. Culture of mixed testis cell populations from cryptorchid mice with BMP4 prior to transplantation into a recipient host reduced the number of colonies formed when compared to untreated samples. This indicates that BMP, similar to activin (Section III.D.3), reduces the germ cell stem cell capacity, in contrast to GDNF which supports it (Section VII). In light of the demonstrated effect of BMP4 on c-kit expression, it appears that BMPs drive germ cells out of the stem cell state and into the differentiation pathway.

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Kate L. Loveland and David M. Robertson TABLE 14.3 Mice with Reproductive Phenotypes Due to Genetically Modified Expression of TGFβ Superfamily Signaling Components

Gene

Reproductive phenotype

References

Inhibin α knockout (Inha)

Gonadal and adrenal tumors

[60]

Inhibin α/p27 double knockout

Enhanced tumor progression relative to inhibin α knockout

[61]

Inhibin α/cyclin D2 double knockout

Decreased tumor progression relative to inhibin α knockout

[179]

Activin βB subunit knockout (Inhbb)

Reproductive abnormalities in females

[181, 182]

Activin βB subunit inserted into activin βA subunit knockout at βA locus (BK)

Delayed fertility onset in males

[71]

Activin Type IIA receptor (ActRIIA)

Delayed fertility in males, reduced testis size; females infertile; 25% perinatal lethal; Sertoli cell and germ cell effects

[68, 183, 184]

ActRIIA/FSHα subunit double knockout

Reduced spermatogonia, males fertile

[185]

Activin βA transgene expression in spermatocytes; metallothionein promoter

Male infertility; spermatogenesis disruption 3 wk postpartum

[186]

MIS knockout

Leydig cell hyperplasia; males sterile due to blocked sperm efflux

[73]

MIS type II receptor knockout

Same as MIS knockout

[187]

BMPR1A knockout, specifically in Müllerian ducts

Pseudohermaphrodite males have uteri and oviducts

[74]

BMP2 knockout

Primordial germ cell (PGC) deficient

[188]

BMP4 knockout

PGC deficient; male infertility

[88]

BMP8A knockout

Spermatogenesis failure after initiation in ~50% adult males

[95]

BMP8B knockout

PGC deficient; male infertility

[89, 94]

BMP7/8B double knockout

Male infertility

[96]

GDNF knockout heterozygotes

Male infertility; loss of spermatogonia

[127]

GDNF overexpression transgenic; hEF-1α promoter

Male infertility; spermatogonia do not differentiate; testicular tumors formed

[127]

Smad 1 knockout

PGC deficient

[90]

Smad 5 knockout

PGC deficient

[91]

Smad 4 transgenic; MIS promoter

Male infertility due to germ cell loss and Leydig cell hyperplasia in one line

[189]

GDF7 knockout

Abnormal seminal vesicles; male infertility, though sperm appeared normal

[190]

Follistatin transgenic lines; metallothionein promoter

Some lines with spermatogenic arrest; Leydig cell hyperplasia; some lines with female infertility

[191]

Note: In the case of mouse strains with gene knockouts relating to this family, the phenotype is embryonic lethal and does not shed light on the role of these factors in reproduction. Reviews in [2, 83] provide comprehensive discussion of many of these phenotypes, as well as phenotypes relating to other physiological systems. See also [68, 69] for discussion.

Evidence for the function of other BMPs has been derived from gene targeting studies in mice (Table 14.3). The deletion of BMP8B or BMP8A can lead to male infertility, with lesions noted in primordial germ cell numbers (BMP8B), in germ cell proliferation at the onset of spermatogenesis, and in spermatocyte survival in the adult testis [94, 95]. The BMP8B transcript is synthesized by round spermatids in the adult testis, and these cells may therefore be contributing to the pool of regulatory cues that drive differentiation of the

less mature germ cell types in the seminiferous epithelium. The expression pattern of BMP7 in the mouse testis overlaps with that of BMP8A and BMP8B in the juvenile and adult, and evidence for functional redundancy has been derived from analysis of BMP7/BMP8B double mutants [96]. Currently, no comprehensive picture exists of the sites of production of the BMPs or BMP receptor subunits in the developing or adult testis, and knowledge about Sertoli cell expression is particularly deficient.

Chapter 14 The Transforming Growth Factor β Superfamily in Sertoli Cell Biology

VI. TRANSFORMING GROWTH FACTOR β PROTEINS A. TGFβ Structure and Signaling Three mammalian genes encode TGFβ proteins: TGFβ1, TGFβ2 and TGFβ3. Initially synthesized as precursor dimeric glycoproteins, the typical hydrophobic signal sequences are first removed to yield the pro-TGFβ dimer. A second cleavage step separates the aminoterminal 112 amino acids, named the latency-activated peptide (LAP), from the carboxy-terminal region that will become the mature 25-k signaling molecule. The mature protein is inactive when complexed with the LAP, which in turn is covalently bound to a binding protein, termed the latent TGFβ binding protein (LTBP). This complex formation appears to be important for folding, secretion, and storage of TGFβ. Activation by several possible mechanisms results in the release of mature TGFβ from these binding proteins. Pathways implicated in TGFβ activation involve proteases and protease activators known to be produced by Sertoli cells [97, 98], acidification of the extracellular milieu, radiation, reactive oxygen species, and integrin α5β6 [99]. The mature proteins are 25-kDa disulfide-bonded dimers. Though structurally quite similar, the expression profiles of the three proteins, and hence factors regulating their expression, are distinct. In general, TGFβ1 subunit mRNA is induced by early response genes, whereas TGFβ2 subunit and TGFβ3 subunit mRNAs are under hormonal and other controls related to development (reviewed in [98]; discussed further in Section VI.C). In addition to the TGFβ type I and type II receptor subunits (TβR-I and TβR-II) that are structurally common within the superfamily, the TGFβs also employ a type III coreceptor subunit, betaglycan, an integral membrane protein. While TGFβ1 and TGFβ3 bind the TβR-II with high affinity in its absence, TGFβ2 binding requires the presence of betaglycan in order to signal. Thus the bioactivity of this specific TGFβ protein is affected by local production of betaglycan. TGFβ signaling can be impeded by inhibin binding to betaglycan (Section VIII.A). In contrast, another TGFβ type III receptor integral membrane protein, endoglin, binds TGFβ1 and TGFβ3 in the presence of the TβR-II, but TGFβ2 does not. Endoglin has also been reported to bind activin A, BMP2, and BMP7 (reviewed in [99]).

B. TGFβ Function in the Testis TGFβ1 mRNA and protein were first detected in Sertoli cells and peritubular cells isolated from the postnatal rat testis, and they were reported to affect

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the migratory and growth characteristics of peritubular cells [100]. Messenger RNAs encoding each TGFβ1, TGFβ2, and TGFβ3 were then identified in peritubular cells and Sertoli cells from the postnatal rat testis and their expression patterns assessed using nuclease protection assays [101]. Total testicular levels of all three ligand mRNAs were highest at birth and declined at or during the onset of spermatogenesis in total testis, whereas a unique pattern of expression for each was reported in Sertoli cells during their maturation. Messenger RNA encoding all three receptor subunits was readily detected in the immature rat testis in both somatic and germ cells, with levels of TβR-I and TβR-II declining through development [102]. In subsequent studies, TGFβ1 protein expression was readily detected by immunohistochemical analysis in the fetal rat testis, appearing to be exclusively expressed in Sertoli cells between 13.5 and 14.5 dpc. This signal was reduced and remained faint from 18.5 dpc onward into postnatal life. Fetal Leydig cells express TGFβ1 protein from 16.5 dpc, as do Leydig cells at all subsequent stages [103]. No signal was detected in germ cells or peritubular cells in the fetus and developing postnatal testis. TGFβ2 protein has been detected in mouse Sertoli cells between 13.5 and 18.5 dpc in the rat [104], with the signal being particularly strong in the first 3 days following the onset of cord formation in the developing gonad. The detection of TGFβ1 and TGFβ2 in fetal (13.5-dpc) and newborn rat Sertoli cells has also been independently reported [105], and TGFβ1 protein has been detected in isolated rat Sertoli cells [106]. TGFβ3 has not been detected in Sertoli cells, but was observed in fetal Leydig cells, gonocytes, and spermatogonia in the immature rat testis [107]. The TGFβ type I and type II receptor proteins were visualized in gonocytes of the rat testis from 13.5 dpc to 3 dpp and in the Leydig cells from 16.5 dpc onward, but no signal was reported in Sertoli cells at these ages [108]. Incubation with TGFβ1 resulted in reduced cord formation in fetal testes (between 13.5 and 14.5 dpc) and inhibition of the epidermal growth factor- and fetal calf serum-stimulated growth of the neonatal rat testis [105]. Culture with TGFβ1 inhibited basal and LH-stimulated testosterone production by rat fetal Leydig cells, probably by reducing mRNA levels of key steroidogenic enzymes [109]. A similar effect for TGFβ3 was also measured in rat Leydig cells in vitro [107]. In accord with these results was the increase in steroidogenic capacity and synthesis of steroidogenic enzymes that resulted from inhibition of TGFβ1 synthesis in cultured pig Leydig cells by administration of an antisense oligonucleotide [110]. An additional effect of TGFβ1 on cell morphology in vitro was reported, and this may reflect the influence of TGFβ1 on cord formation in the early testis [109].

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In vitro culture of testis fragments has been used to examine the impact of exogenous TGFβ on male germ cell development. Although nonproliferating germ cells in the fetal rat testis (16.5 dpc through 2 dpp) were unresponsive to TGFβs, those germ cells that had reentered the cell cycle after birth underwent apoptosis in the presence of TGFβ1 or TGFβ3 [111]. In contrast to the impact of TGFβ, exogenous activin A increased the number of gonocytes at this 3-dpp time point [24]. These data indicate that local production of TGFβ superfamily members and signaling components regulates the onset and progression of spermatogenesis, through its impact on both germ cells and Sertoli cells. Several lines of evidence indicate that TGFβ signaling affects protein synthesis by Sertoli cells in the first wave of spermatogenesis. Most observations were derived from studies with primary cultures of day 19–20 rat Sertoli cells and peritubular cells. The addition of exogenous TGFβ2 to cocultures of Sertoli and peritubular cells, but not to monocultures, promotes cellular aggregation in vitro [112]. These data suggest that local secretion of TGFβ2 by somatic cells may regulate cord formation in vivo during this period of rapid growth, similar to its proposed role in the fetal testis. In another study, production of plasminogen activator inhibitor-1 was stimulated by addition of TGFβ1 to postnatal Sertoli cell cultures [113]. A structure vital to both differentiating and adult Sertoli cell function is the tight junction (reviewed in [114]), which maintains the distinct basal and luminal compartments required for germ cell maturation. The potential influence of TGFβs on the establishment and maintenance of this structure was examined using in vitro cultures of 20day-old rat Sertoli cells, with junctional integrity assessed by measuring transepithelial resistance (TER; [115]). As tight junctions formed and TER values increased, TGFβ2 subunit and TGFβ3 subunit mRNA levels decreased. Once these junctions formed, the levels of these mRNAs increased. The addition of exogenous TGFβ3 to cultures affected the deposition of the key junctional proteins occludin, zonula occludens-1, and claudin-11 during tight junction formation, and it also disrupted those that had already formed. An in vivo study using cadmium chloride to disrupt the blood–testis barrier in rats has revealed that TGFβ3 regulates occludin synthesis and the blood–testis barrier formed by these tight junctions via the p38 MAP kinase pathway [116]. These data convincingly demonstrate the influence of TGFβ proteins in the regulation of this crucial aspect of Sertoli cell function.

C. Regulation of TGFβ Synthesis Growth of the fetal testis is regulated by an interplay of hormones and locally produced factors (see [117]

for an excellent review). Based on the functional data cited earlier, factors that influence TGFβ synthesis are likely to have a profound impact on testis development. The secretion of TGFβ1 in the day 20.5 fetal rat testis has been attributed to both Sertoli cells and Leydig cells, as it was observed to be under the combined regulation of LH and FSH [118]. FSH has been shown to suppress expression of TGFβ2 [101, 112], an interaction that has been proposed to affect cord formation at the onset of rat spermatogenesis. The addition of epidermal growth factor to cultures of newborn rat testis increased TGFβ1 mRNA and decreased TGFβ3 mRNA after 24 hr [105]. The addition of retinoic acid to in vitro cultures of day 0 rat testes stimulated production of TGFβ3 subunit mRNA within 24 hr, and TGFβ1 and TGFβ2 subunit mRNAs were increased after 72 hr in the presence of both retinoic acid and retinol [119]. This may be a mechanism that provides a balance to stimulatory signals in the fetal and newborn testis, because the addition of retinoic acid in this study inhibited the actions of FSH, epidermal growth factor, and calf serum, each of which stimulates growth in its absence. Receptors for retinoic acid have been identified on interstitial and germ cells but not on Sertoli cells [119, 120], which highlights that a complex integration of regulatory signals coming from different cell populations must affect testis growth and function. Evidence suggests that TGFβ protein production is regulated by germ cells. Isolated rat Sertoli cells synthesize a TGFβ1 subunit mRNA transcript but secrete negligible amounts of this protein in vitro. The addition of enriched populations of spermatocytes or round spermatids results in increased expression of this gene and secretion of TGFβ1 protein [106].

VII. GLIAL-DERIVED NEUROTROPIC FACTOR A. GDNF Structure and Synthesis GDNF is a relatively distant member of the TGFβ superfamily of proteins with the characteristic cysteine knot structure of three intermolecular double bonds and one intramolecular bond present in the dimeric ligand [121]. Along with its reasonably close relative, neurturin (NRTN), these Sertoli cell–derived products have been shown to influence germ cell physiology at several stages of development. Two other family members, artemin and persephin [122], have not been identified in the testis to date. GDNF signaling is mediated by a glycoslylated GPI-linked ligand binding subunit, either GDNF family receptor α-1 (GFRα-1; also known as TrnR1) or

Chapter 14 The Transforming Growth Factor β Superfamily in Sertoli Cell Biology

α-2 (GFRα-2; also named TrnR2). Two signaling pathways for GDNF have been described, one dependent on the membrane-bound protein tyrosine kinase c-ret and the other independent of c-ret. The GFRα-1/ c-ret complex mediates signaling of both GDNF and NRTN. GFRα-2 appears to bind preferentially to NRTN, whereas GFRα-1/ret appears to bind both ligands with equivalent affinity [123]. GDNF binding to GFRα-1 shows positive cooperativity of ligand– receptor interactions at low ligand concentrations and negative at high ligand concentrations [124]. This was proposed to illustrate how local GDNF concentrations will specify the signal received by the target cell, with full receptor occupancy occurring at biologically relevant concentrations, and signal attenuation occurring at higher concentrations. Signaling in the absence of c-ret has also been described, and a mechanism by which this occurs was recently uncovered. GDNF can bind to GFRα-1 and signal through N-CAM, and the presence of GDNF can reduce homotypic binding between N-CAM molecules on adjacent cells [125]. Whole mount in situ hybridization analysis of mouse identified GFRα-1, GFRα-2, c-ret, NTRN, and GDNF mRNAs in the embryonic mouse testis at 14 dpc [126]. All three receptor subunits were undetectable at 16 and 18 dpc, times when the male germ cells are not proliferating and Sertoli cell proliferation is escalating, while the ligands remained at detectable levels. GDNF mRNA was detected in Sertoli cells of the newborn mouse, and its receptor subunits, c-ret and GFRα-1, were observed in a subset of the spermatogonia present at 2 wk postpartum [127]. The receptor-positive spermatogonia were further characterized as lacking immunoreactive c-kit protein. Because the c-kit receptor tyrosine kinase is a marker for differentiating spermatogonia [128], these data indicate that there are two functionally distinct spermatogonial populations present in the developing testis at this time. GFRα-1 protein has been visualized using immunohistochemistry on spermatogonia in the 6-dpp mouse testis [129], and GFRα-1 and ret have been observed in a relatively high proportion of the undifferentiated spermatogonia present in five different conditions of germ cell arrest [130]. In some studies of the adult rodent testis using in situ hybridization, GFRα-1, GFRα-2, and NTRN mRNAs have been detected, whereas GFRα-3, c-ret, and GDNF have not [126, 131]. In contrast, GFRα-1 mRNA and protein were detected in adult human Sertoli cells, whereas GFRα-2 was found in Leydig cells [132]. This apparent difference between the rodent and the human may reflect species-specific differences in the function of these pathways or in the reagents and methods used to detect them. NTRN mRNA was detected in zygotene and early pachytene

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spermatocytes (stages X–XII and I–II, respectively [131]). The c-ret mRNA autoradiographic signal detected in this study was most intense at stages X–I, and it appeared to be in Sertoli cells. N-CAM on gonocytes and Sertoli cells has been implicated as a crucial mediator of interactions required at the onset of spermatogenesis, as the gonocytes migrate from the center to the periphery of the developing cord [133]. Messenger RNA encoding the receptors for artemin and persephin, GFRα-3 and GFRα-4, respectively, have been identified in the testis, but the cellular localization of these receptors or their preferred ligands is unknown [122, 134]. GDNF immunoreactive material was detected in peritubular cells and in cells of the microvessels, while GFRα-1 was detected in interstitial endothelial cells and in spermatocytes [135]. Based on its ability to regulate permeability across the blood–brain barrier, these authors proposed that local production of GDNF affects function of tight junctions in the testicular endothelium.

B. GDNF Function in the Testis GDNF has profound effects on events in the early postnatal testis, with effects on both Sertoli and germ cells described. A fourfold stimulatory effect of GDNF on 6-day-old rat Sertoli cell proliferation was observed following 3 days of testis fragment culture with recombinant rat GDNF (100 ng/mL) only when rat FSH (200 ng/mL) was present. This effect was blocked by the addition of specific antibodies [136]. In contrast, overexpression of GDNF in a mouse transgenic model did not appear to affect Sertoli cell proliferation, indicating that the observed in vitro effect may be indirectly mediated through the actions of GDNF on germ cells where its receptors are localized [127]. The in vivo effects of overexpression and deletion of GDNF have been studied in mouse models. In mice heterozygous for GDNF (GDNF+/–), the reduced synthesis of this protein results in a loss of germ cells in the postnatal testis. Overexpression of human GDNF under the control of the human testis-specific translation elongation factor-1α promoter in a transgenic mouse causes accumulation of undifferentiated spermatogonia, ultimately causing spermatogenic disruption [127]. Malignant testicular tumors of aneuploid cells bearing germ cell markers develop in these mice, indicating that cells related to these undifferentiated spermatogonia may contribute to the formation of testicular tumors in humans. It is of interest that targeted deletion of the NTRN gene in mice appears to reduce survival of some spermatocytes and spermatids, but its effect on spermatogenesis appears to be lost shortly after the first wave of spermatogenesis [137].

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Our emerging understanding of GDNF in testicular biology is that this Sertoli cell product is required to maintain spermatogonial stem cells. This has been directly assessed through in vitro culture of spermatogonia in the presence of various factors, followed by transplantation to assess stem cell activity [93]. Cells from adult mouse testes rendered cryptorchid to enrich for stem cells were cultured on various feeder layers for 7 days. Following transplantation to nude mouse recipients, the extent of donor cell colonization was measured after 2 months. Culture with GDNF increased the stem cell activity in these cultures to 160% of untreated controls, whereas incubation with either activin (to 30% of controls) or BMP4 (to 40% of controls) reduced the stem cell potential of the population. The observation that GFRα-1-positive cells do not express c-kit [137] is relevant in light of the finding that murine spermatogonial stem cells lack surface c-kit receptors [138]. We now understand the functional significance of these two spermatogonial populations, the former as the stem cell population, and the latter as those committed to differentiate into sperm. In another intriguing twist, germ cells from the transgenic mice expressing human GDNF in spermatogonia were transplanted into a recipient testes prepared by irradiation [139]. This transfer recapitulated the phenotype seen in the donor mice, with clusters of ret-positive spermatogonia. Another approach to generating elevated numbers of stem cells in vivo involved electroporation of a cDNA encoding human GDNF into immature (12 dpp) or germ cell-depleted mice [140]. In each case, the expression of GDNF in these animals was correlated with expansion of the population of undifferentiated spermatogonia with stem cell characteristics. These models should provide relatively convenient sources of germ cells for analysis of spermatogonial stem cell biology.

VIII. ANTAGONISTS OF TGFβ SUPERFAMILY SIGNALING A. Inhibin Until recently, our knowledge of inhibin’s function related predominantly to its capacity to block activin signaling. Our understanding of its ability to act more broadly to impact on signaling by additional TGFβ superfamily ligands has been further expanded to include investigation of whether inhibin is a bona fide signaling molecule in its own right. Based on its capacity to antagonize activin signaling in the pituitary and other tissues, it had been presumed that inhibin antagonized activin’s action simply by

competing for occupancy of the ActRII receptors. It was also recognized, however, that since inhibin’s ability to compete with activin was poor, having about a 10-fold lower affinity for receptor binding, that other factors may be responsible for its physiological role. Studies by Lewis et al. [141] showed that an accessory binding protein called betaglycan (or TGFRIII), previously shown to potentiate TGFβ2 bioactivity, was able to bind inhibin (Kd 600 pM) but not activin, forming an effective competitive inhibitor of activin binding to its receptor. Inhibin, again in association with betaglycan, has also been shown to block BMP binding to the BMPRII, a receptor subunit to which activin can also bind [142]. These data were striking in that the IC50 for inhibin’s inhibitory effects on BMP action in HepG2 cell line was decreased 100-fold into the low pM range. These studies strongly suggest that inhibin, in conjunction with betaglycan, is a potent antagonist of both activin and BMP action. Betaglycan has been found in adult Leydig and in germ cells [30, 102, 143, 144]. These observations have yet to be reconciled with situations in which inhibin and activin both provoked a similar response, for example, in stimulating steroidogenesis in porcine Leydig cells [141] [145]. Interactions with other signaling regulators may contribute to this phenomenon. The mechanisms whereby inhibin exerts its effects remain unclear, and the current understanding is that inhibin’s impact within the testis may be restricted to the downregulation of signaling by activin and some BMPs. However, attempts to identify an inhibin binding protein with signaling capacity are ongoing. The presence of inhibin in stage-dissected tubule segments reduced in vitro DNA synthesis in intermediate spermatogonia and in preleptotene spermatocytes, whereas the addition of activin increased DNA synthesis in these cells. Binding of 125I-labeled inhibin to germ and somatic cells has been reported [146]. Findings such as these have spurred interest in searching for an inhibin signaling receptor. Additional studies have shown that inhibin was able to bind to Leydig and Sertoli cell lines (TM3 and TM4, respectively) with very high affinity (Kd 50 pM) yet showed little or no evidence of activin binding [147]. Inhibin binding proteins have been identified on a variety of other cell types, including pituitary, bone, and adrenal [148]. These findings suggest that other cell surface molecules are involved in mediating the biological impact of inhibin. Crosslinking studies with radiolabeled inhibin identified at least five proteins on the Sertoli and Leydig cell lines, two of them being full length and shortened forms of betaglycan, with the remaining three being unknown. No specific inhibin

Chapter 14 The Transforming Growth Factor β Superfamily in Sertoli Cell Biology

receptor has been identified to date nor have genomic searches identified an inhibin-specific serine-threonine kinase [149].

B. Follistatin Follistatin is another protein isolated on the basis of its ability to influence pituitary FSH secretion ([150, 151]; reviewed in [152]). The product of a gene family unrelated to the TGFβ superfamily, follistatin is produced as several protein isoforms that block the binding of activin and to a lesser extent, BMP, to its receptor ([153, 154]; reviewed in [85]). Two mRNA forms are produced by alternative splicing, one including and one excluding exon 6 [155, 156]. The absence of exon 6 leads to the generation of the 288-amino-acid isoform (FS288), which is generally found in association with cell surface proteoglycans such as heparin sulfate. Inclusion of exon 6 results in formation of FS315, the soluble protein isoform predominant in serum [157]. Additional variations in the follistatin protein configuration result from glycosylation and proteolysis. Follistatin mRNA and protein have been detected in Sertoli cells at all stages of postnatal testicular development [24, 158], but its absence from the early fetal testis, in contrast to its presence in the fetal ovary, has been noted [159–161]. Follistatin has also been identified as having the capacity to block BMP signaling, most notably BMP15 in the ovary [162], and its local impact on Sertoli cell development and function has yet to be fully elucidated.

C. Bambi Bambi (BMP and activin membrane-bound inhibitor) is a structural homologue of TGFβ type I receptors that lacks the intracellular serine/threonine kinase domain. Bambi is structurally conserved from Xenopus to humans, and it was shown to form a complex with type II receptor subunits that could block signal transduction in the presence of TGFβ, activin, and BMPs [163]. Hence, cells that synthesize Bambi protein would be predicted to show decreased responsiveness to stimulation by many of the TGFβ superfamily members. Expression of Bambi mRNA in the postnatal rat testis has been observed in Sertoli cells of all ages, and it is regulated during germ cell development. It appears to be initially expressed within germ cells as the gonocytes transform into spermatogonia [43]. This upregulated expression coincides with the reduced activin A protein and increased follistatin mRNA and protein observed shortly after birth in developing germ cells [24, 71, 192]. In cultures of 3-dpp rat testis

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fragments, Bambi mRNA expression was downregulated by the addition of activin A [43]. This mRNA was shown to be upregulated in mouse embryonic fibroblasts by culture with BMP4 [164], another factor expressed by Sertoli cells at the onset of spermatogenesis [92] when Bambi mRNA is elevated in germ cells. Bambi mRNA appears highly expressed in pachytene spermatocytes and round spermatids, but it is absent from elongating spermatids. Its localization to these meiotic and postmeiotic cells may relate to a role in regulating TGFβ superfamily ligand action in the adult testis.

D. Crim1 Crim1 is an antagonist of BMP signaling that acts by binding to BMPs intracellularly, slowing their release from the cell [165]. Similar to Bambi, Crim1 is synthesized as a transmembrane protein, and it is structurally related to chordin, due to the presence of an IGF-binding protein motif and multiple cysteine-rich repeats. Crim1 mRNA shows striking gender-specific expression in the fetal mouse gonad at 13.5–17.5 dpc, and it appears to be localized to Sertoli cells [166]. This expression pattern presents an interesting contrast to the upregulation of follistatin in the ovary at the same time [161].

E. α2-Macroglobulin The protease inhibitor α2-macroglobulin is another Sertoli cell–derived secretory product that can affect influence signaling [167]. It has the capacity to bind inhibin, activin, and follistatin and may thereby change the effective local concentration of each of these factors [168], although there is little evidence in support of this possibility.

IX. CLINICAL RELEVANCE OF INHIBIN AND ACTIVIN It has been recognized for many years that serum FSH levels are elevated in men with infertility, and this has been attributed to effects of dysfunction of the spermatogenic process. Based on the presumed biological role of inhibin, it was anticipated that there should be a concomitant fall in serum inhibin associated with these elevated serum FSH levels. Early studies [169] investigating the relationship between serum total inhibin forms and FSH in fertile and infertile men showed no such correlation. The development of serum inhibin assays specific for inhibin A, B, and pro-αC [170, 171] have enabled this relationship to be further explored.

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Inhibin A is undetectable in adult male serum. Serum inhibin B and pro-αC levels have been detected throughout prenatal and postnatal life. Inhibin B is primarily if not totally produced by human Sertoli cells, although pachytene spermatocytes may be a contributing source by producing the inhibin βB subunit [17]. Pro-βC is produced by Leydig and Sertoli cells. In the prepubertal boy there is a positive correlation between serum FSH and inhibin B similar to that seen in in vitro studies, which may reflect the increase in Sertoli cell number during this period. In the adult male, serum inhibin B is inversely correlated with FSH in both fertile and infertile men [14, 15]. However serum inhibin, like FSH, also correlates with testicular size and sperm count. Additional studies have shown that serum inhibin is elevated with FSH but not LH treatment and that attempts to reduce gonadotropin levels by steroid treatment in normal men have led to a partial fall (40%) only in inhibin levels [172]. On the other hand, severe disruption of the testis by chemotherapy and X-irradiation led to a very rapid and marked depression of serum inhibin levels [173]. These findings suggest that the degree of suppression of the spermatogenic process is an important determinant of circulating inhibin. With steroid treatment, many of the steps of spermatogenesis are only partially suppressed—with azoospermia initially resulting from a failure of spermiogenesis [174]. Spermatagonial numbers, for example, fell only by 50%. Chemotherapy and X irradiation induced marked and immediate lesions at the spermatogonial level, although direct effects on the Sertoli cell cannot be excluded. These results suggest that germ cells, particularly at the earliest stages, contribute to the regulation of inhibin, by mechanisms that remain to be elucidated. Studies attempting to identify the germ cell–Sertoli cell mechanisms responsible have resulted in conflicting results [175, 176]. In a recent study of monkeys [177] in which gonadotropins were suppressed with a gonadotropinreleasing hormone (GnRH) antagonist, serum inhibin B levels were suppressed to a limited extent (15–30%), similar to that observed with steroidal treatment. However, the addition of testosterone implants or LH treatment resulted in a further ~50% fall in inhibin B levels. These findings suggest that LH/testosterone have an inhibitory influence on inhibin B subunit levels, adding a further complexity to the understanding of its regulation. At this stage it is unclear if serum inhibin B values are clinically useful, although some claims have been made that the combination of serum FSH and inhibin is superior to serum FSH alone as a marker in monitoring spermatogenesis [11, 12].

X. IMPLICATIONS OF INTEGRATED TGFβ SUPERFAMILY FUNCTION IN THE TESTIS: WHAT WE DO AND DON’T KNOW The importance of BMPs in germline specification, primordial germ cell generation and survival, and spermatogonial differentiation have been highlighted by many investigations, although the mechanisms by which they achieve these outcomes are in large part unknown. There is much to learn about how TGFβ superfamily signals affect fetal testis development, including how specific aspects of cord formation, Sertoli cell proliferation, and somatic cell maturation are affected. In the juvenile testis, understanding how regulated TGFβ superfamily signaling contributes to establishment of final Sertoli cell numbers may drive new methods for in vitro and in vivo expansion of the Sertoli cell population that will facilitate augmentation of germ cell development. The data presented in this chapter collectively support the concept that regulated actions by TGFβ superfamily members and their antagonists drive key events in Sertoli cell development and function and in germ cell differentiation. Some of these factors interact functionally with FSH [24, 56, 57, 130, 136], so this family of ligands is crucial for coordinating germ cell maturation and Sertoli cell function. Interaction with other known regulators of Sertoli cell maturation (reviewed in [170]) is an area requiring further investigation. Great advances have been made in the area of stem cell biology, with new models developed to use Sertoli cell overexpression of GDNF to drive expansion of the stem cell population [139, 140]. Integration of TGFβ superfamily signaling is emerging as a vital regulator of spermatogonial stem cell development, with GDNF sustaining the stem cell phenotype while activin and BMPs promote spermatogonial differentiation. Our increasing ability to control events at the onset of spermatogenesis that affect maintenance and differentiation of the germline stem cell population may provide new therapeutic options in the treatment of male infertility. Development of contraceptives that sustain the stem cell population but prevent completion of spermatogenesis will also be supported by this knowledge. It is in the adult testis that the local effects of TGFβ superfamily members become even more challenging to investigate, due to the extreme complexity of the testicular architecture layered on top of the extravagant number of interacting signaling networks. Advances have been made through production of knockout and

Chapter 14 The Transforming Growth Factor β Superfamily in Sertoli Cell Biology

transgenic animals in which the expression pattern of one or more genes affects Sertoli cell function or other aspects of testicular physiology (Table 14.3). The influence of TGFβs on Sertoli cell junctions has yielded insights into the nature of their formation [114]. It may prove useful to integrate this knowledge with clinical findings that serum inhibin levels change rapidly in response to certain types of damage [173], so that the integrity of Sertoli cell function can be more accurately assessed. Understanding the links between cytokines, TGFβ superfamily members and immunosuppression in the testis may illuminate mechanisms that can be exploited to achieve therapeutic outcomes in this and other organs. To fully understand the complex integration of TGFβ superfamily signaling inputs and their impact on testis development and function, it is vital to identify the sites where these components exist. Some of these have been intensively investigated (Tables 14.1 and 14.2), and others remain to be determined. It is interesting to note that the TGFβs, activins, and GDNF are much more highly expressed in the fetus and/or neonate than in the maturing and adult testis [71, 101, 126, 192]. There is also evidence of transient elevation of BMP4 and the ActRII at the onset of spermatogenesis [29, 92]. These and other lines of evidence indicate that TGFβ signaling governs events in early testis development, including the onset of spermatogenesis and Sertoli cell differentiation. Although the list of superfamily members is relatively stable, the recent identification of new receptor subunit functions and new antagonists continues to offer insights and provide challenges for our efforts to study testicular physiology. There are many components of the signaling pathway for which the sites of synthesis have not been systematically studied. Similarly, there are very few studies of testis biology that compare the impact of different ligand actions on the same functional endpoint, although it is quite apparent from the survey presented here that there are many potential points of overlap. Competition and enhancement may be the result of integrated signaling by TGFβ superfamily ligands, and the next decade should enhance our understanding of this. Some headway has been made by breeding animals together that bear single gene mutations. Insights into the functional importance of inhibin α and BMPs have been gained in this way [61, 96, 179]. Conditional knockouts will overcome the deficit of information about postnatal function of activin and TGFβs, because mice lacking many of these genes do not survive past birth (reviewed in [2]). As a final challenge, developing an understanding of how the TGFβ superfamily interacts with other signaling pathways [181] will be required

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to fully understand the mechanisms that underpin testis development and function.

Acknowledgments Supported by fellowships (K.L.: 143792; DR: 241000) and program grants (K.L.: 1147386; D.R.: 983120) from the National Health and Medical Research Council of Australia.

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V SERTOLI CELL TRANSCRIPTIONAL REGULATION

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C H A P T E R

15 Transcription Factors in Sertoli Cells JAIDEEP CHAUDHARY AND MICHAEL K. SKINNER Center for Reproductive Biology, School for Molecular Biosciences, Washington State University, Pullman, Washington

role in sex determination [6–8] to the adult stage of maintaining spermatogenesis. Each of these is associated with, and regulated by, a series of coordinated gene expression events at various periods of development [9–12]. For example, the molecular events leading to the initiation of Sertoli cell differentiation at the time of sex determination involves a coordinated and sequential expression of a number of DNA binding proteins such as WT-1, SF-1, Sry, Sox9, GATA-6, Dmrt1, and DAX-1 [13–17]. The expression of these proteins is achieved through an intricate and complex transcriptional regulatory network. The transcriptional regulation of Sertoli cell gene expression in large part determines the unique differentiated state and functional status of the cell throughout development. Therefore, the unique set of transcription factors present and how they interact to coordinate gene expression determines the state of Sertoli cell differentiation required to maintain testis function and spermatogenesis. This chapter reviews what is currently known regarding the transcription factors present in Sertoli cells and how they may coordinate specific cellular functions.

I. INTRODUCTION II. TRANSCRIPTION FACTORS III. TRANSCRIPTIONAL CONTROL OF SERTOLI CELL FUNCTION IV. TRANSCRIPTION FACTORS EXPRESSED IN SERTOLI CELLS V. REGULATION OF SERTOLI CELL GENES BY COMBINATORIAL INTERACTIONS OF TRANSCRIPTION FACTORS VI. CONCLUSION AND FUTURE DIRECTIONS References

I. INTRODUCTION Sertoli cells throughout development differentiate and undergo a number of functional and structural modifications [1]. These modifications start with Sertoli cell fate determination at the time of male sex determination in embryonic day 11–13 mice [2]. Following this initial event, Sertoli cells undergo a rapid phase of proliferation and differentiation [1]. The functional maturation at puberty is achieved by exit from the cell cycle that corresponds with the formation of the blood–testis barrier [3]. In the adult, the primary functions of Sertoli cells are to support spermatogenesis by (1) providing the microenvironment for the growth and development of the germ cells, (2) transmitting hormonal/environmental cues to the developing germ cells through paracrine signaling mechanisms, and (3) providing the structural support for the developing germ cells [4, 5]. Therefore, a number of distinct differentiated and functional states of the Sertoli cell exist from the earliest period in the embryo with their SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

II. TRANSCRIPTION FACTORS Transcription factors are DNA binding proteins (trans-acting) that bind to DNA regulatory elements located cis- to the target genes. Transcription factors are generally considered final targets of signal-transduction pathways [18–20]. The level of expression of transcription factors and their activities determines whether their target genes are transcribed and to what extent.

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Jaideep Chaudhary and Michael K. Skinner

These regulators of gene expression in turn are tightly regulated by a multitude of signaling pathways influenced by cell–cell interaction, growth factors, hormones, extracellular matrix, and the stage of development. This is exemplified by the observations that the gene expression profile of Sertoli cells associated with germ cells at different stages of spermatogenesis are different [9, 21–29]. In response to follicle stimulating hormone (FSH), the prepubertal cells express genes involved in proliferation and, following puberty, the cells express genes involved in maintaining differentiation and spermatogenesis [30, 31]. Therefore, alterations in the transcriptional regulation of the Sertoli cell determine changes in cellular differentiation and function. In general, the transcriptional control unit, or the promoter, present immediately upstream of the transcriptional start site of the gene is a cluster of binding sites for the RNA polymerase complex and transcription factors [32, 33] (Fig. 15.1). Most of the Sertoli cell gene promoters have a conserved TATA box, which binds the basal transcription factor initiation complex (TFIID, IIB, and IIF). These TATA box-containing promoters are highly regulated in that these promoters are switched on or off in response to a specific stimulus. Alternatively, some promoters of the housekeeping genes such as the androgen binding protein (ABP) [34]

and FSH receptor [35, 36] are devoid of TATA boxes but instead contain a G/C-rich sequence and an initiator region (Inr) [37, 38]. These TATA-less promoters can allow initiation of transcription at multiple sites over a broad region, often generating transcripts with multiple 5’ ends [39]. The organization of the typical promoter is shown in Figure 15.1. In addition to the promoter, an enhancer element can also stimulate transcription in an orientation-independent manner [40, 41] (Fig. 15.1). These enhancer elements can be located distal to the promoter or in the introns and 3’ UTR regions. The enhancer elements can also bind multiple gene-specific transcription factors. As an added level of control, the bound transcription factors can interact to form cell specific transcription factor complexes. These interactions can be mediated by regulatory proteins (coactivators/repressors), which may not directly bind to the DNA, but facilitate interactions between DNA bound transcription factors [42, 43] (Fig. 15.1). Equally important are the transcription factors and regulatory proteins that repress the transcription and are often identified as silencer regions on the promoter or enhancers [44]. These reversible modifications are important steps in the control of cellular gene expression.

III. TRANSCRIPTIONAL CONTROL OF SERTOLI CELL FUNCTION The functional and developmental changes associated with Sertoli cell differentiation and functions are a result of stage-specific activation/repression of specific transcription factors. Some of these transcription factors such as Sry [45] and DAX-1 [46, 47] are expressed only during a particular stage of development. Factors, such as C/EBPβ [48] and androgen receptor, are dependent on the presence/absence of specific stimuli such as FSH and specific stages of spermatogenesis [49], respectively. Thus, to achieve the complexity of functions required by Sertoli cells, such as sex determination and spermatogenesis, it is apparent that these cells achieve a degree of transcriptional control that can be achieved by modulation of the activities of specific transcription factors by one or more of the following mechanisms:

FIGURE 15.1 The organization of a typical eukaryotic promoter. In general, the eukaryotic promoters contain a TATA box within the proximal −30 bp of the transcriptional initiation site, which binds the basal transcriptional machinery. The promoter upstream of the TATA box can be organized into proximal (about −500 bp), distal (−500 to −1000 bp), and enhancer (more than −1000 bp). These regions can bind sequence-specific transcription factors. The bound transcription factor can also physically interact either directly or through accessory proteins in order to form a cell-specific transcriptional complex.

1. Phosphorylation/dephosphorylation, for example, CREB phosphorylation in response to FSH [50] 2. Protein–protein interactions, for example, interactions between bHLH, CREB, and CBP in transferrin promoter [51] 3. Ligand-dependent activation of the nuclear hormone receptors, for example, activation of

Chapter 15 Transcription Factors in Sertoli Cells

androgen receptor and retinoic acid receptors by binding of their cognate ligands, androgen and retinoic acid, respectively 4. Expression during a particular stage of development, for example, spermatogenesis stage-dependent expression of androgen receptor 5. Availability of DNA binding sites (methylation/demethylation), for example, Sertoli cell–specific demethylation of the E-box response element in FSH receptor promoter [52] A large number of transcription factors have been identified in Sertoli cells, mostly through comparing genetic disorders in Drosophila, Caenorhabditis elegans, humans, and mice, leading to a male reproductive phenotype such a sex reversal (ambiguous gonads, vanishing testes, persistent Müllerian duct syndrome, loss or decrease in fertility and testis size). These elegant studies have resulted in the identification of key Sertoli cell–specific transcription factors and their postulated mechanism of action. These include Sry, DAX-1, Dmrt1, CREB, AR, and GATA among many others listed in Table 15.1. The complex interplay and

TABLE 15.1 Categorization of Transcription Factors in Sertoli Cells 1. Transcription factors with basic domains a. Basic leucine zipper (bZIP); jun, fos, CREB, ATF, C/EBP b. Basic helix–loop–helix (bHLH); E2A, E2–2, Id1, Id2, Id3, Id4, REB c. bZIP-bHLH: USF, USF2, c-myc 2. Transcription factors with zinc-finger domains a. Nuclear receptors: AR, ER, RAR, RXR, PPAR, TR, SF-1, DAX-1 b. Cys4 zinc fingers: GATA, FOG c. Cys2His2 zinc-finger domains: TFIIIA, Sp1, SP3, Sp4, WT1 3. Transcription factors with helix–turn–helix motifs a. Paired box: Pem b. LIM homeodomain: Lhx9 c. Winged helix/Forkhead: Trident 4. β scaffold factors with minor groove contacts a. Rel/Ankyrin: NFκB family b. HMG: Sox, Sry 5. Others a. STAT: STAT-1 and -2 b. CBP c. Dmrt1

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hierarchy of these factors at the Sertoli cell “systems level” remain to be investigated.

IV. TRANSCRIPTION FACTORS EXPRESSED IN SERTOLI CELLS Recent progress in high-throughput gene expression profiling technology has allowed us to develop a comprehensive understanding of the Sertoli cell transcription factor “Transcriptome.” The Sertoli cells cultured from postnatal 20-day-old rats express a large and diverse group of transcription factors as listed in Table 15.2. The expression of some of these transcription factors is based on previous studies such as the retinoic acid receptor α, upstream transcription factor USF-1, or GATA-4 and GATA-6. The high levels of expression of p53, orphan nuclear receptors, Drosophila hairy, and enhancer of split (HES) are novel observations and suggest a functional significance of these factors in Sertoli cells. In response to FSH, the expression of some of the transcription factors remains unchanged, suggesting that they may be involved in maintaining basal Sertoli cell functions or may undergo phosphorylation or other post-transcriptional modifications in response to specific stimuli in order to be active. As expected, the expression of the majority of transcription factors is increased in response to FSH (Table 15.2). Most notable among these are C/EBPβ, which may be involved in mediating the FSH-cAMP-PKA response. The highest increase in the expression levels is observed for the orphan nuclear receptors Nr4a2 and Nr4a3 (40-fold average) within 2 hr of FSH stimulation (Table 15.2B). A large number of transcription factors also decrease in response to FSH, which include C/EBPδ, retinoic acid receptors, GATA4, and thyroid hormone receptor α. The dynamics of activation of certain transcription factors in response to FSH over a period of time (24 hr) also correlates well with the previously published reports. These include cAMP-responsive element modulator (CREM), which is induced after 4 hr of FSH stimulation, a delayed response when compared to C/EBPβ. The data provided in Table 15.2 should serve as a comprehensive overview of the basal and FSH-induced Sertoli cell transcription factor profile. The physiological function and significance of each of these transcription factors should be considered in relation to mechanisms that can modify their activities as listed earlier. In the following sections, the Sertoli cell transcription factors are discussed that have a previously identified functional significance in terms of stage-specific expression (outlined in Table 15.3), expression in response to hormones and growth factors, and/or that have a

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Jaideep Chaudhary and Michael K. Skinner TABLE 15.2A Sertoli Cell Transcription Factor: FSH Responsiveness

Transcription Factor Class/Sub class/name/Access. no.

0 hr Signal

2 hr xChange

4 hr xChange

1. Transcription Factors with Basic Domains 1.a Basic Leucine Zipper Jun

AI175959

c-jun

429

0.5

Atf3

M63282

Activating transcription factor 3 (ATF3)

234

1.4

1.0 0.5

Junb

AA891041

jun B proto-oncogene

171

5.7

2.7

Crem

S66024

cAMP responsive element modulator (CREM)

113

5.8

11.2

Creb1

X14788

cAMP response element binding protein 1 (CREB)

22

1.6

1.5

Cebpd

M65149

CCAAT/enhancerbinding, protein (C/EBP) delta

106

0.4

0.4

Cebpb

X60769

CCAAT/enhancer binding protein (C/EBP), beta

67

7.5

2.4

Dbp

J03179

D site albumin promoter binding protein

247

0.3

0.0 1.2

1.b. Basic Helix–Loop–Helix Hes1

D13417

Hairy and enhancer of split 1 (Drosophila)

212

2.1

rE12

S77532

Helix–loop–helix transcription factor E12

64

0.9

1.0

Hif1a

Y09507

Hypoxia inducible factor 1, alpha subunit

43

1.7

2.1

Usf1

AA891717

Upstream transcription factor 1

702

0.7

0.5

Usf2

X90823

Transcription factor USF2

46

1.0

1.4

U61184

Aryl hydrocarbon receptor nuclear translocator

65

0.9

1.0

1.c. bHLH-ZIP

1.d. bHLH-PAS Arnt

Note: The data are the mean of two different experiments and are within a 15% coefficient of variation (CV). The Xchange represents increases (bold) or decreases (underline) of the transcription in cultured rat (20-day-old) Sertoli cells treated with FSH for various time intervals. The data were generated by using the rat Affymetrix chip 434. The analysis was obtained from a data set previously described by Dr. Derek McLean and Dr. Michael Griswold [324].

TABLE 15.2B Sertoli Cell Transcription Factor FSH Responsiveness Transcription Factor Class/Sub class/name/Access. no.

0 hr Signal

2 hr xChange

4 hr xChange

8 hr xChange

24 hr xChange

2. Transcription Factors with Zinc-Finger Domains 2.a. Nuclear Receptors Rara

U15211

Retinoic acid receptor, α

749

0.5

0.5

0.6

0.6

Nr1h2

U14533

Ubiquitous Receptor/Liver X receptor β

662

0.6

0.7

0.8

0.5

Thra

M31174

Thyroid hormone receptor α

608

0.7

0.7

0.6

0.9

Nfyc

AA875121

Nuclear transcription factor-Y γ

302

1.0

1.1

1.2

1.1

Nr0b1

X99470

DAX-1

265

0.9

0.7

0.5

1.1

Nr5a1

D42156

SF-1

133

0.3

0.2

0.2

0.2

Nr0b2

D86580

SHP (Small Heterodimer Partner)

109

0.2

0.2

0.2

0.4

Rxra

L06482

Retinoid X receptor alpha

83

0.3

0.6

0.3

0.2

Ppard

U40064

Peroxisome proliferator activated receptor δ

68

0.7

0.8

0.6

0.7

Nr4a1

U17254

IEG transcription factor NGFI-B

65

24.0

13.9

1.8

2.1

Nfya

M34238

Nuclear transcription factor-Y α

56

0.6

1.0

1.2

0.7

Nfix

AB012234

Nuclear factor I/X

52

0.5

0.7

0.6

1.1

51

3.7

3.1

1.9

2.0

8

39.5

30.5

8.0

2.5

Nr1d2

U20796

Rev-Erb Alpha

Nr4a3

AI176710

Nuclear Orphan Receptor 1

Continued

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Chapter 15 Transcription Factors in Sertoli Cells TABLE 15.2B Sertoli Cell Transcription Factor FSH Responsiveness—cont’d Transcription Factor Class/Sub class/name/Access. no.

0 hr Signal

2 hr xChange

4 hr xChange

8 hr xChange

24 hr xChange

Nr4a2

U01146

NOT, RNR-1, H2F-3, NURR1

6

40.6

26.8

4.9

3.6

Nr1d1

M25804

NR subfamily 1, group D, member 1

205

0.4

0.4

0.3

0.7

Nr2f6

AF003926

NR subfamily 2, group F, member 6

181

1.3

2.5

1.7

1.0

2.b. Cys4 Zinc Fingers Gata4

L22761

GATA-binding protein 4

306

0.5

0.9

0.7

0.4

Gata6

L22760

GATA-binding protein 6

161

1.7

1.9

1.2

0.9 1.3

2.c. Cys2His2 Zinc-Finger Domains Tieg

AI172476

TGFB inducible early growth response

112

1.8

1.6

1.6

Wt1

X69716

Wilms tumor 1

70

1.6

1.8

1.4

1.4

Egr1

M18416

Early growth response 1

86

1.2

0.6

0.5

1.1

Egr4

AI145177

Early growth response 4

74

1.8

2.0

1.3

1.2

Klf9

D12769

Kruppel-like factor 9

47

4.1

3.8

2.1

1.1

Note: The data are the mean of two different experiments and are within a 15% coefficient of variation (CV). The Xchange represents increases (bold) or decreases (underline) of the transcription in cultured rat (20-day-old) Sertoli cells treated with FSH for various time intervals. The data were generated by using the rat Affymetrix chip 434. The analysis was obtained from a data set previously described by Dr. Derek McLean and Dr. Michael Griswold [324].

TABLE 15.2C Sertoli Cell Transcription Factor FSH Responsiveness Transcription Factor Class/Sub class/name/Access. no.

0 hr Signal

2 hr xChange

4 hr xChange

8 hr 24 hr xChange xChange

3. Transcription Factors with Helix–turn–Helix motifs 3.a. Homeo Domain Tcf1; HNF1

X54423

Transcription factor 1

56

0.7

0.6

0.4

0.6

AF053100

Paired box gene 4

51

0.5

0.5

0.2

0.8

3.b. Paired Box Pax4

3.c. Tryptophan Clusters Ets1

AI175900

v-ets E26 oncogene homologue 1 (avian)

Irf1

M34253

Interferon regulatory factor 1

81

0.6

0.9

0.9

0.8

222

1.6

0.8

1.0

1.6

2.9

1.0

4. Beta Scaffold Factors with Minor Groove Contacts 4.a. Rel/ Ankyrin L26267

Nuclear factor κ B p105 subunit

138

1.5

4.6

Stat1

AA892553

Signal transducer and activator of transcription 1

104

0.5

0.4

0.1

0.6

Stat3

X91810

Signal transducer and activator of transcription 3

82

0.6

0.5

0.7

0.4

Stat5b

X91988

Signal transducer and activator of transcription 5b

41

1.7

1.3

1.2

1.5

X13058

Tumor protein p53

306

0.6

0.7

0.7

0.6

Nfkb1 4.b. STAT

4.c. Class p53 Tp53

Note: The data are the mean of two different experiments and are within a 15% coefficient of variation (CV). The Xchange represents increases (bold) or decreases (underline) of the transcription in cultured rat (20-day-old) Sertoli cells treated with FSH for various time intervals. The data were generated by using the rat Affymetrix chip 434. The analysis was obtained from a data set previously described by Dr. Derek McLean and Dr. Michael Griswold [324].

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Jaideep Chaudhary and Michael K. Skinner TABLE 15.3 Expression of Transcription Factors in Relation to Developmental Stage of Sertoli Cellsa

Transcription factor

Undifferentiated gonad

Sex determination

FSH receptor Androgen receptor

?

?

Aromatase

Embryonic differentiation

Prenatal/prepubertal

Puberty/adult

++

++

++

++

++

++

++

++

–

Transferrin

–

–

+

+

++

Sry

–

+++

–

–

–

SF–1

++

+++

++

+

+

FOG2

+++

+++

+/–

+

++

Sox9

++

+++

+

+

++

FOG1

–

–

+

++

++

GATA–2

++

–

–

–

–

GATA–1

–

–

–

+

++

GATA–4

+

++

+

+

+

NFκB CREB

+(?)

+(?)

++

CREM

+(?)

+ (?)

++

+

+

+ ++

++

WT–1

++

Jun Fos

?

?

?

++

Lhx9

?

++

++

?

?

DAX–1

?

+

++

++

++

a

The expression of FSH receptor, androgen receptor, aromatase, and transferrin are also indicated as markers of functional differentiation of Sertoli cell. Legend: +, present; –, absent; ?, expression not known.

potential mechanism of action in regulating Sertoli cell genes. This is discussed in regards to the transcription factor categorization outlined in Table 15.1.

A. Transcription Factors with Basic Domains 1. Basic Leucine Zipper The characteristic feature of basic leucine zipper (bZIP) transcription factors is the conserved 60- to 80-amino-acid bipartite alpha helix. The alpha-helix at the C terminal contains leucine every seven amino acids. The sequence-specific DNA binding is achieved by the highly charged basic region toward the N-terminal of bZIP transcription factors. The bZIP transcription factors bind to a double-stranded consensus DNA sequence as homodimers or heterodimers (Fig. 15.2). Unlike other transcription factors, the presence of bZIP proteins has been shown only in eukaryotes [53]. a. C/EBP CCAAT/enhancer binding protein (C/EBP) transcription factors are involved in diverse cellular

FIGURE 15.2 Schematic of basic leucine zipper transcription factors. The leucine zipper mediates dimerization and the charged basic domain is involved in DNA binding. Significant variation between the length of the N and C terminals is observed within the family members and may affect the dimerization.

Chapter 15 Transcription Factors in Sertoli Cells

processes such as proliferation, differentiation, metabolism, inflammation, and numerous other responses and are highly expressed in hepatocytes, adipocytes, and hematopoietic cells. C/EBP dimers bind to the palindromic DNA consensus sequence 5’: –A/GTTGCG/ CTCAA/T–3’ [54, 55]. Structure–function analyses of these proteins have identified several trans-activation domains, some of which can physically interact with general transcription factors present in the initiation complex. At least six members of the family have been isolated and characterized (C/EBPα–C/EBPζ). The functional diversity of the C/EBP family is further expanded by the generation of polypeptides through the use of alternative translation initiation sites and through interactions within the family and with other transcription factors [56]. Heterodimeric interaction partners of C/EBP include Fos, Jun [57], ATF4 [58, 59], ATF2 [60], and CREB [61]. Largely considered constitutively acting factors, recently C/EBPs have also been shown to mediate cAMP responses. A number of cAMP-responsive gene promoters also contain binding sites for C/EBP. Generally, these binding sites are located within the region of the promoter that is responsible for mediating the acute responsiveness to cAMP. Collective evidence now suggests that C/EBPs may have both constitutive and cAMP-inducible activities, and are now considered a cAMP-responsive nuclear regulator. The role of C/EBP in mediating cAMP responses can also be due to the ability of C/EBP dimers to bind related CRE sites [62]. The functions of the C/EBP family of transcription factors in energy metabolism and as a downstream effecter of cAMP responses are also functions in Sertoli cells. C/EBPα, β, δ, and ζ messenger RNA (mRNA) and proteins are present in Sertoli cell primary cultures but only the β and δ isoforms are induced in the presence of FSH or cAMP [48]. The β isoform appears to be predominant since this is the primary isoform observed in oligonucleotide gel shift experiments. The kinetics of C/EBP expression suggests that the early induction of C/EBP isoforms by cAMP may play a role in FSH-dependent regulation of late response genes in Sertoli cells. These observations are also supported by data presented in Table 15.2. C/EBPβ may mediate the cAMP responses elicited by FSH because its expression is increased within 2 hr by more than 25-fold in response to FSH. In contrast, the expression of C/EBPδ expression is decreased more than 2-fold in response to FSH. The significance of C/EBPδ downregulation following FSH treatment is intriguing and suggests that isoform-specific homoand/or heterodimerization may be involved in limiting FSH action.

257

b. CREB CREB (cAMP response element binding protein) is a large superfamily and consists of the closely related factors CREM [63], ICER (an alternate spliced form of CREM) [64], and ATF-1 (activating transcription factor 1) (Fig. 15.3). A high degree of homology within the bZIP domain of CREB, CREM, and ATF-1 allows them to form homo- and heterodimers and to bind to the same palindromic cis-regulatory element TGACGTCA [65]. While many CREB binding sites are comprised of variations of this consensus motif, almost all harbor the core sequence CGTCA. The activation of the cAMP pathway by the binding of FSH to its cognate receptor and subsequent activation of CREB and associated downstream events influencing spermatogenesis have been extensively studied and reviewed elsewhere [66, 67]. The immediate effect of cAMP or FSH on Sertoli cells is the phosphorylation of CREB, which usually occurs within 1 to 5 min [50]. The almost instant phosphorylation and activation of CREB at serine 133 ensures the expression of downstream regulatory genes such as c-fos [66]. The presence of at least two CRE sites in the proximal 300 bp of the CREB promoter is required for cAMP-inducible expression of the gene and supports the model of FSHand cAMP-mediated CREB autoregulation of its own promoter [50]. This autoregulatory mechanism may explain the dramatic stage-specific oscillations in Sertoli cells of CREB messenger RNA levels during the 12-day cycles of spermatogenesis in rat seminiferous tubules [21]. c. CREM CREM and its spliced variants α, β, and γ bind to a consensus CRE with high affinity and specificity similar to CREB. Unlike CREB, CREM lacks the transcriptional

FIGURE 15.3 Typical organization of the CREB family of transcription factors. Q1, Q2: represent the transactivation domain and are present in all CREB isoforms and CREMt, but absent in ICER and CREMα; KID: kinase inducible domain is present in all CREB and CREM isoforms but not in ICER; Basic: mediates binding to CRE sequence; leucine zipper: mediates dimerization, NLS: nuclear localization signal.

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activation domain. Therefore, CREM competes with the binding of CREB to the CRE (Fig. 15.3). In addition to CREB, the Sertoli cells also express the repressor ICER, an isoform of CREM [68] (Fig. 15.3). ICER is expressed in response to FSH and its induction accompanies downregulation of CREB and FSH receptor transcript by binding to a CRE-like sequence in the regulatory region of the gene leading to long-term desensitization [26]. The gene expression profile of the transcriptional repressor CREM reported in Table 15.2 validates the previous results, in that it is induced at least 4 hr after stimulation of Sertoli cells by FSH [69]. This delayed expression of ICER may be involved in regulating the expression of early event genes such as C/EBPβ, which are induced within 2 hr of FSH stimulation. d. The AP-1 Complex The AP-1 transcription factor complex consists of a dimer between members of the fos, jun, and ATF families of proteins [70, 71]. In addition to fos, jun, and ATF, CREB proteins can also form part of the AP-1 complex. Whereas fos proteins can only heterodimerize, the jun proteins can both homo- and heterodimerize with fos [72], ATF2 [73], and ATF3 members to form transcriptionally active complexes. The jun-fos dimers preferentially bind to the TRE (TGACTCA, TPA response element) [74]. The preference for the binding changes to the consensus CRE response element, TGACGTCA, when one of the members of the dimer is ATF or CREB [75, 76]. The jun homodimers bind DNA at low affinity and are generally considered transcriptionally inactive [77]. Preliminary results from our laboratory have also shown that members of the AP-1 family, specifically, ATF-3 can interact with scleraxis, a member of the basic helix–loop–helix (bHLH) transcriptional family (unpublished observations). cfos Given its proto-oncogene status, cfos is the most actively studied transcription factor in any cell system. In Sertoli cells cfos has been implicated as an early event gene that is rapidly and transiently induced in response to FSH [78–80]. Upon stimulation with FSH and cAMP, the expression of cfos is observed within 30 min. The kinetics of induction of cfos in relation to other Sertoli cell genes such as transferrin, plasminogen activator [81], ABP, and inhibin-α subunit [81] support the notion that cfos is an immediate early gene. This is also supported by the observation that treatment of Sertoli cells with cfos blocks FSH-mediated activation of transferrin [82] and their attachment and spreading on contact with extracellular component [83]. The later observation can be extended to suggest the role of cfos in the formation and maintenance of the blood–testis barrier and basal lamina. In addition to FSH, other

growth factors, possibly from germ cells, such as basic fibroblast growth factor (FGF) [84], leukemia inhibitory factor (LIF) [85], growth hormone releasing hormone (GHRH), nerve growth factor (NGF), interleukin 6 (IL-6), and interferon γ [86] also have a rapid stimulatory effect on cfos expression. In cultured immature porcine Sertoli cells, FGF has been shown to stimulate cell proliferation and plasminogen activator activity by early induction of the cfos gene independent of cAMP and the calcium/ phospholipid pathways [84, 87]. Collectively, these studies suggest that cfos expression in Sertoli cells is regulated by diverse signaling pathways. The kinetics of induction of cfos by hormones and growth factors ranges from 15 min to 2 hr with return to basal levels between 6 and 18 hr [81, 85, 86]. Some of these differences observed in the time of induction of cfos following treatments may be due to culture conditions, concentrations of hormone used, and time of treatment after Sertoli cell isolation. These results suggest that cfos activation may be an important early event and lack of its function/expression may lead to gross Sertoli cell abnormalities. Mice lacking cfos are viable and fertile but lack osteoclasts [88]. But the homozygous chimeric mice generated by embryonic stem cells targeted at the c-fos locus show reduced placental and fetal weights and significant loss of viability at birth. Among other abnormalities these mice also show delayed or absent gametogenesis, lymphopenia, and altered behavior [89]. The effect on gametogenesis supports a potential critical role for cfos in Sertoli cells. Jun (c-jun, junB, and junD) Consistent with its expression elsewhere, junD is also highly expressed in the embryonic testis and in purified postnatal and prepubertal Sertoli cells (unpublished observation). High levels of expression of JunD suggest the importance of the AP-1 transcription factor complex in testicular development and Sertoli cell differentiation. Consistent with this observation, the male junD knockouts have reproductive phenotypes that include reduced fertility with age, increased inhibin, and decreased FSH serum levels [90]. In spite of being highly expressed, the regulation of junD has not been extensively studied in Sertoli cells. The expression of other members of the jun family such as junB is increased in response to FGF and FSH [81]. JunB can alternate or sometimes antagonize the activity of c-jun. Consistent with these observations, FSH negatively regulates the expression of c-jun [81]. Interestingly, the c-jun mRNA levels are increased, following retinoid (vitamin A) stimulation of the Sertoli cells [91] and in response to tumor necrosis factor alpha (TNFα) it is phosphorylated [92]. The differential dynamics of c-jun and junB expression in

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response to FSH and vitamin A is interesting and will need further evaluation regarding the role of the jun proto-oncogene family in Sertoli cell function. [92]. 2. Basic Helix–Loop–Helix Transcription Factors The bHLH transcription factors are critical cell-type determinants and play important roles in cell growth, differentiation, and sex determination [93]. A bHLH domain that is conserved from yeast to mammals characterizes members of this family [94]. The bHLH domain consists of two amphipathic helixes separated by a loop that mediates homo- and heterodimerization adjacent to a DNA-binding region rich in basic amino acids [95, 96] (Fig. 15.4). The bHLH dimers bind to an E-box (CANNTG) DNA consensus sequence present in a wide variety of promoters [96, 97]. The E-box response element has been shown to influence the promoters of a number of Sertoli cell genes [98–101]. The bHLH proteins have been classified into two distinct classes. The ubiquitously expressed class A bHLH proteins consist of E2–2 [102], HEB [103], and E12 and E47 (the differentially spliced products of the E2A gene) [96]. These dimerize with tissue-restricted and developmentally regulated class B proteins such as MyoD and neuroD [104, 105]. The members of the Id (Inhibitor of differentiation/DNA binding) family modulate the transcriptional activity of class A and B bHLH heterodimers. The four known Id proteins, Id1 [106], Id2 [106], Id3 [107], and Id4 [108], share a homologous HLH domain, but lack the basic DNA binding region. Thus, the Id proteins act to sequester bHLH proteins (preferentially binding to class A) by forming inactive dimers to prevent binding of bHLH proteins to the E-box sites [109] (Fig. 15.4). Therefore, Id proteins function as dominant negative bHLH transcription factors [110]. Alternatively, Id proteins can also interact with several other non-bHLH proteins and influence cellular function. For example, Id proteins can interact with proteins such as retinoblastoma (Rb) tumor suppressor protein to promote cell division [111] and ETSTCF proteins to modulate the intracellular response to growth factors [112]. The potential role of bHLH proteins in regulating Sertoli cell function was hypothesized based on the observations that in Drosophila, daughterless and ASC (acheate-scute-complex) have a role in sex determination [113]. Subsequent promoter analysis of Sertoli cell genes also showed the presence of conserved E-box response elements. Systematic studies involving mutations in the E-box used reverse transcriptase–polymerase chain reaction (RT-PCR) to determine known and ubiquitously expressed bHLH proteins; these studies strongly suggested their significance in regulating Sertoli cell function [98, 99]. Sertoli cell expressed genes

FIGURE 15.4 Schematic of basic helix–loop–helix family of transcription factors. In general, the bHLH transcription factor heterodimerizes. The dimerization is mediated by the HLH domain. The changed basic domains of the dimer bind the conserved E-box response element (CANNTG). The inhibitor of differentiation (Id) family of HLH transcription factors lacks the basic domain. The heterodimers between bHLH and Id proteins therefore fail to bind DNA and initiate transcription.

indicate that transferrin, ABP, FSHR [114, 115], SF-1 [116], c-fos [99], and the RIIβ subunit of cAMPdependent protein kinase promoter [117] are regulated by an E-box response element. The ubiquitously expressed proteins E47 and REBα are expressed in Sertoli cells [118]. Surprisingly, the negatively acting bHLH proteins Id1, Id2, Id3, and Id4 are also expressed in postpubertal terminally differentiated Sertoli cells [28, 51]. The suppression of proliferation of prepubertal proliferating Sertoli cells treated with thyroid hormone, retinoic acid, and testosterone is accompanied by an increased expression of Id2 and Id3 [119]. Longterm exposure (48 hr) of postpubertal Sertoli cells to FSH induces Id4, but downregulates Id1 expression. In contrast, serum induces the expression of Id1, Id2, and Id3 but suppresses Id4 [51]. Short-term treatment of postpubertal Sertoli cells (4 hr) with FSH significantly upregulates Id3 (unpublished observations). In a similar experiment, a biphasic expression of Id2 is observed. The first peak of expression is observed at around 4 hr and a second peak appearing at about 8 hr. Such a biphasic expression pattern of Id2 is also observed in human diploid fibroblasts after serum

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stimulation, which corresponds to the G1 phase and G1–S transition, supporting their role in proliferation [120]. The postpubertal Sertoli cells fail to proliferate after FSH induction, but together these results suggest that these cells may be competent to reenter the cell cycle. The mechanisms by which Id proteins promote the cell cycle are diverse but may involve suppression of p21, p27, cyclin A, or E/CdK2 and interactions with pRb [121]. Given the role of Id proteins in cell proliferation, the functional significance of Id protein expression in terminally differentiated and postmitotic Sertoli cells is unclear. Contrary to the hypothesis that Id proteins have redundant functions, the results in these studies suggest that Id1, Id2, Id3, and Id4 are differentially regulated and may have distinct functions [51]. Id1 may act to maintain Sertoli cell growth potential, whereas Id2, Id3, and Id4 may be involved in the differentiation and hormone regulation of Sertoli cells. Recent observations from our laboratory have suggested that the additional bHLH proteins such as scleraxis, ITF-4, and HEY (hairy and enhancer of split, Drosophila homologue) are also expressed by Sertoli cells (unpublished observation). The significance of these transcription factors in Sertoli cells is currently being investigated. 3. bZIP-bHLH: USF, USF2, and cMyc The bZIP bHLH family can be considered a family of composite transcription factors with two domains, the bZIP and the bHLH. The bHLH domain is considered the primary DNA binding domain, whereas the bZIP domain may be involved in protein–protein interactions. The presence of the bZIP domain also raises the possibility that these transcription factors may interact with other bZIP proteins such as cfos, jun, and CREB to form cell-specific transcription factor complexes. a. USF The two isoforms of USF, USF1 [122] and USF2 [123], are ubiquitously expressed in many cell types [124]. The highly conserved C-terminal bZIP-bHLH domain allows identical dimerization and DNA-binding specificities. Both of the isoforms bind to E-boxes as heterodimers [125]. USF1 and USF2 have been shown to regulate the expression of genes involved in the metabolism of glucose response. Consequently, the USF1 and USF2 knockout mice have a diminished glucose response [126, 127]. USF1 is expressed at high levels in Sertoli cells and in response to FSH its expression levels decrease by more than twofold. USF2 is expressed at a significantly lower level than USF1, but may be important in negatively regulating certain components of the FSH signaling pathway,

such as RIIβ, the regulatory subunit of cAMP-dependent protein [128]. The role of USF1 and USF2 in regulating Sertoli cell-specific expression of the FSHR gene has been extensively studied [36, 100, 115] and reviewed [129]. Both USF1/USF2, possibly as a heterodimer pair, bind to the E-box in the proximal FSHR promoter [100, 114]. The downregulation of USF1, 4 hr after FSH stimulation, a profile similar to ICER, may also be involved in negatively regulating FSHR expression (Table 15.2). In cells expressing FSHR genes such as Sertoli cells, the USF1/USF2 binding Eboxes sites are unmethylated, whereas these sites are methylated in non-FSHR-expressing cells. The methylated sites may interfere with the binding of general transcription factors such as USF leading to repressive chromatin structure in the nonexpressing cells [52]. b. c-myc In general, c-myc promotes cell growth and proliferation, inhibits terminal differentiation and sensitizes cells to apoptosis [130]. Consistent with these functions, c-myc expression in response to FSH [131] and testosterone [132] is observed in proliferating prepubertal and early pubertal Sertoli cells, but not in nonproliferating, postpubertal Sertoli cells [131, 132]. The presence of the E-box (CANNTG), which can bind the c-myc-max heterodimer in the promoter of Sertoli cell genes, may suggest its importance in regulating their expression. C-myc expression is also induced by IL-6 and interferon gamma (IFNγ) in primary Sertoli cells established from immature testis, suggesting its role in Sertoli cell proliferation in response to these mitogens [133].

B. Transcription Factors with ZincFinger Domains The transcription factors within this family are characterized by a short stretch of ~30 amino acids containing at least two conserved cysteines and histidines. The conserved C2H2 residues coordinate a zinc ion, allowing the finger to assume a compact β-turn with cysteines and an α-helix with histidine. DNA recognition by zinc-finger transcription factors is typically determined by amino acid sequence within the 30 amino acids and number of zinc fingers (three or more) [134]. 1. Nuclear Receptors The nuclear receptors are composed of three domains: (1) the NH2-terminal transcriptional activation domain (A/B domain), (2) the DNA-binding domain (DBD), and (3) the C-terminal ligand-binding domain [135] (Fig. 15.5). The nuclear receptors containing zinc

Chapter 15 Transcription Factors in Sertoli Cells

FIGURE 15.5 Structure of a typical nuclear hormone receptor. The N-terminal region (A/B transactivation domain) contains constitutively active transactivation region(s) (AF-1) and several autonomous transactivation domains (ADs). The highly conserved C domain functions as a DNA-binding domain and may also serve as a dimerization domain with other nuclear receptors. The liganddependent transactivation function of nuclear receptors is achieved by the binding of respective ligands to the E domain or ligand-binding domain.

fingers bind DNA as a dimer with each monomer recognizing one of the two hexanucleotides as an inverse or a direct repeat. The nuclear receptor superfamily has more than 48 known members [136, 137]. The nuclear hormone receptors binds specific steroids, terpene-derived molecules, and peptides. The majority of superfamily members have no recognized ligand and are therefore termed orphan receptors [138]. In Sertoli cells the following major groups of this superfamily are known: Group 1: Steroid receptors (bind DNA as homodimers); androgen receptors (ARs), GGTACAnnnTGTTCT; and estrogen receptors (ERs): AGGTCAnnnTGACCT Group 2: Receptors that dimerize with RXR (cis retinoic acid receptor) such as thyroid hormone (TR), AGGTCATGACCT; all-trans retinoic acid (RAR), AGGTCAnnnnAGACCA; and peroxisome proliferator-activated receptors (PPAR) Group 3: Orphan nuclear receptors (bind DNA as a monomers and/or dimers): SF-1, DAX-1 a. Androgen Receptor The androgen receptor is one of the earliest [139, 140] and most studied transcription factors in Sertoli cells [141, 142]. Over the years a number of studies have shown that both FSH and androgens are involved in regulating its expression. In cultured immature Sertoli cells, short-term (5-hr) exposure to FSH downregulates androgen receptor expression [143]. Long-term exposure of cultured immature Sertoli cells to FSH

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increases both AR protein and mRNA. The effects of FSH on AR expression can be mimicked by cAMP suggesting that the effect of FSH on AR are mediated by the PKA pathway [143]. Activation of the PKC pathway leads to the downregulation of AR. In vivo, a complex interplay between androgens and FSH influencing transcriptional, translational, and post-translation regulation of androgen receptor expression has been proposed. The earliest time point when androgen receptor expression appears in Sertoli cells is postnatal day 5, and it persists through adulthood. By day 21, at the initiation of spermatogenesis, the AR expression in Sertoli cells is predominantly observed in stages II–VII of the spermatogenic cycle [49, 144]. This stage-specific expression of AR may be an important mechanism for regulating Sertoli cell responsiveness to testosterone and its subsequent control of spermatogenesis [145]. The stage-specific expression of AR in adult Sertoli cells suggests that germ cell regulatory factors may also influence its expression. TNFα, a secretory product of round spermatids, has been shown to stimulate the expression of AR in Sertoli cells [146]. The highly regulated AR gene expression at the promoter lever is due to three κB enhancer elements that interact with Sertoli cell p50 and RelA nuclear factor kappa B (NFκB) proteins. The overexpression of these NFκB subunits in Sertoli cells stimulates AR promoter activity. In vivo, TNFα may stimulate NFκB binding to the AR promoter, leading to the increased promoter activity and AR expression in primary cultures of Sertoli cells [146, 147]. The androgen receptor gene has a repetitive DNA sequence in exon 1 that encodes a polyglutamine stretch. Within the normal polymorphic range this (CAG)(n) length is inversely related to the transcriptional activity of the androgen receptor and is directly reflected in the efficiency of spermatogenesis, but not overall fertility. For example, men with short CAG repeats have the highest sperm output within the normal fertile population. AR gene mutations are also common and cause androgen insensitivity syndrome with altered sexual differentiation in XY individuals [148, 149]. In addition to binding to its cognate response element, the AR also directs the assembly and stabilization of the basal transcription apparatus. Androgendependent cofactors and CREB-binding protein (CBP) are influenced by AR target gene promoters to enhance transcription. Regulation of AR activity in response to diverse signaling pathways have also been reported. For example, interactions between AR and members of the STAT family, PIASx and PIAS1 (protein inhibitor of activated signal transducer), have been demonstrated. The expression of PIAS1 is observed in Sertoli cells after puberty. In addition to Sertoli cells, PIAS1 expression is also present in Leydig cells and germ cells

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suggesting its role in androgen-dependent initiation and maintenance of spermatogenesis [150]. A small nuclear RING finger protein, SNURF (or RNFN), also regulates androgen-receptor dependent transcription. The 3.0-kb SNURF transcript is predominantly expressed in Sertoli cells of both immature and mature testis [151]. The molecular actions of AR on Sertoli cells remain to be fully elucidated. b. Estrogen Receptors The male mice homozygous for a mutation in the estrogen receptor gene (ER knockout; ERKO) are infertile [152]. These earlier studies showing lack of fertility in ERKO knockout mice were confirmed to be that of the ERα because male mice lacking the estrogen receptor beta (ERβ) exhibit no compromised fertility [153]. Because ERα is essentially absent in the Sertoli cells [154, 155], it was postulated that the infertility was a result of a defect in germ cell function. Recently, sperm transplantation studies have shown that the altered sperm function that is characteristic of the ERα knockout male [152] is the result of the loss of ERα actions in the reproductive tract rather than in the germ cell. In the double ERKO males, the α-βERKO indicates an overall phenotype of that of αERKO [156]. ERα is expressed only in Leydig cells, whereas ERβ is widely expressed in various testicular cell types such as Leydig, some peritubular cells, Sertoli cells, and germ cells, except spermatids and meiotic spermatocytes [157]. In the human testis, a spliced variant of the ERβ is also observed. The wild-type ERβ1 and its spliced variant, the ERβ2, are both detected in human testis. The expression of ERβ1 is most intense in pachytene spermatocytes and round spermatids, but low levels of expression are also detected in Sertoli cells. Highest levels of expression of ERβ2 protein are detected in Sertoli cells and spermatogonia. In the fetal human testis (12–19 wk of gestation) ERβ1 and ERβ2 are also expressed in some, but not all Sertoli cells [158]. Given the expression profile of both the estrogen receptors and the reproductive phenotypes in the knockouts, the functions of these two estrogen receptors are not redundant. Specific Sertoli cell actions of ER are unclear at the present time. c. Retinoic Acid Receptors The retinoic acid receptors (RAR) can be largely divided into two broad families with each family having at least three subtypes (α, β, and γ) encoded by a different gene. RARα, RARβ, and RARγ bind all-trans and 9-cis retinoic acid. RXRα, RXRβ, and RXRγ preferentially bind 9-cis retinoic acids. The heterodimerization between RAR and RXR, homodimerization between RXR, and heterodimerization of RXR and RAR with

other transcription factors can generate a large repertoire of transcriptional complexes that bind specific retinoic acid response element(s) and/or other DNA response elements [159–161]. The various retinoic acid receptors are widely expressed in fetal, neonatal, and adult Sertoli cells, suggesting the significance of vitamin A–dependent signaling in Sertoli cell function [162–164]. RARβ and RARγ, and RXRγ expression is first observed in the Sertoli cells of fetal testis and continues to be expressed in the adult [165, 166]. In the juvenile testis, the α isoforms of RAR and RXR are also observed. In the adult, both RAR and RXR and all of their subtypes are expressed [167]. Retinoic acid increases Sertoli cell proliferation in testicular explants from 14.5-day-old rats, an effect mediated by RARβ [165]. Some redundancy may exist between the expression of RAR subtypes in the Sertoli cells. The expression of RARα and RXRβ seems to be required for proper Sertoli cell function because the mice homozygous for deletion of either of these receptors are sterile due to altered Sertoli cell function [168–170]. The Sertoli cells in the RXRβ homozygous mutants show progressive accumulation of unsaturated triglycerides and those with RARα mutants show frequent vacuolation. The overall phenotype of the two-retinoid receptor knockout males is similar to the Sertoli cells of vitamin A– deficient animals, which are infertile [171]. The pathways regulated by RAR subtypes in the Sertoli cells are complex and are regulated at multiple levels. The dimerization between various subtypes and other transcription factors such as PPAR allows the dimer to bind different response elements, allowing activation/repression of selective genes. The nuclear/ cytoplasmic localization and expression of individual receptor subtypes in response to hormones/growth factors can also determine the timing and magnitude of expression of target genes [163]. For example, using cultured Sertoli cells from 20-day-old rats, retinoic acid in a ligand-dependent manner induces the translocation of RARα to the nucleus, which can be blocked by FSH [172]. The translocation of RARα to the nucleus can also be achieved in a ligand-independent manner by PKC and MAPK pathway [173]. Retinoic acid itself increases the expression of RARβ, whereas in vitamin A-deficient rats retinol increases the expression of RARα [174]. d. Thyroid Hormone Receptors Overall thyroid hormones (THs) can increase testis size and sperm function [175, 176]. The increased testicular size may be due to the effect of TH on increasing Sertoli cell proliferation [177–179] by regulating p27 (Kip1) activity. In the Sertoli cells, the thyroid

Chapter 15 Transcription Factors in Sertoli Cells

hormone receptor α isoform is observed [180]. The two α isoforms 1 and 2 are expressed differentially during prepubertal and postpubertal stages of Sertoli cell development. The thyroid hormone binding α 1 isoform is expressed at higher levels during the prepubertal stages, whereas the nonthyroid hormone binding α 2 isoform is the predominant isoform in postpubertal Sertoli cells [181]. Thyroid hormone may also be involved in regulating the expression of AR mRNA [177, 182]. e. Orphan Nuclear Receptors SF-1 SF-1 belongs to the steroid hormone receptor superfamily with an N-terminal zinc-finger DNA binding domain. SF-1 (steroidogenic factor 1, also known as adrenal 4-binding protein, Ad4-BP), initially identified as a tissue-specific transcriptional regulator of cytochrome P450 steroid hydroxylases, is vital for endocrine development and function [183, 184]. The mouse Ftz-F1 gene locus coding for SF-1 closely resembles the Drosophila fushi tarazu factor 1 (Ftz-F1), which regulates the developmental homeobox gene fushi tarazu [185]. Unlike most other nuclear receptors, SF-1 binds to the extended CCA/GAGGTC sites as a monomer [186]. Additional DNA binding specificity is achieved through amino acid residues C terminal to the DNA binding domain, which recognizes specific nucleotides 5’ to the AGGTCA site [187]. The C-terminal ligand binding domain has no known ligand; hence, it is referred to as an orphan nuclear receptor. SF-1 expression is observed in the indifferent gonad at embryonic day 9 in mice, the earliest stage of gonadal development when the intermediate mesoderm condenses into the urogenital ridge. At embryonic day 12.5, the time of sex determination, SF-1 expression becomes dimorphic, increasing in the testis and decreasing in the ovaries. In the developing testis, SF1 transcripts are detected in both the Leydig cells and the Sertoli cells [188, 189]. The SF-1 knockout mice die shortly after birth from adrenocortical insufficiency and exhibit male-to-female sex reversal of the external genitalia. Because their gonads regress before male sexual differentiation normally occurs, the internal and external urogenital tracts of SF-1 knockout mice are female irrespective of genetic sex [190, 191]. In humans, a mutation in SF-1 leading to sex reversal has also been reported [192]. DAX-1 DAX-1 (dosage-sensitive sex reversal-adrenal hypoplasia congenital critical region on the X chromosome, gene 1) belongs to the nuclear receptor superfamily of ligand activated transcription factors based on a sequence/structure similarity with other nuclear receptors, especially at the C-terminal domain

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[193, 194]. DAX-1 lacks the zinc-finger DNA binding domain [195]. The absence of any known ligand puts DAX-1 in the orphan receptor category. DAX-1 contains a unique amino terminal domain with 3–5 repeats of a 65- to 67-amino-acid motif that forms two putative zinc-finger domains. Unlike other nuclear receptors, this zinc-finger domain may not function as a central DNA binding domain but may mediate protein–protein interactions with SF1 [196, 197], ER [198], and AR [199]. The role of DAX-1 at a post-transcriptional level has also been proposed based on the observation that DAX-1 can be found associated with polyribosomes, complexed with polyadenylated RNA [200]. Functionally, DAX-1 seems to act at multiple levels to repress the expression of genes involved in steroid hormone metabolism through a potent transcriptional repression domain present in its C terminus, which is similar to the nuclear receptors’ ligand binding domain [196]. Most of the known clinical manifestations involving DAX-1 mutations such as hypoplasia congenita (AHC), which is consistently associated with hypogonadotropic hypogonadism (HHG), are present in the C-terminal domain of DAX-1 protein. These mutations may lead to cytoplasmic retention of DAX-1 and transcriptional silencing due to the lack of interactions with transcriptional cofactors [201]. DAX-1 plays a significant role in the sex determination pathway. The DAX-1 deficiency in male mice leads to hypogonadism and sterility [202, 203]. The progressive degeneration of the germinal epithelium, independent of abnormal gonadotropin and testosterone production suggests that lack of DAX-1 impairs the function of somatic cells early during development [203]. The earliest expression of DAX-1 is seen in the Sertoli cells at the time of cord formation in the testis at embryonic day 12.5 (E12.5) in mice [204]. Between days E13.5 and E17.5, the expression of DAX-1 markedly decreases in Sertoli cells and increases in the interstitial cells [204]. In the female, DAX-1 is expressed in the entire ovarian primordium from E12.5 until E14.5. After E14.5, the expression of DAX-1 decreases and between E17.5 and postnatal day 0 (P0) is limited to cells near the ovarian surface epithelium. In contrast, DAX-1 expression is persistent in postnatal Sertoli cells but rare in Leydig cells [204]. The expression of DAX-1 in the embryonic Sertoli cells may also be regulated by Wnt-4 [205]. In humans, duplication of 1p31-p35 harboring the Wnt-4 gene is associated with upregulation of DAX-1. Interestingly, this upregulation of DAX-1 leads to XY sex reversal [205]. These results suggest that sex determination is sensitive to the dosage of DAX-1 and excess DAX-1 may in fact be antagonistic to Sry. The DAX-1 expression in postpubertal Sertoli cells peaks with the first spermatogenic wave and is

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subsequently regulated and increases during androgensensitive phases of the spermatogenic cycle [46]. In cultured Sertoli cells, DAX-1 expression is downregulated by the pituitary hormone FSH [46] (Table 15.2B). f. Cys4 Zinc Fingers GATA The GATA transcription factor family (GATA-1 through -6) contains a highly conserved DNA binding domain, consisting of two zinc-finger domains (CX2CX17CX2C), and it binds to a nucleotide sequence (A/T)GATA(A/G). Subtle sequence variations within the core response element (i.e., GATA, GATT, GATC) and flanking region may allow differential binding among coexpressed GATA family members within a specific tissue. The GATA family of transcription factors is widely expressed during embryonic stages and in the adult [206]. The targeted deletions of individual GATA proteins have revealed important functions in the development of various organs and systems, such as hematopoiesis (GATA-1, -2, and -3) and cardiogenesis (GATA-4) [206–208]. The coordinated expression of GATA-4 has been implicated in testicular development and function. GATA-4 protein is detected as early as E11.5 in the developing somatic cell lineages in the gonads of both XX and XY mouse embryos [209]. Following sex determination by E12.5–E13.5, GATA-4 expression is confined to the Sertoli cells and is markedly downregulated in the XX somatic cells, suggesting its role in sexual differentiation and determination [209, 210]. In humans, GATA-4 is expressed in the early fetal testicular development to adulthood and peaks at 19–22 weeks of gestation in Sertoli cells [211]. A similar role for GATA-4 in swine testicular development has also been proposed [212]. In support of the essential role of GATA-4 in testis differentiation, a number of studies have been performed to identify the genes activated by GATA-4, which include steroidogenic enzymes [213], steroidogenic acute regulatory protein [214], inhibin-α [215], and Müllerian inhibiting substance [209, 216]. In addition to GATA-4, the expression of GATA-6 has also been observed in late fetal, neonatal, juvenile, and adult Sertoli cells [210]. In the postpubertal Sertoli cells, GATA-6 and -4 levels are unaltered, but those of GATA-1 decrease in response to FSH [217]. GATA-1 expression is also observed in prepubertal Sertoli cells. After puberty, the GATA-1 expression is observed in Sertoli cells associated with stage VII, VIII, and IX seminiferous tubules [29]. The uniform expression of GATA-1 in the testis of germ-cell–depleted models such as W/Wv, jsd/jsd, or cryptorchid mice has led to the hypothesis that GATA-1 expression may be negatively regulated by germ cells [22]. In the absence of GATA-1, the testes develop normally with

no obvious effect on spermatogenesis and no change in the expression of putative GATA-1 target genes and other GATA factors [218]. GATA-4 knockout mice die by 9.5 days postcoitus and exhibit profound defects in ventral morphogenesis, including abnormal foregut formation and a failure of fusion of the bilateral myocardial primordial [219, 220]. This early embryonic lethality of GATA-4 knockout mice has been a limitation for studying the role of GATA-4 during various stages of gonadal development. This limitation has been partially addressed by generating knockout mice for FOG-2 (friends of GATA-2) or mice homozygous for a targeted mutation in GATA-4 that abrogates the interaction of GATA-4 with FOG [221] as discussed next. FOG FOG family proteins contain between eight and nine zinc-finger domains. These highly conserved C-X2-5CX12HX2-5H domains (CCHH ZnFs) have been shown to interact with the N-terminal domain of GATA proteins [222]. The interactions between the two members of the FOG family, FOG-1 and FOG-2, with GATA-1 and GATA-4 have been extensively studied and appear to repress the GATA-dependent transcription in cardiac morphogenesis [223, 224]. The interaction between GATA-4 and FOG-2 has also been proposed as an important step in gonadal development [29, 225]. FOG-2 expression correlates with that of GATA-4 and can be observed in the male urogenital ridge [29]. FOG-2 is transiently expressed in the mouse Sertoli cells at E12.5 [29]. The downregulation of FOG-2 expression after E12.5 may have important implications because FOG-2 can repress the GATA-4 dependent trans-activation of the MIS promoter [29, 216, 225]. The expression of FOG-2 and GATA-4 in the urogenital ridge may also be required for Sry expression. The mouse embryos homozygous for a null allele of FOG-2 exhibit significantly decreased levels of the Sry transcript at E11.5, whereas WT1 and SF1 expression is normal. In contrast, the expression of Sox9, MIS, and Dhh genes downstream of Sry were decreased. Similar results obtained in mice homozygous for a targeted mutation in GATA4 (GATA4(ki)) abrogates the interaction of GATA-4 with FOG cofactors. Collectively, these results suggest that FOG-2 and GATA-4 interactions play significant roles in various stages of sex determination and Sertoli cell differentiation [221]. g. Cys2 His2 Zinc-Finger Domains SP1 Family of GC-Box Binding Transcription Factors The presence of GC boxes has been shown to play a critical role in directing promoter specificity to the Sertoli cell. A multitude of transcription factors

Chapter 15 Transcription Factors in Sertoli Cells

expressed by Sertoli cells such as Sp1, Sp3, Egr-1, Rnf6, Ap-2, and Sp3 bind the GC box. The significance of the GC boxes is not whether they bind any cell-specific transcription factors, but more in terms of formation of a cell-specific complex that aids in stabilization of the transcriptional initiation complex. The GC-box binding transcription factors have been shown to influence Sertoli cell expressed genes such as the specificity of KL (Kit ligand) expression by Sertoli cells in response to FSH [226], ABP expression [227], stage-specific expression of L-cathepsin promoter [228], SF-1 expression [229], Dmrt1 expression [230], clusterin promoter activity [231], and activation of the TATA-less promoter Mer, a receptor tyrosine kinase [232]. Inhibin-α subunit and CREB [233] have also been shown to be directly dependent on the presence of a GC-rich region in the respective proximal promoters. Wilms’ Tumor 1 The Wilms’ tumor 1 (WT1) transcription factor has four contiguous C2H2 zinc-finger motif at the C terminal and an N-terminal transcriptional regulatory domain. Loss of function of WT1 was initially identified in Wilms’ tumor, a pediatric kidney cancer [234, 235]. Subsequently, several other complex phenotypes such as gonadoblastoma, congenital malformations, and 46XY gonadal dysgenesis were associated with the loss of WT1. The role of WT1 as a sex-determining gene was proposed based on mutations in WT1 in individuals with Danys-Drash syndrome or with Frasier syndrome, both of which resulted in gonadal dysgenesis [236]. Mice with targeted deletion of the WT1 gene may survive to birth but have no gonads regardless of their genetic sex. This observation supports the notion that, like SF-1, WT1 may act upstream of Sry. These homozygous null mice also have no kidneys and exhibit a wide range of anomalies in heart, lung, and mesothelium [237]. Consistent with these phenotypes, WT1 gene expression has been localized to Sertoli cell [238]. WT1 is a complex gene. At least 24 isoforms are now known as a result of alternative start sites, splicing, and RNA editing (U => C editing). These isoforms can lead to diverse activities of WT1 such as RNA processing and interactions with DNA as a homodimer. The transcriptional activity of WT1 is dependent on the presence of the KTS domain, a string of lysine, threonine, and serine between zinc fingers 3 and 4. The presence or absence of the KTS tripeptide can determine the strength of WT1 transcriptional activity. In general WT1 (–)KTS is considered to be a transcriptional activator, whereas the (+)KTS isoform is generally localized to the spliceosomal site [236, 239, 240]. The (–)KTS isoform can regulate the expression of MIS, Sry, and androgen receptor in Sertoli cells [241–243].

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C. Transcription Factors with a Helix–turn–Helix Motif 1. Homeobox Genes The homeobox genes contain a conserved 183-bp sequence coding for a 61-amino-acid homeodomain, which is responsible for sequence-specific DNA binding [244]. Functionally, the homeobox genes are involved in biological processes such as control of cell identity [245], cell growth and differentiation [246], cell–cell interactions, and cell–extracellular matrix interactions [247]. The diversity of regulatory pathways influenced by homeobox genes suggests that these transcription factors may have a significant role in Sertoli cell biology, from initiation of Sertoli cell phenotype to regulation of Sertoli cell function in the adult testis. Members of the homeobox family, such as Oct4 [248, 249] and PAX, have been shown to be involved in germ cell development. The only homeobox gene reported to have a role in Sertoli cell function is the Pem gene. a. Pem The orphan homeobox gene Pem encodes a homeodomain transcription factor related to the Prd/Pax gene family [250]. Pem is induced in Sertoli cells at day 9 postnatal in mice, at the initiation of meiosis in germ cells [251]. Following puberty, Pem expression is restricted to androgen-dependent stages IV–VIII of the seminiferous epithelium cycle [251]. Consistent with this observation, Pem gene expression is regulated by androgens and other cell types [252, 253]. The proximal 0.6-kb 5’-flanking sequence is sufficient for age- and stage-specific expression of the mouse Pem gene in Sertoli cells. Transgenic mice overexpressing Pem specifically in Sertoli cells have an increased number of preleptotene spermatocytes and elongated spermatocytes with double- and single-stranded breaks, respectively, with no obvious anomalies in spermatogenesis, fertility, or fecundity. Pem gene expression in Sertoli cells may be involved in the regulation of the expression of secreted or cell-surface proteins that may serve to control premeiotic DNA replication, DNA repair, and/or chromatin remodeling in the adjacent germ cells [254]. 2. LIM Homeodomain: Lhx9 Lhx9 (LIM/homeobox gene 9) encodes a transcription factor implicated in various developmental processes, including gonadogenesis [13]. In the rat, Lhx9 expression is observed in the undifferentiated gonads but disappears as epithelial cells differentiate into Sertoli cells and begin to express MIS. The mutually exclusive expression of Lhx9 and MIS suggests that Hx9 may negatively regulate MIS gene expression [255].

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3. Winged Helix: WIN-1/Trident Winged helix (WH) proteins are a large family of putative transcription factors that may regulate mesenchymal-to-epithelial transitions and maintain cellular differentiation [256, 257]. Family members share a highly conserved 100-amino-acid DNA-binding domain, which was first identified in HNF-3 proteins [257]. Targeted disruptions of a number of WH genes have revealed essential functions of WH proteins in development for cell fate determination, cellular proliferation, and cellular differentiation [257–260]. The role of WH proteins in mesenchymal-to-epithelial transitions during kidney and brain development was established by targeted disruption of BF-2 and BF-1, respectively [261, 262]. An increasing number of WH proteins are also involved in the transcriptional regulation of cell-specific genes, which suggests a role in integrating transcriptional gene networks [263–265]. A novel member of the WH family, rWIN, is expressed in Sertoli cells [266]. During embryogenesis, the expression of Trident, the mouse homologue of rWIN/HFH-11, is observed in all proliferative cells, but not in resting cells [267]. The human homologue HFH-11 is highly expressed in spermatocytes and spermatids [268]. In contrast, spermatogonia undergoing active proliferation reportedly do not express HFH-11 [268]. Long-term stimulation (72 hr) of cultured Sertoli cells with FSH decreases rWIN expression. The dynamics of the regulation of rWIN expression suggest that rWIN is transiently upregulated within 30 min of FSH stimulation. Such a transient increase in rWIN expression is analogous to the induction of the immediate early gene c-fos. The transient upregulation of rWIN expression in response to FSH may be required for the expression of Sertoli cell functional genes such as transferrin [266].

D. Beta Scaffold Factors with Minor Groove Contact 1. Rel/Ankyrin NFκB NFκB is a family of closely related transcription factors that bind to the kB DNA motif as a dimer [269]. NFκB was originally discovered in the nucleus of B cells, where it binds to the κ chain of the immunoglobulins. The various members of the NFκB family and the dimerization/processing scheme are outlined in the Figure 15.6. Within the nucleus, the activity of NFκB is also modulated by phosphorylation. The possibility of preferential dimerization and phosphorylation in response to appropriate stimuli makes NFκB an attractive transcription factor family to study mechanisms of cell-specific gene regulation in both normal and diseased states. The roles of NFκB in tumorigenesis have been extensively studied and include regulating the expression of immunoregulatory and inflammatory genes, antiapoptotic cell survival genes, cell proliferation genes, and differentiation genes [270, 271]. The RelA and p50 NFκB DNA binding activity is also present in Sertoli cells [27, 272]. Importantly, NFκB proteins can specifically bind to the κB enhancer motifs within the promoter of the CREB gene [273]. The role of NFκB in germ cell–Sertoli cell interactions has also been proposed [272]. TNFα, an NFκB-activating cytokine produced by round spermatids located adjacent to Sertoli cells, stimulates the elimination of IκB, translocates additional NFκB to the nucleus, and increases NFκB binding to CREB promoter κB enhancer elements [146, 273]. TNFα also stimulates transcriptional activity of the CREB promoter. These data demonstrate that NFκB contributes to the upregulation

FIGURE 15.6 Mechanism of action of REL proteins. The reticuloendotheliosis family (REL) of proteins is divided into two broad categories. The first consists of CREL, RELA, and RELB. The proteins are synthesized in mature forms as opposed to the second group of REL proteins the NFκB1 (p105) and NFκB2 (p100). The p105 and p100 undergo proteolytic cleavage to generate the p50 and p52 forms, respectively. This proteolytic cleavage removes the C-terminal ANKYRIN repeats. Together, the REL proteins can form the transcriptionally active homo- or heterodimers. The transcriptional activity of these dimers can be inhibited by inhibitor of κB (IκB). IκB proteins consist of an N-terminal regulatory domain followed by a series of ANKYRIN repeats.

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of CREB expression in Sertoli cells and raises the possibility that NFκB may induce other Sertoli genes required for spermatogenesis [273]. In addition, the activators of the PKA signaling pathway such as forskolin or FSH also increase NFκB DNA binding activity [27]. The limited studies and the potential role of NFκB in the regulation of cellular function make this family an attractive candidate for further research. 2. High Mobility Group High mobility group (HMG) proteins are nonhistone chromosomal proteins that contain two highly conserved DNA binding HMG-box domains. The superfamily is largely divided into three subfamilies: HMG1, HMG2, and HMGB4. Acting primarily as architectural proteins, they facilitate the assembly of nucleoprotein complexes by bending and binding preferentially to distorted DNA, thus effecting recombination and initiation of transcription [274, 275]. In contrast to other transcription factors, the HMG proteins have no transactivation domain. HMG-box proteins might be targeted to particular DNA sites in chromatin by either protein–protein interactions or recognition of specific DNA structures [276]. The significance of these proteins can be appreciated from the observations that HMG1 is one of the most abundant nuclear proteins in all mammalian cells. a. Sry Sry, a member of the Sox family of HMG box proteins, is considered a testis-determining gene and is located on the Y chromosome [277–279]. Sry is expressed for a short period of time (10.5–12 and 11.5–13 days postcoitus in the mouse and rat, respectively) in selected epithelial cells of the genital ridge and is sufficient for initiating their differentiation into Sertoli cells [2, 45, 280] (Table 15.3). The differentiation of Sertoli cells initiates testis differentiation from the indifferent gonad. Transgenic mice overexpressing Sry develop as males even with an XX karyotype and, when deleted, the chromosomally male mice adopt a female phenotype [278, 281, 282]. The molecular mechanism of Sry function is an area of intense research and is complicated by the fact that Sry has been evolving very fast, as indicated by significant sequence differences among various mouse strains [283]. Outside the consensus HMG box, the Sry sequences are highly divergent between species. Based on the function and mechanism of action of other HMG-box proteins, it can be predicted that Sry may have other partners with which it interacts via its HMG box [190, 284, 285]. In humans a single

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34-kDa protein called SIP-1 (Sry interacting protein) has been identified through yeast two-hybrid interaction studies [286]. SIP-1 is ubiquitous, nuclear, and interacts via its two PDZ protein binding domains with the most C-terminal seven amino acids in human Sry. The murine homologues of SIP-1 have not been found, possibly because the non-HMG-box segments of the Sry protein are significantly different from the human Sry. Another possibility that needs careful evaluation is that more than two proteins may be involved in this interaction, and formation of the complex is dependent on the availability of all of the proteins of the complex. Alternatively, the presence of a C-terminal trans-activation domain in the mouse Sry and its absence in the human gene may suggest that a “functional Sry unit” in humans may require interactions with SIP-1 and not in mouse. Potential phosphorylation of a protein kinase capable of phosphorylating an N-terminal domain serine residue may modulate the DNA-binding activity of Sry. A similar phosphorylation residue is absent in the mouse Sry [287]. The rapid evolution of the Sry sequence, divergence between species of regions outside the HMG box, selective protein interactions, and functional modification such as phosphorylation suggest the HMG-box alone is sufficient for the activity of Sry. b. Sox Family of Transcription Factors The expression of at least two Sox proteins, Sox3 and Sox9, is also observed in the Sertoli cells of the gonad. Sox3 is most closely related to Sry and is largely considered to be the evolutionary precursor of Sry [288]. Sox3 is expressed in both XX and XY gonads during the critical period of sex determination [289]. On the other hand, Sox9 expression is more widespread and is present in the chondrocytes, pancreas, and heart among other tissues [290, 291]. In the developing gonad, Sox9 is initially expressed in both males and females [292]. The Sox9 expression becomes restricted to the male gonad following the initiation of Sry expression, suggesting that after sex determination Sry may be involved in regulating the expression of Sox9 [13]. The localization of the Sox9-positive cells around the testicular cords suggests that it is expressed primarily in Sertoli cells [292, 293]. Unlike Sry, the expression of Sox9 is biphasic and decreases gradually until birth but is increased immediately before puberty at 15 days postnatal and persists in the adult mice seminiferous tubules [293]. The expression profile of Sox9 suggests that it is required for the important phases of aggregation and reorganization of the Sertoli cells and germ cell differentiation. The consensus motif for Sox proteins has been defined as the heptameric sequence 5’-(A/T)(A/T)CAA(A/T)G-3’ [294].

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The close association between Sox9 and Sry expression suggests that it may be important for Sertoli cell differentiation and sex determination [295–296]. Such a function is also supported by the fact that ~75% of karyotypically male patients with heterozygous Sox9 mutations develop as intersexes or XY females. Thus, Sox9 is expressed in Sertoli cells that also express Sry. However, whereas Sry expression is transient, Sox9 expression is maintained throughout further testis development. Sox8, another member of the Sox family, is also expressed in the developing mouse testis around the time of sex determination [297].

E. Others 1. STATs The STAT (signal transducer and activator of transcription) family is a group of transcription factors that translocates signals from the cytokine, hormone, or growth factor activated receptors to the nucleus. The receptor-recruited STATs are phosphorylated on a single C-terminal tyrosine. The homo- and heterodimers of STATs between phosphotyrosine of one STAT and the SH2 domain of the interacting STAT bind to sequence-specific DNA sites. Out of seven known STAT family members (STAT1–7), only STAT-3 and STAT-1 activities have been observed in response to cytokines such as LIF [85], IL-6, and IFNγ [86] in Sertoli cells. FSH appears to suppress STAT-1 and STAT-3 expression, but stimulates STAT-5 (Table 15.2C). 2. CBP Both CBP and p300 are highly homologous nuclear proteins originally cloned for their ability to interact with phosphorylated CREB [298] and with the adenovirus E1A protein [299], respectively. Both CBP and p300 are thought to function as crucial links between diverse signal transduction and transcriptional pathways and to play an essential role in the cellular processes of growth and differentiation [300–302]. Several protein motifs in CBP and p300, such as a bromodomain, a KIX domain, and three regions rich in Cys/His residues (C/H domains), are conserved in CBP/p300 species ranging from Drosophila to mammals. The CBP/p300 consists of flexible modules that can accommodate interactions with multiple activators of transcription and integrate signals derived from multiple pathways. These modules serve as binding sites for sequence-specific transcription factors and other components that regulate gene expression. The CBP/p300 interactions with transcription factors include the bHLH myogenic proteins; CREB;

the oncogene product Myb; the retinoic acid, estrogen, glucocorticoid, and thyroid hormone receptors; AP1 complex; components of the basal transcription machinery such as transcription factor IIB (TFIIB); and TATA-binding protein [300–302]. These observations suggest that CBP/p300 might constitute a physical nexus between enhancer binding proteins and components of the basal transcription machinery. In addition, CBP/p300 may disrupt repressive chromatin structures through its intrinsic or associated histone acetyl transferase activity [303, 304]. In Sertoli cells, the coactivators, CBP/p300, integrate the synergistic actions of the bHLH proteins and CREB on the transferrin promoter and are required for optimal FSH stimulation of the transferrin promoter (Fig. 15.7). Based on the effect of CBP/p300 antisense oligonucleotides on wild-type and E-box and PRII mutants of the transferrin promoter, it is speculated that CBP/p300 may form a ternary complex involving bHLH and CREB transcription factors. The formation of a ternary complex by CBP/p300 may facilitate integration of transcriptional regulators such as bHLH proteins and CREB in Sertoli cells. This is postulated to result in positive cross-talk between the transcription factors and the signal transduction pathways in Sertoli cells. The possibility that CBP/p300 may also integrate the basal transcription machinery to the bHLH and CREB transcription factors is also likely [305]. 3. Dmrt1 The presence of Dmrt1 as one of the key sexdetermining genes in mammals was proposed from the studies on C. elegans and Drosophila genes MAB-3

FIGURE 15.7 Proposed model for the regulation of transferrin gene expression in Sertoli cells. bHLH and CREB proteins are required for the transcriptional activity of the transferrin promoter. An interaction between the bHLH and CREB proteins is facilitated by CBP/p300. This cooperative interaction may be required for Sertoli cell-specific transferrin gene expression.

Chapter 15 Transcription Factors in Sertoli Cells

and doublesex, respectively [306]. MAB-3 and doublesex genes have a highly conserved DM domain with cysteines and histidines [307, 308]. The DM chelates zinc, but this zinc-binding domain is distinct from other zinc-binding motifs [309]. The DM domain is probably the most conserved sex-determining domain across phyla. Consistent with its role in sex determination, overexpression of Dmrt1 has been shown to induce sex reversal in many species. In humans, the role of Dmrt1 as a potential sexdetermining gene was obtained through karyotyping sex-reversed males that showed monosomy in chromosome 9p with a loss of the Dmrt locus that harbors the Dmrt1 gene [308, 310, 311]. In mice, the Dmrt1 gene was cloned by using a degenerate PCR approach. Dmrt1 in mouse is expressed exclusively in the XX and XY genital ridge prior to sexual differentiation. The expression becomes restricted to seminiferous cords during progression of testis development and is observed in both Sertoli cells and germ cells [312]. In the postnatal testis, Dmrt1 expression increases by postnatal day 10. As germ cells proliferate, the expression of Dmrt1 drops around the third week of postnatal development [313]. In cultured Sertoli cells, FSH and cAMP (PKA pathway) are known to upregulate expression, whereas phorbol esters (PKC pathway) downregulate Dmrt1 expression [314]. The expression of Dmrt1 in Sertoli cells is regulated by a combination of ubiquitously expressed transcription factors Sp1, Sp3, and Egr1 [230].

V. REGULATION OF SERTOLI CELL GENES BY COMBINATORIAL INTERACTIONS OF TRANSCRIPTION FACTORS The cumulative observations from a number of studies suggest that only a small number of transcription factors and their combinatorial interactions may be required to regulate the expression of large numbers of genes expressed by Sertoli cells. Examples of some of the genes known to be regulated by the combinatorial interactions are discussed next. In most of these studies, these mechanisms have been studied using in vitro systems of cultured Sertoli cells transfected with a specific promoter region of genes of interest. The strength/activity of the promoter is usually measured by its ability to induce the expression of a reporter gene such as β-galactosidase, luciferase, chloromphericol, or acetyl transferase. Alternatively, the timing of expression of a particular transcription factor (e.g., DAX-1) in relation to a potential gene of

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interest (e.g., MIS) can also provide some clues as to the regulatory mechanisms in the cell. Although these in vitro studies have yielded a great deal of useful information, these data should also be evaluated in terms of the complexity of in vivo chromatin dynamics and interactions.

A. Regulation of the Transferrin Gene Expression Cell-specific expression of the iron-binding protein transferrin is in part mediated through the regulation of its promoter [315]. Serum transferrin is produced by hepatocytes, but cells that create a blood barrier such as Sertoli cells in the testis and choroid plexus epithelium in the brain also express the transferrin to provide iron to cells sequestered within the serum-free environment. Both mouse and human transferrin promoters have been used to identify the potential conserved response elements and/or regions involved in Sertoli cell-specific expression of the transferrin gene. A remarkable (80%) homology between the proximal human and mouse transferrin promoters suggests that the transcriptional control mechanisms between the species are conserved. Comparisons of the regulatory regions and cell-specific expression (hepatocytes versus Sertoli cells) have revealed that the enhancer element in the –3600/–3300 region are functional in hepatoma cells, but are unable to increase transcription in Sertoli cells. The conserved proximal region I and II (PRI and PRII) in the proximal 150 bp of the promoter are sufficient to achieve optimal transferrin gene expression in Sertoli cell cultures. In response to FSH, CREB, or CREB-like cAMP responsive proteins, possibly C/EBPβ, can bind to the PRII site [316, 317]. The presence of E-box response elements in the proximal promoter is also known to regulate the transferrin promoter activity in Sertoli cells [98]. As an added level of control to achieve specific gene expression, the bHLH proteins bound to the E-box can physically interact with transcription factors binding to the PRII site. This interaction can be mediated through CBP/p300 [305]. The CBP and p300 are the only known histone acetyltransferases (HATs) that are capable of acetylating all four core histones. The E-box and PRII are approximately 220 bp apart, which corresponds to one turn of the DNA around the nucleosome complex. Such proximity and the intrinsic HAT activity of CBP/p300 further support the hypothesis that CBP/p300 may form a ternary complex involving bHLH and CREB proteins [305] (Fig. 15.7). The presence of negatively acting bHLH Id proteins and their expression profile in response to FSH may help to fine tune the transferrin gene expression by inhibiting the bHLH proteins [51].

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B. Regulation of the MIS Promoter In males MIS is secreted normally by fetal and adult Sertoli cells and in the females by postnatal granulosa cells. MIS regulates testicular development by initiating the regression of Müllerian ducts. The dynamics of its expression in Sertoli cells led to the hypothesis that MIS may be one of the earliest sex-specific targets of the Sry transcription factor. The role of other transcription factors involved in the sex determination pathway in the regulation of the MIS promoter also has been reported. For example, upon ligand binding, SF-1 may regulate Sertoli cell–specific expression of the MIS promoter [318]. It has been speculated that this ligand may be specific to Sertoli cells. The cell specificity of MIS expression has been localized to the proximal 180 bp of the promoter, which harbors the consensus SF-1 binding site [318]. SF-1 may help to maintain quantitative levels of MIS expression, but loss of HMG-box sequences that bind Sox9 may lead to abnormal male sex differentiation. Interactions between GATA-4 and SF-1 [319], Sox9 and SF-1 [320], and WT-1 and SF-1 [242] have also been proposed in the regulation of the MIS promoter. These interactions are mediated by direct protein– protein interactions. The transcriptional synergism between GATA-4 and SF-1 can be disrupted by DAX-1. The repression of the MIS promoter by DAX-1 may involve direct interactions with SF-1 bound to DNA and may not necessarily involve GATA-4 [321]. The mutually exclusive expression of GATA-1 and MIS during prepubertal development of Sertoli cells has led to the hypothesis that GATA-1 may inhibit MIS gene expression [322]. The presence of FOG2 in Sertoli also suggests that it may modulate GATA-4–dependent MIS expression [216]. The presence of SF-1, FOG2, GATA-4, DAX-1, and Sox9 in multiple cell lineages, but restricted MIS expression to Sertoli cells, supports a mechanism involving a Sertoli cell–specific cooperative interaction between these transcription factors.

C. Regulation of SF-1 Promoter The sexually dimorphic pattern of SF-1 expression observed during male gonadal differentiation suggests the role of similarly expressed transcription factors in its regulation. For example, the proximal SF-1 promoter harbors a conserved Sox binding site AACAAAG (Sox-BS1) [323]. This site binds Sox9 with high affinity, but other Sox proteins, which are also coexpressed with SF-1 such as Sox8 and Sox3, can also bind this site with lower affinity. In addition to the Sox-BS1 site, other sites such as the USF1 and USF2 binding E-box site, the CAAT site, sequences with

overlapping E-box and CCAAT sites, and multiple GC-rich Sp1/Sp3 are also required for optimal SF-1 promoter activity [229].

VI. CONCLUSION AND FUTURE DIRECTIONS Over the years and with an ever-increasing knowledge of the genome, the studies addressing the mechanisms involved in Sertoli cell gene regulation have matured from a linear approach that is based on characterization of specific cis-acting regulatory sequences and their cognate transcription factors to a more global approach that considers regulation in terms of the complete functional unit that simulates a specific gene expression pattern in vivo. This global approach is required to acknowledge the role of transcription factors, coactivators, corepressors, chromatin remodeling enzymes, histone acetylases, deacetylases, kinases, and methyl-transferases in regulating Sertoli cell-specific gene expression. For example, E-box response elements are required for the expression of many Sertoli cell genes, as discussed in this chapter. The consensus site for an E-box (CANNTG) occurs approximately once every 300 bp in the human genome. All of these sites are obviously not bound by bHLH transcription factors. Therefore, it is reasonable that some E-box sites have a preference in terms of sequences surrounding them or their state of methylation. In addition, accessory mechanisms may be involved that can include the presence or absence of other transcription factor coactivators such as CREB, C/EBP, and CBP/p300, as in the case of the transferrin promoter. Together these mechanisms may serve as a “homing mechanisms” that steer a particular transcription factor toward a very small subset of potential target sites. In recent studies the concept of cell-specific transcription factor complex involving many of the concepts discussed earlier is also being evaluated. For example, the transferrin gene expression may involve combinatorial interactions between bHLH, CREB, and CBP/p300 [305]. The formation of a complex on the serum response element of the cfos promoter may involve interactions between bHLH proteins and serum response factors, which may be specific to Sertoli cells [99]. A complex interplay of stage-specific expression of transcription factors and the corresponding expression of functional genes such as MIS supports this hypothesis. Promoter analysis for conserved binding sites in coregulated genes can also be useful to develop a consensus of the Sertoli cell transcriptional regulatory networks. Microarray experiments in combination with

Chapter 15 Transcription Factors in Sertoli Cells

other techniques such as genome-wide analysis of in vivo transcription factor–binding using “ChIP on a chip” (chromatin immunoprecipitation on a chip using transcription factor–specific antibodies) and specific computer algorithms will help in assembling complete Sertoli cell–specific transcriptional regulatory maps. For example, in a large-scale gene expression profiling of Sertoli cell genes, the genes can be grouped in a pattern definition process such as selecting coregulated genes with similar transcription factor core motifs. The complexity of this initial data can be further reduced by selecting a second common transcription factor motif. Following the process of verification, such as biologically relevant overexpression, reverse genetics and knockout experiments are needed. A consensus can be developed on the potential regulatory networks involving similar transcription factors and their interactions. For example, the role of the E-box elements in regulating the transferrin gene promoter was considered as a potential conserved process due to its presence and validation in a number of other Sertoli cell gene promoters. The current review helps categorize the various transcription factors in Sertoli cells. The FSH regulation of these transcription factors (Table 15.2) and developmental regulation (Table 15.3) have been also reviewed. Currently very few cell-specific transcription factors have been shown to promote or induce Sertoli cell differentiation. In contrast, observations support the proposal that unique combinations of more widely expressed transcription factors are likely involved in Sertoli cell–specific gene expression. How this wide variety of transcription factors interacts to form unique regulatory networks in Sertoli cells will be the primary future research to elucidate the transcription regulation of Sertoli cell differentiation and development.

Acknowledgments The authors wish to thank Dr. Derek McLean and Dr. Michael Griswold for sharing the rat Sertoli cell microarray data. The authors also wish to thank Mr. Jim Shima and Ms. Jacqueline Ague for analysis of the microarray data.

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C H A P T E R

16 Structure and Regulation of the FSH Receptor Gene LESLIE L. HECKERT Department of Molecular and Integrative Physiology, University of Kansas Medical Center, Kansas City, Kansas

cells, which causes an increase in intracellular calcium levels [11, 12]. At the cellular level, FSH stimulation of Sertoli cells results in increased protein synthesis, increased estradiol synthesis, and increased transcriptional activity of specific target genes [7, 13, 14]. The biological ramifications of FSH action in the testis are to induce Sertoli cells to establish their full cellular complement within the seminiferous tubules and to promote their maturation and ability to assist germ cell development, ultimately contributing to the final spermatogenic capacity of the testis [15–20]. Importantly, delivery of the FSH signal depends on the correct expression of its receptor, and thus Fshr gene regulation is a critical component of the final biological effect of the hormone. Therefore, efforts to fully understand the mechanisms required for FSH signaling in the testis have included numerous studies focused on identifying the mechanisms responsible for activation and expression of the Fshr gene in Sertoli cells. Other interests in the Fshr gene have focused on identifying the cell-specific events responsible for the highly restricted expression pattern observed for Fshr. Because Fshr is only expressed in Sertoli cells in the male, it appears likely that the transcription factors that direct this exquisite cell specificity will not only be relevant to Fshr gene expression but will be key proteins that define gene expression patterns unique to Sertoli cells. To date, analysis of the transcriptional mechanisms required for Fshr expression have been limited to studies on its proximal promoter region, using techniques that

I. INTRODUCTION: FSH RECEPTOR BIOLOGY II. FSHR AND ITS GENE III. TRANSCRIPTIONAL REGULATION OF FSHR—THE FSHR PROMOTER References

I. INTRODUCTION: FSH RECEPTOR BIOLOGY Follicle-stimulating hormone (FSH) and testosterone are the two major signals that control sperm production in the testes. Both of these hormones elicit their actions through specific receptors located in Sertoli cells, which, when bound by hormone, stimulate various cellular processes necessary for Sertoli cells to properly nurture the developing germ cells. The receptor for FSH, i.e., the follicle-stimulating hormone receptor (Fshr), is located at the cell surface, where it contains both extracellular and intracellular domains connected by an α-helical segment that spans the membrane (Fig. 16.1; for a review of Fshr see [1]). Fshr is a member of the G-protein coupled receptor (GPCR) family that, accordingly, couples to a small intracellular GTPase (G-protein) that links FSH binding to cellular response (Fig. 16.1). In males, Fshr is expressed exclusively in Sertoli cells, where it preferentially couples to Gs, thereby inducing adenylyl cyclase activity and synthesis of the second messenger cAMP [2–10]. Although the mechanism is not as well understood, FSH also regulates ion channels on the plasma membrane of Sertoli SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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FIGURE 16.1 The different domains of the FSH receptor partitioned into the 10 exons of its gene (see color plate). Fshr is a 675-amino-acid cell-surface receptor with a prominent amino-terminal domain, a rhodopsinlike transmembrane domain, and a carboxy-terminal intracellular domain (top). Comparison of the protein structure to the encoding exons of the gene (bottom) revealed that the extracellular portion of the receptor is encoded by the first 9 exons and the transmembrane/intracellular domain is encoded by exon 10. Exons 2 through 8 are similar in length (68–77 bp, listed on top of each exon) and contain one of the nine leucine-rich repeat structures that characterizes the receptor’s extracellular domain. Colored portions of the receptor protein match the color of the corresponding encoding exon (bottom, rectangles). The number of amino acids encoded by exons 1 through 9 is represented by a similar number of colored segments. Intron sizes, in base pairs, for rat (blue) and human (red) are indicated in the intervening regions between exons (bottom).

Chapter 16 Structure and Regulation of the FSH Receptor Gene

include transient transfections, DNA/protein binding assays, and transgenic mice. Although our understanding of Fshr expression has grown with respect to the mechanisms controlling basal transcription and its regulation by FSH, the mechanisms that drive cellspecific expression remain elusive. This chapter summarizes our current understanding of the Fshr gene and the transcriptional processes that are necessary for its expression and regulation in Sertoli cells.

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pairs (68–77 bp) and coding for one of the nine leucinerich repeat structures that characterizes the receptor’s extracellular domain (Fig. 16.1) [21, 25, 26, 29–32]. The amino acids encoded by exons 2, 3, 4, and 7 follow very similar repeat patterns, whereas those from exons 5, 6, and 8 differ slightly at their carboxy termini. Exon 9 diverges from exons 2–8 with respect to size and the specific repeat pattern, but does have an arrangement of aliphatic amino acids characteristic of a leucine-rich repeat [26]. Exon 10 is the largest of the exons and encodes the entire intracellular domain and all seven of the putative membrane-spanning domains (Fig. 16.1).

A. Cloning Fshr

B. The Fshr Gene and Evolution

Cloning of the Fshr cDNA in 1990 had a significant impact on our understanding of the receptor, by providing new insight into its protein structure and relationship to other known receptors ([21]; for additional reading, see [1, 14]). Fshr was identified as a 675-aminoacid cell-surface receptor with a prominent aminoterminal domain located on the extracellular surface of the Sertoli cell (Fig. 16.1) [21]. Cloning of the receptor also revealed that this domain consists of a series of leucine-rich repeats that are required for hormone recognition and binding and are typified by a pattern of similarly positioned aliphatic amino acids (leucine, isoleucine, valine, and phenylalanine) [21–23]. The cDNA sequence also showed that Fshr’s amino-terminal domain is tethered to the cell through a rhodopsinlike transmembrane segment that threads through the membrane seven times and ends with a string of 63 amino acids in the intracellular space (Fig. 16.1). This transmembrane-spanning domain is predicted to exist predominantly as an α-helical structure and is characteristic of all members of the GPCR family (reviewed in [24]). Cloning of the Fshr gene soon followed publication of its cDNA, providing important insight into the exonic partitioning of the different receptor domains and the evolutionary relationship of Fshr to other members within the gene family [25–27]. The Fshr gene was initially cloned and characterized from a rat genomic library and revealed a structure consisting of 10 exons and nine introns that stretch along more than 84 kb of DNA sequence (Fig. 16.1) [26]. Identical structures were confirmed in several other species, including human, murine, and tammar wallaby [25, 27, 28]. Comparison of the protein structure to the encoding exons of the gene identified an intriguing division of receptor domains (Fig. 16.1). The extracellular portion of the receptor is encoded by the first nine exons. Exons 2 through 8 have notable structural similarity, with each exon consisting of a similar number of base

Within the GPCR family, the FSH receptor is part of a subfamily that includes the glycoprotein hormone receptors, luteinizing hormone receptor, thyroid stimulating hormone receptor, and other more recently discovered leucine-rich repeat-containing GPCRs or LRGs [21, 24, 31–33]. Among members of this GPCR subfamily, the extracellular domain, like that of Fshr, consists of a series of leucine-rich repeats that function in hormone binding and recognition [22, 23, 26, 34, 35]. Phylogenetic analysis identified three distinct subgroups for the glycoprotein hormone/LRG receptors and based on their protein sequence similarities, the glycoprotein hormone receptors and LRGs from sea anemone, Caenorhabditis elegans, and Drosophila were placed into one subgroup while the other LRGs (LGR4, LGR5, snail LRG, relaxin receptor) were divided into the other two [32, 36]. A common evolutionary origin for these receptors was also evident from the striking structural similarities among their genes. For each, the large extracellular domain is partitioned among a continuous series of similarly sized exons that, typically, contribute a single leucine repeat to the receptor, whereas the rhodopsin-like transmembrane/intracellular domain is encoded by a single, large exon (Fig. 16.2) [22, 23, 26, 29–31, 34, 37–40]. The number of leucine-rich repeats differs among some of these receptors and this is the predominant factor responsible for the variation in exon number observed between receptors. Notably, the division of one leucine-rich repeat into a single exon has led to speculation that the receptor’s extracellular domain evolved from tandem duplication of an ancestral leucine-rich module [26]. The evolutionary relationship between these receptors is also evident from the similarity in intron phasing of the genes. For the Fshr, Lhr, Tshr, and sea anemone genes, the phasing is identical with each intron in phase 2, whereas for the Drosophila Lrg-1 gene, 12 out of 16 introns are phase 2 with seven positions identical to that of the mammalian

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FIGURE 16.2 Gene structures for the glycoprotein hormone and sea anemone receptors (see color plate). Exons are depicted as rectangles with their number given below and their size in base pairs given above. Gray boxes represent leucine-rich repeats and the blue boxes the seven membrane-spanning domains. The structures of the glycoprotein hormone receptor genes were derived from the rat Fshr, human TSHR, and rat Lhr [26, 29–30]. The structure for the sea anemone receptor gene was derived from studies by Vibede et al. [37].

genes [26, 29, 30, 41]. Thus, it appears that Fshr, Tshr, and Lhr share a common ancestral gene with the Drosophila Lgr-1 and sea anemone receptors, and this ancestral gene likely arose prior to cnidarian evolution [37].

C. Fshr and Genome Analysis Advancements The explosion of genome sequencing projects has provided a wealth of new sequence data on the genomes from a wide range of species and has afforded new opportunities to disclose functionally important regions through the use of comparative genomics [42]. This approach takes advantage of known sequences from two or more species to identify evolutionarily conserved sequences, based on the assumption that these sequences are under selective pressure to be maintained and are therefore functionally important, i.e., exons or gene regulatory elements. DNA sequences of the rat and human FSHR/Fshr genes were used to perform pairwise sequence analysis and identified more than 40 nonrepetitive conserved regions (Fig. 16.3) [43]. Ten of these corresponded to exons (Fig. 16.3, triangles), which were between 83% (exon 1) to 93% (exons 3 and 8) identical. In addition, there are 41 conserved sequences (Fig. 16.3, circles 1–41) that did not correspond to Fshr exons, other known coding sequences, or repetitive sequences, and these ranged from 76% to 96% identity.

By comparing the location of the conserved sites on each gene, it was observed that the sites are conserved not only with respect to their sequences but to their arrangement on the gene as well, suggesting that both sequence and position are important to their function. The pattern of conserved sites also showed a cluster of highly conserved sites located 5’ flanking sequence and first intron. DNase I hypersensitivity mapping was used to examine part of this region in the rat Fshr gene and showed that conserved sites 4, 5, and 7 were hypersensitive to DNase I digestion, identifying them as sites of altered chromatin structure. Because hypersensitivity often indicates functionality, the correlation between conservation and hypersensitivity supports the value of the comparative genomics approach and suggests that the analysis of each site will dramatically extend our understanding of Fshr gene regulation, which is currently limited to studies on the proximal promoter region.

III. TRANSCRIPTIONAL REGULATION OF FSHR—THE FSHR PROMOTER A. General Promoter Features Studies on the rat, murine, ovine, and human FSHR/Fshr genes have provided a significant amount

Chapter 16 Structure and Regulation of the FSH Receptor Gene

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FIGURE 16.3 Annotation of the aligned sequences for the rat and human Fshr genes (see color plate). The sequence of the rat Fshr gene (supercontig accession #NW_044190) was aligned to the human FSHR gene (chromosome 2 genomic contig accession #NT_022184) using the default parameters for pairwise Blast analysis through the National Center for Biotechnology website. All sequences having greater than 75% identity were annotated and labeled 1 through 41. Exon positions are denoted as flags. Each conserved sequence is marked with a colored circle and a corresponding line that marks the position of the sequence on both the human and rat genes.

of information on Fshr transcriptional regulation. These studies have focused exclusively on characterization of the 5’ flanking sequences and have revealed both important similarities and differences between the promoters of the four species. Comparison of the four promoter sequences revealed a significant level of conservation within the first 1050 bp upstream of the translational start codon, indicating an evolutionary pressure to maintain the sequences and implicating functional importance for this region (Fig. 16.4) [27, 44–47]. Examination of the aligned sequences has assisted with the identification of potential regulatory elements in the Fshr promoter and provided a measure of conservation for those functional elements identified experimentally (Fig. 16.4) [26, 45–47]. Some of these elements, such as the E box, are conserved among each of the species, whereas others, such as FP2 in the ovine promoter, appear to be limited to a specific species. The transcriptional start sites were mapped for each of the above species, and their relative positions to that of the translational start codon differed significantly (Fig. 16.4, bent arrows). In the rat gene, transcriptional start sites were mapped to positions –80 and –98 relative to the start codon [26]. These sites correspond well to those identified for the human gene but differed considerably from those reported for the murine and ovine genes, which were located at positions –534 and –163 bp, respectively (Fig. 16.4 and [45–47]).

Sequences around the transcriptional start sites for the human and rat genes are within highly conserved regions of the promoter, whereas that around the murine start site is partially conserved and that of the ovine site appears to be unique. In addition, examination of the sequences near the transcriptional start sites of each species revealed that initiation of Fshr does not require a canonical TATA motif, a feature that is found in more than half of the known human genes [48]. Different classes of TATA-less core promoters have been described and include those rich in cytosine and guanine (CpG-rich promoters), as well as those dependent on an initiator element (Inr). This latter element is found within the rat and human FSHR/Fshr core promoters (i.e., the transcriptional start sites and flanking sequence that extends ~35 bp upstream and or downstream) and is defined as a discrete element that is functionally similar to the TATA box but can function independently [49]. Because core promoter characteristics are thought to contribute to combinatorial gene regulation, this particular feature of the FSH receptor promoter may likely determine its response to specific transcription factors and thus help direct its cell specificity. Our current understanding of the Fshr promoter comes predominantly from studies that employed transient transfection and DNA/protein binding studies to identify important regulatory elements and their

FIGURE 16.4 Aligned sequences for rat, murine, human, and ovine Fshr promoters (see color plate). Sequences for the rat, murine, human, and ovine Fshr promoters were aligned, with adenosine of the translational start codon denoted as +1. Bent arrows mark the transcriptional start sites with the corresponding species and initiation base shown for each. Base positions are indicated for the rat gene in the left margin. Identified DNA elements or footprinted regions (FP1–FP4 ovine) are boxed in blue and encompass only species in which the element was characterized. Red asterisks mark cytosines within the rat Fshr promoter that were differentially methylated in nonexpressing and expressing cell types. Red bases shaded in gray indicate 100% identity between species and blue shaded bases indicate 75% identity.

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cognate binding proteins. This approach has provided insight into basal promoter function, but unfortunately has revealed little with regard to the mechanisms responsible for cell-specific expression [44, 50–53]. The rat, murine, ovine, and human promoters have been analyzed using different promoter lengths, reporter vectors, and cell systems that should be recognized when comparing promoter function and considering the observed differences. In the following four sections, the studies contributing to our understanding of the Fshr promoter are summarized, with each section devoted to results from a single species.

B. The Rat Fshr Promoter 1. Cell Specificity The 5’-flanking region of the rat gene has been used extensively in studies on Fshr transcriptional regulation in Sertoli cells. In efforts to reveal the mechanisms that direct cell-specific expression of Fshr, transient transfection studies were used to compare basal promoter activity across different cell types, both Fshr expressing and nonexpressing. However, the success of this approach has been limited, because promoter activity was not entirely restricted to those cells that express the receptor and, thus, characterization of basal promoter activity has not identified any candidate transcription factors that restrict Fshr expression to Sertoli cells [52, 54]. However, studies examining induced promoter activity in Fshr-negative cells showed that the orphan nuclear receptor steroidogenic factor 1 (Sf-1) directly regulates the rat and mouse promoters and has provided new insight and potential mechanisms for Fshr cell specificity [51, 55]. At this juncture, the role of Sf-1 in determining the expression profile of Fshr has not been explored but its presence in Sertoli cells and its limited expression profile has made it a good candidate for directing cell specificity. However, Sf-1 cannot act alone in directing Fshr transcription to Sertoli cells because it is also expressed in several additional tissues that lack Fshr. Therefore, if Sf-1 helps limit Fshr expression, it is likely part of a combinatorial array of transcription factors that directs Fshr cell specificity. Unfortunately, our current understanding of the mechanism directing Fshr cell specificity is incomplete, and the available data suggest that sequences outside of the investigated promoters are needed to direct cell-specific expression of Fshr.

FIGURE 16.5 Relative activity of rat Fshr promoter mutants. Transient transfection analysis was used to measure promoter activity of different amounts of 5’ flanking sequence from the rat Fshr gene. Activity is relative to the largest promoter construct (–5000 bp). (Adapted from [52].)

important regulatory elements that reside both 5’ and 3’ to the transcriptional start sites [50, 52, 54, 56]. Promoters of various lengths were included in the transcriptional studies of the rat Fshr gene, with the largest fragment extending out approximately 5000 bp 5’ to the start of translation [52, 54]. Transient transfection analysis of various promoter deletions indicated that most of the regulatory elements necessary for basal promoter function reside within the first 200 bp upstream of the ATG start codon. Accordingly, deletion of the rat promoter to this position did not adversely effect transcriptional activity, when assayed in either primary cultures of Sertoli cells or the mouse Sertoli cell line, MSC-1 (Fig. 16.5) [50, 52, 54]. In fact, promoter activity increased with some deletions, suggesting that repressor elements are located between –5000 and –2800 and between –313 and –197 bp, relative to the translational start codon (Fig. 16.5 and [52]). Promoter regions associated with transcriptional repression were also observed in the mouse and ovine promoters, but, to date, no mechanisms have been uncovered to explain repression through these sites. Because the majority of basal activity was found to be within the first 200 bases of the promoter, emphasis was initially placed on identifying important elements within this region. 3. The Proximal Promoter

2. Promoter Function

a. The E Box

Analysis of the rat Fshr promoter, using transient transfections and mutagenesis, has identified several

Transient transfection analysis of various mutant promoters in either primary rat Sertoli cells or MSC-1

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FIGURE 16.6 Schematic representation of the rat Fshr proximal promoter (see color plate). The rat Fshr gene is represented by the gray rod. Exon 1 is colored blue and important regulatory elements are colored gray. The DNA sequences across the Inr, E2f, and E box are given. Red bases within the E2f site represent those identified as most functionally important. Red bases within the Inr region represent the reverse GATA sequence. Bent arrows show the transcriptional start sites and ATG represents the translational start codon. The graph (lower left) represents the relative activity of each E box promoter mutant (sequence is shown to the right of the graph) relative to that of the wild-type promoter. Bases in lowercase signify the mutation and underlined bases denote the core E box sequence. The series of Usf dimers above the E box denotes their relative binding abundance when assayed in vitro using Sertoli cell nuclear extracts. Starred bases within the E box designate those protected from methylation by in vivo footprint analysis. (Adapted from [52].)

cells identified the E box, Inr, and E2f regulatory elements (Fig. 16.6) [50, 52, 56]. Mutation of the E box had the single largest impact on rat Fshr promoter activity, whereas more modest effects were observed with mutations in the E2f and Inr sites. In general, an E box is a DNA element that has the consensus sequence CANNTG and is bound by transcription factors in the basic helix–loop–helix (bHLH) family. The Ns located in the central two positions represent any base but most commonly are represented by either CG or GC. In the rat promoter, the E box (5’-CACGTG-3’) is located from –119 to –124 and shows a high degree of sequence conservation with the other species (Fig. 16.4). With the exception of a single G-to-A change at the first guanine of the human sequence (5’-CACGTG-3’ to 5’-CACATG-3’), each of the six core bases is conserved. Mutagenesis of bases within the core and flanking sequences of the E box revealed that a change in the core’s central CG dinucleotide to either GC or GA diminished promoter activity 60–80% (Fig. 16.6) [52]. Surprisingly, mutation of these central bases was better tolerated than changes in the flanking bases outside the element (5’-GGTCACGTGACTT-3’ to 5’-GtaCACGTGtaTT-3’). To determine if the human E box sequence was functional, the fourth base of the

rat sequence was changed from guanine to adenine (5’-GTCACGTGACTT to GGTCACaTGACTT) and the promoter activity determined. In this case, changing the rat sequence to that of the human had little impact on promoter activity, suggesting that the E box is functional in the human promoter (Fig. 16.6). Further investigation of the E box included studies on the endogenous Fshr gene using in vivo genomic footprinting. To this end, primary rat Sertoli cells were treated with dimethyl sulfate (DMS) and, following isolation of genomic DNA, the methylation status of promoter guanines was determined by piperidine cleavage and ligation-mediated PCR [53]. Comparison of the methylation patterns around the Fshr E box from DNA isolated from DMS-treated Sertoli cells, DMS-treated Fshr-negative cells, and untreated Sertoli cells identified two core guanines within the Fshr E box that were protected from methylation in cultured Sertoli cells but not in untreated DNA or DMS-treated samples from the nonexpressing cells (Fig. 16.6, starred bases) [53]. These data showed that there are protein(s) bound to the Fshr E box on the endogenous gene, thereby blocking the ability of DMS to methylate these sites. Importantly, these studies also suggest that occupancy of the E box depended on active transcription of the Fshr gene.

Chapter 16 Structure and Regulation of the FSH Receptor Gene

b. The USF Proteins Protein/DNA binding studies were used to characterize factors bound to the E box in vitro and identified the transcription factors Usf1 and Usf2 as the predominant Sertoli cell proteins bound to this element [52, 53]. These ubiquitously expressed transcription factors are members of the bHLH family, and within this family, form part of a subgroup designated the basic helix– loop–helix zip (bHLH-Zip) proteins [57–59]. In addition to the HLH and basic domains, proteins in this subgroup have a leucine zipper domain that contributes to protein dimerization. Like other family members, the target element for Usf1 and Usf2 is an E box, and for these transcription factors, the preferred element has the same sequence as that found in the Fshr promoter (RYCACGTGRY) [60, 61]. To bind DNA, Usf1 and Usf2 must form either homodimers or Usf1/ Usf2 heterodimers, and studies using nuclear extracts from either Sertoli cells or whole testis indicated that Usf1 homodimers and Usf1/Usf2 heterodimers are the predominant forms binding the Fshr E box (Fig. 16.6) [52, 53]. Binding of Usf2 homodimers to the E box was barely detectable. Additional support for Usf1 and Usf2 in Fshr transcriptional regulation was provided by studies that demonstrated a strong correlation between bases required for promoter function and those required for Usf binding (Fig. 16.6). Thus, mutations with the greatest impact on promoter activity also had the greatest impact on Usf binding and vice versa [52]. More direct evidence for Usf regulating Fshr transcription was obtained by cotransfection of wild-type and mutant forms of the Usf proteins with Fshr promoter/ reporter constructs. In these studies, the Usf proteins activated the Fshr promoter via a mechanism that required binding to the proximal E box [53]. In addition to Fshr, Usf1 and Usf2 have been shown to regulate a number of different genes but an in vivo or physiological role for these transcription factors has not been firmly established. Mice lacking either Usf1 or Usf2 were developed by two different groups and both demonstrated mild to moderate phenotypic changes [62, 63]. This contrasts with the dramatic developmental defect (embryonic lethality) observed when both transcription factors are lost and shows that Usf1 and Usf2 direct critical developmental events and compensate for each other when one gene is absent [62]. However, compensation was incomplete, because postnatal defects were apparent in some tissues. For example, in the liver, Usf1 homodimers could not completely substitute for either the Usf1/Usf2 heterodimers needed for transcriptional response to glucose or the Usf2 dimers required for hepcidin expression [64, 65]. Furthermore, Usf-2 null mice showed a modest

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growth defect and males had reduced fertility and a decreased life span, while Usf1-deficient mice were prone to epileptic seizures [62]. The fertility defect observed in Usf-2 null mice has not been studied in detail, so it is uncertain if Fshr expression is altered or if it is a contributing factor to the decreased fertility. Because the Usf proteins appear to be an important regulator of the Fshr gene, it is intriguing how these ubiquitously expressed proteins participate in the highly cell-specific transcriptional mechanism for Fshr. The observation that the Usf proteins regulate transcription through the E box and that the E box of the endogenous Fshr gene is occupied by proteins only in Fshr-positive cells indicates that the promoter is accessible to the Usf proteins in expressing cells, whereas in nonexpressing cells the promoter is nonpermissive for Usf binding. A likely explanation employs a series of Sertoli cell transcription factors, perhaps including Sf-1, whose combined properties restrict Fshr expression. These proteins would function within the Fshr gene and facilitate binding of Usf to the proximal E box, thereby activating transcription. This facilitated binding may occur through direct recruitment of the Usf proteins to the promoter or from induced changes in the promoter, such as methylation of bases within the E box that influence Usf binding or in the chromatin configuration around the proximal promoter, which would then permit binding of the Usf proteins to the E box. c. The E2f, GATA, and Inr Elements In addition to the E box, the E2f, GATA, and Inr elements reside within the core promoter region of Fshr, but the extent to which these elements have been studied is considerably less when compared to the E box. Like the E box, though, each of these elements was implicated in control of the endogenous Fshr gene by in vivo genomic footprint analysis, which showed these sites were protected from methylation in DMStreated Sertoli cells [50, 55, 56]. Analysis of various promoter mutants by transient transfections and in vitro protein/DNA binding studies have confirmed the functional importance of these sites and demonstrated sequence-specific interactions between Sertoli cell nuclear proteins and each of these elements [52, 56]. For the E2f site (5’-TTTTCGCGC-3’), placement of mutations within the Fshr promoter (positions –38 to –46) that included either the entire site or more refined, smaller mutations showed that this element was necessary for full promoter activity [52]. Examination of the element’s sequence revealed a strong match with a consensus E2f site and partial conservation between the rat, human, mouse, and ovine promoters (Fig. 16.4). If the site is functional for

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each promoter, the conserved and likely relevant bases do not conform to an E2f binding site, suggesting that a different protein is responsible for the promoter activity. Characterization of the E2f site showed that the largest impact on promoter activity occurred with mutation of the second GC pair, which decreased promoter activity to the same degree (~50%) as a full block-replacement mutant (Fig. 16.6, red bases) [52]. Interestingly, the position of this dinucleotide is conserved in each of the species (Fig. 16.4). DNA/protein binding studies showed that several Sertoli cell protein complexes bound to the element but none were identified [56, 66]. However, studies that employed an antibody against E2f1, one of three activators in the E2f family, suggested that it is not a component of the protein complex bound to this element [56]. Therefore, at this juncture the element has been identified but the protein that elicits the transcriptional activity is unknown. The functional contribution of the Inr to Fshr transcriptional activity was determined by characterization of promoter/reporter constructs carrying 3’ promoter deletions and specific mutations within the element [50, 52]. The sequence spanning the major transcriptional start sites (–103 to –75) has two Inr-like sequences that cover both start sites and resemble the consensus Inr sequence, PyPyAN[T/A]PyPy (reviewed in [49]). Sequence-specific protein complexes bound this region (–117 to –30) and were competed by a wellcharacterized Inr from the deoxynucleotidyltransferase gene, demonstrating similarity in the recognition sequences of the protein complexes binding these Inrs [50]. Within the region between the two transcriptional start sites, there is an inverted GATA sequence, TATC, that, when mutated, significantly diminished Fshr promoter activity [56]. The importance of this site suggested that members of the GATA transcription factor family function either independently or in conjunction with the Inr binding proteins to regulate Fshr promoter activity [56]. A role for the transcription factor GATA-1 was implicated, as protein/DNA binding studies demonstrated that a GATA-1 antibody cross-reacted with Sertoli cell proteins bound specifically to this site. In addition to GATA-1, several other unidentified proteins interact within this promoter region and may also contribute to Fshr transcription [56]. GATA-1 is a well-known regulator of genes in hematopoietic cell lineages and belongs to a family of transcription factors that function in tissue differentiation and development and bind to the consensus sequence WGATAR [67]. The Fshr GATA motif differs slightly from this consensus, where a guanine replaces the weak base prior to the GATA sequence (Fig. 16.6).

In addition to its expression in erythroid cells, GATA-1 is found in the Sertoli cells of the testis, where it is produced from a testis-specific mRNA that is transcribed from a promoter located 5’ to the erythroid transcript’s first exon [68, 69]. The remainder of the transcript is generated by exons that are shared in the two cell types and encodes the GATA-1 protein. GATA-1 expression in the testis occurs in a stage-dependent manner in which it is restricted to stages VII, VIII, and IX of mouse spermatogenesis [68]. This is similar to that for the rat Fshr, which is first detected in Sertoli cells just prior to sperm release at stage VIII with the highest expression observed during stages IX and X [70, 71]. However, because there are differences in rat and mouse seminiferous cycles, the exact level of overlapping expression is not currently known but is likely sufficient to support a mechanism of direct regulation by GATA-1. d. Regulation by Steroidogenic Factor 1 and cAMP More recent studies on the rat promoter demonstrated that the orphan nuclear receptor Sf-1 regulates Fshr transcription [51]. In these experiments, transcriptional activity rose substantially when an expression vector for Sf-1 was added to an Fshr promoter/ transfection assay [51]. Cotransfection of the Sf-1 expression vector with various promoter mutants revealed that two regions were needed for full Sf-1 response: one between –2800 and –743 and the other within the first 200 bases of the promoter. Within the core promoter, mutation of the E box eliminated all activation by Sf-1, demonstrating a requirement for this element in response to Sf-1. Notably, the E box was not only required for the response observed within the core promoter but also for response generated through the upstream site(s). Thus, the E box appears to play an important role in coordinating the response to this transcription factor. Binding studies using the Fshr E box and either in vitro translated Sf-1 or nuclear extracts containing the protein failed to show direct binding of Sf-1 to the E box. This suggested that Sf-1 functions by interacting with proteins bound to the E box, thereby implicating the Usf proteins in integration of the Sf-1 signal [51]. The proposed interaction with the Usf proteins was further evaluated by examining Sf-1 activation of the Fshr promoter in the presence of dominant negative forms of Usf1 and Usf2. The results demonstrated that in the presence of mutant Usf proteins, lacking either the transactivation domain or the DNA binding domain, activation by Sf-1 was significantly attenuated. Thus, stimulation by Sf-1 requires interaction with the Usf transactivation domain [51]. Notably, this Usf-dependent Sf-1 activity was unique to the Fshr

Chapter 16 Structure and Regulation of the FSH Receptor Gene

promoter, as altering Usf levels did not impact Sf-1’s ability to activate the equine luteinizing hormone α-promoter. Although the mechanism of Sf-1 and Usf cooperative regulation is not fully understood, the finding that these transcription factors act in concert to induce transcription has provided important insight into Fshr gene regulation and the transactivation function of Sf-1. Sf-1 regulation was also observed with the mouse Fshr promoter and was shown to require a specific binding site for induced promoter activity (see later discussion) [55]. This binding site (SLBS-3; CAAGGACT), however, did not align with sequences in the

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5’ flanking region of the rat Fshr gene (~35,000 bp) but did matched a sequence nearly 6800 bp into the first intron. This particular site has not been evaluated in terms of regulating the rat gene. Unfortunately, identification of specific upstream sites required for Sf-1 regulation of the rat promoter was problematic. Sf-1 response of sequential promoter deletions resulted in incremental decreases in transactivation and specific sites could not be determined with significant accuracy. This suggests that, within the upstream region of the rat promoter, response to Sf-1 is due to multiple Sf-1 binding sites that spread over the region from –2800 to –763 (Fig. 16.7A).

FIGURE 16.7 Summary of the regulatory features for Fshr promoters from different species (see color plate). The (A) rat, (B) human, (C) mouse, and (D) ovine Fshr promoters are represented. The 5’ flanking region of each gene is signified by a rod. Areas of the promoters that are shaded either red or green represent regions with a negative or positive transcriptional effect, respectively. Exon 1 is shown as a blue cylinder and bent arrows denote the transcriptional start sites. Positions of the transcriptional start sites, relative to the translational start codon ATG, are given above the arrows. Names of identified regulatory elements are shown in black lettering below the genes and their corresponding positions are indicated as cylinders. Green cylinders represent positive regulatory elements and red cylinders represent negative ones. Proteins identified to bind a regulatory region are shown on top of their binding region as shaded circles or ovals, with the corresponding name of the factor adjacent to it. Sequences for the mouse SLBS-3 and ovine CACC sites are provide over the elements. Black numbers above the promoter indicate base positions, in kbp, relative to the translational start codon. The green shaded bar below the rat promoter (A) shows the Sf-1 responsive region and the orange arrow denotes the interaction between Sf-1 and the Usf proteins. Dotted red or green lines delineate repressor or activator regions, respectively, common to at least three promoters.

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Sf-1 is an important regulator of endocrine function and sex determination and its expression is primarily restricted to cells of the gonads, adrenal, pituitary, and ventral medial hypothalamus, where it regulates transcription of specific genes within these tissues [72–86]. This orphan receptor regulates transcription of genes encoding important steroidogenic enzymes as well as those involved in the production of the gonadotropin hormones, i.e., αGSU, luteinizing hormone (LH) β subunit, gonadotropin releasing hormone receptor, and inhibin α subunit [77–90]. With Fshr added to the list of Sf-1 target genes, regulation of the FSH axis can be finely tuned through Sf-1 activity in the testis, which affects both hormone production, through regulation of testicular inhibin, and hormone response through regulation of Fshr [82]. Sf-1 activates the inhibin α subunit promoter and this is further stimulated in the presence of activated protein kinase A (PKA), the major intracellular signal stimulated by FSH [82]. The actions of Sf-1 on inhibin suggest that, in the presence of Sf-1, increasing FSH levels will significantly enhance inhibin levels, which feed back on the pituitary to decrease FSH production. Remarkably, activated PKA was shown to block Sf-1 stimulated transcription of the Fshr promoter [51]. Hence, with Sf-1 poised to regulate these two genes, FSH stimulation (via PKA activation) can produce opposing transcriptional effects that result in increased α inhibin and decreased Fshr, a result that would dramatically decrease FSH signaling in Sertoli cells. The role of FSH and cAMP on Fshr levels in Sertoli cells has been the focus of many studies. In Sertoli cells, Fshr mRNA levels are significantly reduced in response to FSH or agents that stimulate the cAMP pathway [91–93]. In contrast to earlier studies, more recent reports demonstrate that this decrease in Fshr occurs at the transcriptional level [91, 92]. In these studies, a direct transcriptional effect was indicated by demonstrating that the transcriptional inhibitor actinomycin D blocks receptor mRNA downregulation and by the observation that heteronuclear Fshr RNA decreased in the presence of FSH [91, 92]. Notably, there were no negative effects of FSH observed on the –2700-bp Fshr promoter but rather, unexpectedly, activity of the –383-bp promoter increased nearly threefold in the presence of FSH, an event that required an AP-1 site located at position –213 (Figs. 16.4 and 16.6) [91]. FSH-induced protein complexes bound the AP-1 element and were shown to cross-react with an antibody against the transcription factor c-fos. Although these studies did not identify a specific mechanism for Fshr repression, they did provide evidence that an AP-1 complex containing c-fos may activate Fshr under some physiological conditions [91, 94]. As indicated in

the previous studies, cAMP dramatically decreases Fshr promoter activity when stimulated by the transcription factor Sf-1. This observation may explain the inability to observe decreased promoter activity under the standard transient transfection conditions discussed earlier. Thus, in the transfection studies, the level of Sf-1 may be insufficient to observe the effects of cAMP. e. Studies in Transgenic Mice Studies in transgenic mice were used to delineate regions of the Fshr promoter needed for cell-specific expression. Two separate studies employed 5000 bp of the rat promoter to drive reporter gene expression in transgenic mice [53, 54]. In the first study, β-galactosidase was used as a reporter and its expression was measured in various tissues by Northern blot analysis [54]. Evaluation of liver, thyroid, kidney, ovary, testis, and epididymal RNA samples revealed expression only in testis and ovary of two independent lines. The results suggested that the first 5000 bp of the Fshr 5’ flanking sequence was sufficient for expression in the correct tissues, but, unfortunately, there was no confirmation that this occurred in the correct cell types. In the second study, reverse transcriptase polymerase chain reaction (RT-PCR) was used to measure the reporter Cre recombinase in RNA samples collected from testis, ovary, heart, lung, liver, kidney, brain, bladder, stomach, spleen, and eyes [53, 95]. These studies, in addition to the 5000-bp promoter, included a second transgene having only 198 bp of 5’ flanking sequence. For 8 of the 5000-bp Cre transgenic lines examined, expression was observed in both testis and brain, while three additional lines had considerable expression in other tissues. Further examination of the temporal expression pattern of the transgene revealed a significant difference when compared to that of the endogenous gene. Thus, in Sertoli cells, Fshr is expressed throughout postnatal development, and if the required transcriptional machinery is within the transgene, it should also be present at these times. However, when assayed early in postnatal development (i.e., 10 days), transgene expression was absent. The smaller –198 Cre transgene lacked the ectopic brain expression observed with the larger construct and was only present in the testis. However, like the –5000-bp Cre transgene, its temporal expression did not match that of the endogenous gene. Closer inspection of the –198-bp Cre animals revealed that testis expression in mice either 27 or 50 days of age was mostly, if not entirely, due to inappropriate expression in germ cells. These inaccurate expression patterns led to the conclusion that the promoter sequences did not contain enough information to properly restrict or express the receptor to Sertoli cells of the testis.

Chapter 16 Structure and Regulation of the FSH Receptor Gene

Notably, however, examination of the cell types expressing the transgene was limited to studies on the smaller construct, leaving open the possibility that Cre expressed from the 5000-bp promoter occurred in Sertoli cells at a later time than the endogenous gene. If true, this would suggest different transcriptional mechanisms for early (embryonic/early postnatal) and late (postpubertal) expression of the Fshr gene. Interestingly, studies on the paralogous LHR gene provided support for this theory, because they suggested different mechanisms for early and late Lhr transcription [96]. Two kb of the murine Lhr promoter behaved remarkably similar to the Fshr promoter in transgenic mice. Hence, three of five Lhr transgenic lines expressed the transgene in the testis but failed to express it in the ovary and all lines showed ectopic expression in the brain. Furthermore, testis expression of the Lhr transgene was not observed between weeks 1 and 3 after birth, but appeared at 5 wk of age in germ cells and Leydig cells. Because the transgene did not follow the temporal expression of the endogenous gene, separate transcriptional mechanisms for early and late expression were implicated, which, as indicated earlier, may also be relevant to Fshr expression. f. Methylation and Chromatin Structure in Fshr Expression Evidence that structural changes in the Fshr promoter contribute to its regulation was provided by data on methylation and chromatin structure. This suggested that Fshr transcription is regulated by methylation/ demethylation of CpG dinucleotides as well as the location of two key nucleosomes across the promoter. In mammals, significant evidence suggests DNA methylation plays a critical role in regulating gene expression [97–99]. The contribution of this modification to cell-specific gene regulation is implicated by the numerous genes with limited expression profiles that exhibit different DNA methylation patterns in expressing and nonexpressing cells and by the observation that gene demethylation correlates with gene activation [97, 100, 101]. The Fshr core promoter does not contain CpG islands but does have a few CpG dinucleotides that are potential sites of methylation [26]. Bisulfite-based DNA sequencing identified seven specific CpG sites within the Fshr core promoter that were methylated in nonexpressing cells and unmethylated in Sertoli cells (Fig. 16.4, bases marked with red asterisks) [66, 102]. The role of methylation in Fshr expression was further explored in the mouse Sertoli cell line MSC-1, which does not actively express Fshr, and showed that the inactive state of Fshr transcription correlated with cytosine methylation of the core promoter [102].

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In these studies, treatment of MSC-1 cells with 5-azacytidine, a chemical that blocks DNA methylation, reactivated transcription of the gene and demethylated the Fshr promoter [103–105]. The correlation between promoter methylation and Fshr gene activation suggests that methylation blocks the access of needed transcription factors to their cognate binding elements. Within the promoter, four of the seven potential 5-methylcytosine residues are located within known protein-DNA binding sites, suggesting that methyl group(s) within cis-acting regulatory elements directly inhibit binding of trans-acting factors (Fig. 16.4). Notably, methylation of the promoter’s E box did attenuate binding of nuclear extracts from Sertoli cells [66]. Nucleosome reconstitution experiments were used to examine the chromatin structure of a 319-bp region of the Fshr proximal promoter, and nucleosome locations were mapped using hydroxyl radical footprinting [44]. This showed that one nucleosome was positioned tightly over a region of the promoter from approximately –200 to –400 bp and a second one positioned weakly over the region from –100 to +100 (Fig. 16.8). Interestingly, the E box appeared to be localized to the linker region and the CpG methylation sites were predominantly situated within the second nucleosome. These data on methylation and chromatin structure

FIGURE 16.8 Nucleosome positions on the rat Fshr proximal promoter (see color plate). Schematic representation of nucleosome positions within 319 base pairs of the rat Fshr promoter as determined by reconstitution studies with hydroxyl radical footprinting. One nucleosome was positioned very tightly over the region of the promoter from approximately –200 to –400 bp (noted as fixed), while the second nucleosome was positioned very weakly over the region from –100 to +100 (noted as mobile). The E box, Inr, and E2f elements are represented as gray boxes in their approximate positions relative to the nucleosomes. Bent arrows represent the transcriptional start sites and location of the seven CpG methylation sites (*) are shown. Red cylinder represents the nucleosome core and the blue ribbon the DNA.

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suggest a model whereby the tightly positioned or fixed nucleosome helps define the promoter location. The promoter is repressed when methyl binding proteins bind methyl groups within the promoter, recruit histone deacetylases and other transcriptional inhibitors, and decrease mobility of the second nucleosome to restrict access to parts of the promoter [106, 107]. In addition, methylation of the E box in the linker DNA and perhaps methylation of other sites such as E2f might directly prevent transcription factor binding. Activation of the Fshr gene would occur when the promoter is demethylated, followed by transcription factor binding and displacement of the second nucleosome. At this juncture, however, the mechanisms that regulate DNA methylation and chromatin structure of the Fshr gene are unknown. It is noteworthy, however, that the proximal 231 bp of the human Fshr promoter, although 80% homologous to the rat and murine promoters, lacks all seven specific CpGs, suggesting that promoter methylation is not a key regulatory step for the human FSHR gene [45].

C. The Human FSHR Promoter The human FSHR promoter has been the subject of few studies, with only one known publication focusing on characterization of FSHR transcriptional activity [45]. As noted earlier, the transcriptional start sites for the human gene are positioned close to those of the rat and there is no obvious TATA sequence. Using primary rat Sertoli cells, Gromoll et al. [45] examined various promoter deletions by transient transfection analysis and showed that removing sequences between –1484 and –225 increased promoter activity nearly twofold (Fig.16.7B). In addition, removal of either the region between –94 and –59 or between –25 and –1 resulted in a significant loss in promoter activity (65 and 90%, respectively) when examined in either Chinese hamster ovary cells or primary cultures of Sertoli cells. The upstream site (site 2, Figs. 16.4 and 16.7B) overlaps with the Inr and GATA sites that were characterized in the rat promoter. The second identified site (site 1, Figs. 16.4 and 16.7B) shows a high degree of sequence conservation but when tested in MSC-1 or primary cultures of Sertoli cells, functional analysis markedly contrasted with that of the rat promoter, which showed that deletion of this region had no effect on promoter activity [52].

D. The Murine Fshr Promoter Like the human, there is a paucity of information on the murine Fshr promoter. Initial studies cloned and sequenced the promoter, identified the transcriptional

start sites, and performed a minimal number of promoter studies (Fig. 16.4) [47]. Preliminary analysis of 5500 bp of the promoter indicated that activity was poor when assayed in primary granulosa cells (1.8-fold above promoterless control), and in contrast to observations with the human promoter, the murine promoter was inactive in Chinese hamster ovary cells [45, 47]. Subsequent studies compared activities of different promoter lengths in several cell types [55]. The longest promoter fragment included 1548 bp of the 5’ flanking sequence, relative to the translational start site, and had the greatest activity in KK-1 and mLTC-1 cell lines (granulosa and Leydig cell lines, respectively), while activity in human embryonic kidney, HEK293, or MSC-1 cell lines was four to five times lower. Deletion to –1110 bp reduced promoter activity only slightly in MSC-1 and HEK cells but resulted in an 85 and 69% loss in mLTC-1 and KK-1 cells, respectively (Fig. 16.7C). Further deletion to –867 recovered all the activity in MSC-1 and HEK 293 cells and part of the activity in KK-1 and mLTC-1 cells, indicating that a repressor element resides in this region. Finally, deletion to –99 or removal of the region between –555 and –99 resulted in complete loss of promoter activity. Because Sf-1 is abundant in steroidogenic tissues, the authors postulated that this transcription factor accounted for the high level of Fshr promoter activity in the two steroidogenic cell lines, KK-1 and mLTC-1 [55]. Further investigation showed that activity of the larger 1548-bp promoter but not the 1100-bp promoter correlated with the endogenous Sf-1 levels in cells used for the transfection analyses. This suggested that Sf-1 regulates the promoter through the region between –1548 and –1100. Examination of this region identified two Sf-1-like binding sites, SLBS-2 and SLBS-3, of which only SLBS-3 (GCCAAGGACT) was shown to bind Sf-1 and be important for Sf-1 mediated Fshr transcription (Fig. 16.7C). Notably, Sf-1 activation of the Fshr promoter was revealed for both the mouse and rat, but differences are apparent in the upstream promoter regions that contain the Sf-1 regulatory elements, with a single site implicated in the mouse (SLBS-3) and multiple weak sites implicated in the rat.

E. The Ovine Fshr Promoter In the past few years, a considerable amount of data has accumulated on the transcriptional regulation of the ovine Fshr promoter. Using this promoter, transcriptional activity was predominantly measured by transient transfection analysis in a mouse Sertoli cell line, 15P-1, and a porcine granulosa cell line, JC-410 [108]. The largest promoter fragment assayed extended

Chapter 16 Structure and Regulation of the FSH Receptor Gene

to –1268 relative to the translational start site and in JC-410 and 15P-1 cells, promoter activity was 25 and 7 times above values for a promoterless control, respectively. Deletion to –807 increased activity by 70 and 24% in 15P-1 and JC-410 cells, respectively (Fig. 16.7D). Interestingly this repressor region is similar to one observed in the mouse promoter and overlaps a region of repression observed in the human promoter (Fig. 16.7). An additional deletion to –519 bp completely eliminated promoter activity in 15P-1 cells, but had no impact in the granulosa cell line. Critical regulatory regions were not identified at similar locations in the other three promoters. Next, deletion to –363 increased promoter activity (6X in 15P-1 and 5.5X in JC-410), again indicating a region of repression (Fig. 16.7D). After deletion to –269, nearly all promoter activity was lost in both cell lines. Remarkably, increased promoter activity was observed in 15P-1 cells with further deletion to –174, which overlaps repressor regions observed in the human and rat promoters. Additional studies on the ovine promoter employed DNase I footprint analysis and identified five protein binding sites within the –519 promoter [108–111]. One of these sites corresponded to the E box, which, in sheep, is located approximately 35 bases 3’ to the transcriptional start site. As in the rat promoter, the ovine E box bound to the transcription factors Usf1 and Usf2 and mutation of the E box reduced promoter activity. A second footprint region (CACC site) was identified between –46 and –67 upstream of the major ovine promoter or –209 to –230 relative to the translational start (Fig. 16.4, FP2) [111]. Curiously, this site is within a larger region identified as having repressor activity, but mutation of the element reduced Fshr promoter activity, suggesting that a complex arrangement of positive and negative elements resides in this region (Fig. 16.7D and [108, 111]). DNA/protein binding studies implicated a member of the Krupple transcription factor family bound this site, but no protein was identified. Comparison of the CACC sequences between the difference species showed that it is unique to the ovine promoter, indicating that the element likely plays a role specific to sheep (Fig. 16.4). Additional studies with the ovine promoter examined proteins binding to a region called RE-3, which is located between –360 and –334 relative to the ATG codon (Fig. 16.4) [112]. DNA/protein binding studies identified sequence-specific complexes binding this region, and antibody supershift assays indicated that the site potentially binds Usf, c-fos, c-jun, Sf-1, and chicken ovalbumin upstream promoter transcription factor-1/2 (COUP-TF I/II; JC-410 cells). COUP-TF binding to FP1, which contains the downstream E box, was also suggested by partial interaction of the binding

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complex from 15P-1 cells with anti-COUP-TFI/II antibodies. In the Sertoli cell line, a mutation within RE-3 (AATAGTGACCCACAGGGACAGTCTTA, mutated bases underlined) reduced promoter activity nearly 70% and abolished phorbol ester stimulation of the promoter [112]. Furthermore, cotransfection studies showed slight activation of this site with a Sf-1 expression vector and repression by COUP-TF, the latter of which also occurred though FP1. Hence, in sheep, it appears that RE-3 is a composite element that is activated by complexes that contain c-fos/c-jun, Usf-1/2, or Sf-1 and that the COUP-TFs repress promoter activity by interfering with the function of these transactivators (Fig 16.7D). Finally, characterization of footprint region 4, FP4, showed that within 15P-1 extracts, the retinoic acid receptor bound this region and studies with the –519-bp promoter demonstrated that the Fshr promoter was repressed approximately twofold in the presence of all-trans retinoic acid and this repression was lost on mutation of the RAR binding site (Figs. 16.4 and 16.7) [110].

F. Summary In comparing data generated from promoters of the different species, it is challenging to come to any generalized mechanism for Fshr transcription. Whereas it seems likely that much of the data apply to Fshr transcription in general, at this point it is difficult to determine if the observed mechanisms are applicable to all or only the examined species. However, several similarities do emerge, suggesting that there are some shared regulatory features (Fig. 16.7). In particular, the E box and its binding proteins Usf1 and Usf2 appear to be important to both rat and sheep Fshr transcription and a region containing this element was important to the mouse promoter. Therefore, it appears that the E box plays a central role in regulation of Fshr transcription, and the Usf proteins, because they are the major binding proteins, are the implicated regulatory proteins. The orphan nuclear receptor Sf-1 was also shown to regulate transcription in two of the four studied promoters, indicating that its regulatory role extends beyond just one species. However, the mechanisms in rat and mouse appear slightly different with respect to the elements used for Sf-1 regulation. A second consistency in the function of the Fshr promoter is the presence of areas of repression that were found in three of the four promoters (enclosed by dashed red lines in Fig. 16.7). One area is located approximately 1000 bp upstream of the ATG codon and is shared between the human, mouse, and ovine promoters. The second region is present in the rat, human, and ovine promoters and is located within the proximal

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promoter region just upstream of the E box. Despite these observations, however, no additional studies have identified the regulatory elements or proteins responsible for the repression. In summary, our knowledge of Fshr transcription has grown significantly during the last decade, but there are clearly areas that require additional exploration in order to better understand how the receptor is restricted to Sertoli cells and how repression is employed in receptor regulation. Although additional answers can clearly be obtained through continued analysis of the promoter region, the identification and characterization of regulatory elements outside the immediate promoter region appears to hold the most promise for advancing our understanding of Fshr transcription. The boon in sequence data from many different species has made this second area of investigation much more attainable, and therefore, it is anticipated that comparative genomics matched with molecular and genetic approaches will escalate our understanding of the regulatory mechanisms required for Fshr expression.

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VI CELL–CELL INTERACTIONS INVOLVING SERTOLI CELLS

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17 The Role of the Sertoli Cell in Spermatogonial Stem Cell Fate MARTIN DYM AND LIXIN FENG Department of Cell Biology, Georgetown University School of Medicine, Washington, D.C.

differentiating germ cells crowded between them. Thus, the germ cells are partially separated from one another by thin laminar Sertoli cell processes (Fig. 17.1). In the early 1970s, we described two compartments within the seminiferous epithelium: a basal compartment occupied by the spermatogonia and early spermatocytes and an adluminal compartment occupied by the more advanced germ cells [2]. We postulated that the tight junctions between adjacent Sertoli cells form an important part of the blood–testis barrier [2]. Substances entering the seminiferous epithelium have ready access to the spermatogonia in the basal compartment, but in order to reach the differentiating germ cells in the adluminal compartment these bloodborne substances such as hormones or growth factors would have to traverse the Sertoli cell. The basal aspects of the Sertoli cell and the adjacent basement membrane form the microenvironment or “niche” that allows for spermatogonial stem cell development [3, 4]. Thus, because the microenvironment in which germ cell differentiation occurs is provided by the Sertoli cells, it is reasonable to suggest that the spermatogenic process, including the regulation of the spermatogonial stem cells, must be largely controlled by this cell type. The complex organization of the seminiferous epithelium as seen in Figure 17.1 supports the concept that the polarity of the Sertoli cell in vivo must be crucial in regulating the germ cells. Thus, the basal portion of the cell is highly specialized and will secrete substances that may act directly on the adjacent spermatogonia, whereas the apical portion of the cell will secrete other substances that have direct access to the

I. INTRODUCTION TO SERTOLI CELLS IN MAMMALS II. SOMATIC CELL–GERM CELL TOPOGRAPHY IN CAENORHABDITIS ELEGANS III. SOMATIC CELL–GERM CELL TOPOGRAPHY IN DROSOPHILA MELANOGASTER IV. SPERMATOGONIAL STEM CELL RENEWAL AND DIFFERENTIATION IN MAMMALS V. RECEPTOR SIGNALING ON TYPE A SPERMATOGONIA IN MAMMALS VI. SIGNALING IN THE GERM LINE IN CAENORHABDITIS ELEGANS AND DROSOPHILA MELANOGASTER VII. FATE OF GERM LINE STEM CELLS: RENEWAL VERSUS DIFFERENTIATION VIII. SUMMARY AND CONCLUSIONS References

I. INTRODUCTION TO SERTOLI CELLS IN MAMMALS Researchers have known for many years that the Sertoli cell, the supporting element in the testis, regulates germ cell renewal and differentiation. In a colorful account of the structure of the testis [1], Elftman spoke of the Sertoli cells “standing on the basement membrane in patterned array not unlike trees in an orchard.” The spermatocytes and early spermatids are arranged in vertical rows along the “trunks” or columns of the Sertoli cells. The deeply excavated lateral aspects of the Sertoli cells conform to the convex contours of the SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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FIGURE 17.1 Sertoli cells and the blood–testis barrier (see color plate). A diagram depicting the morphology of the Sertoli cell and its close relationship to the adjacent germ cells. Localization of the blood–testis barrier and the compartmentalization of the germinal epithelium by tight junctions between adjacent Sertoli cells is also depicted. The primary barrier to substances penetrating from the interstitium is the myoid cell layer. The majority of cell junctions in this layer are closed by a tight apposition of membranes. Over a small fraction of the tubule surface, the myoid junctions exhibit a 200-Å-wide interspace and are therefore open. Material gaining access to the base of the epithelium by passing through open junctions in the myoid layer is free to enter the intercellular gap between the spermatogonia and the Sertoli cells. Deeper penetration is prevented by occluding junctions on the Sertoli–Sertoli boundaries. The tight junctions constitute a second and more effective component of the blood–testis barrier. In effect, the Sertoli cells and their tight junctions delimit a basal compartment in the germinal epithelium, containing spermatogonia and early spermatocytes, and an adluminal compartment, containing the majority of the spermatocytes and spermatids [2]. Substances traversing open junctions in the myoid cell layer have direct access to cells in the basal compartment, but to reach the cells in the adluminal compartment, substances must pass through Sertoli cells.

spermatocytes and/or spermatids. Perhaps this is one of the reasons why the coculture of germ cells with Sertoli cells has not allowed for full germ cell differentiation in vitro. This may be due in part to the fact that Sertoli cells in culture largely lose their polarity.

When grown in culture on basement membrane components, the Sertoli cells regain partial polarity but apparently this is still not sufficient to allow for spermatogonial entry into meiosis in vitro [5]. However, a recent report demonstrated that coculture of spermatocytes with Sertoli cells can produce sperm with fertilizing ability from these earlier germ cells and the subsequent production of pups after single cell injection into eggs. [6]. A number of reviews have been published on various Sertoli cell–germ cell interactions [7–11]. From these reviews and the many other publications describing Sertoli cell–germ cell interactions, a great deal of novel and important information has become available regarding the structure and the secretory products of the Sertoli cell, the cellular interactions between Sertoli cells and germ cells, signaling patterns in the Sertoli cells, and the role of the tight junctional complexes between Sertoli cells; however, there is no clear understanding of the role of the Sertoli cell in the spermatogenic process. In 1857, Sertoli wrote that “the function of these cells is still obscure to me” ([12], translated by Brian Setchell [13], pp. 55–63). This statement is still valid today. The focus of this review will be on Sertoli cell–spermatogonial interactions with an emphasis on the spermatogonial stem cells. Some information is available in the mammalian system on this topic, but a great deal of elegant work has also been carried out in model systems such as Drosophila melanogaster and Caenorhabditis elegans. The latter two systems provide simpler models to explore the coordinated events occurring between somatic cells and germ cells in the gonad.

II. SOMATIC CELL–GERM CELL TOPOGRAPHY IN CAENORHABDITIS ELEGANS The gonad in the nematode C. elegans is a relatively simple model system for studying germ cell–somatic cell interactions. There is a linear organization in the development of the male germ line cells with mitotic cells at one end of an elongate gonad and maturing gametes at the opposite end, as shown in Figure 17.2 [14–16]. Distally, there resides a single somatic distal tip cell (DTC) that could be considered the homologue of the Sertoli cell. The DTC provides the niche that allows for stem cell renewal and differentiation. Next to the distal tip cell is a region where the germ cells are mitotically active, and it is this region that houses the germ line stem cell and transient amplifying germ cells. This latter group is an intermediate population of committed cells with limited proliferative capacity and a restrictive

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FIGURE 17.2 Gametogenesis in C. elegans. The mitotic germ cells that are most distal in the gonad are in contact with the distal tip cell, a somatic element. The mitotic germ cells undergo a series of divisions to give rise to transient amplifying cells; the latter then enter meiosis and in the male sperm are formed; in hermaphrodites sperm and eggs are produced. (Panel a courtesy of Crittenden et al. [15].)

FIGURE 17.3 Stem cell renewal and differentiation in Drosophila.

differentiation potential. The transient amplifying cells give rise to the meiotic pachytene spermatocytes and, finally, in males sperm are formed and in hermaphrodites some sperm but mostly oocytes are produced. The sequential arrangement of the germ cells in C. elegans from the distal to the proximal region is similar to the arrangement observed in a cross section of a seminiferous tubule, with the early germ cells at the base of the tubule and the more mature elements near the tubule lumen. Similarly, a close relationship of somatic cells to early germ cells is present both in mammals and C. elegans as depicted in Figures. 17.1 and 17.2.

III. SOMATIC CELL–GERM CELL TOPOGRAPHY IN DROSOPHILA MELANOGASTER Stem cell renewal and differentiation in Drosophila has been examined extensively by a number of investigators [17, 18]. The general pattern of germ cell differentiation and the relationship to a somatic element are remarkably similar in flies and mammals. At the apical tip of the testis (Fig. 17.3), a group of nonmitotic somatic cells is packed tightly in an area called the hub. An average of seven to nine germ line stem cells surround the hub (Fig. 17.3), and each stem cell is flanked by two somatic cyst progenitor cells. These are

At the bottom of the figure, hub cells are surrounded by eight germ line stem cells. Each stem cell (S) then undergoes an asymmetric division, giving rise to another stem cell as well as to a gonialblast. The gonialblasts continue to divide, producing a pair of interconnected spermatogonia (Spga) then eventually 16 interconnected cells that exit mitosis and enter meiosis and terminal differentiation. The stem cells are surrounded by two cyst progenitor cells and the gonialblasts and the spermatogonia are each surrounded by two cyst cells. The stem cells and the cyst progenitor cells always remain in contact with the hub cells. (Courtesy of Yamashita et al. [95].)

the precursors or the stem cells of the cyst cells that appear to be the homologue to the Sertoli cells in mammals. In an asymmetric division, the germ line stem cell is able to give rise to another germ line stem cell that maintains contact with the hub as well as to a gonialblast cell that moves away from the hub and differentiates. Contact with the hub allows the germ line stem cell to retain stem cell identity, and as the germ line stem cell migrates away from the hub it differentiates. The germ cells continue a series of transient amplifying divisions with incomplete cytokinesis to give rise to 16 interconnected cells, now called spermatogonia, still flanked by two cyst cells. This is remarkably similar to the division of connected spermatogonia in mammals described by Dym et al. in 1971 [19].

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IV. SPERMATOGONIAL STEM CELL RENEWAL AND DIFFERENTIATION IN MAMMALS Spermatogonia proliferate actively, maintain their own numbers, and produce large numbers of spermatocytes. Thus, the spermatogonia are considered to be a stem cell population since the cells are able to renew themselves as well as give rise to differentiated elements. Primates, including monkeys and humans, have three types of spermatogonia: Adark, Apale, and B [20, 21]. The Adark are considered to be a reserve type of stem cell, whereas the Apale comprise a renewing type of stem cell. The type B cells are the differentiated spermatogonia. In rodents, a similar scheme of spermatogonial renewal with a reserve stem cell, the A0, and a renewing stem cell group, the A1 to A4, has also been described [22]. A second pattern of spermatogonial renewal was described several years later by Huckins [23] and Oakberg [24]. In this latter scheme, the stem cell is the Astem or As, a cell that is isolated along the basement membrane of the tubules. The As can divide to either renew itself or give rise to a paired group of cells, the Apaired (Apr). These then produce, in turn, Aaligned (Aal), A1 to A4, In, and type B spermatogonia. This is the

scheme that is accepted by most investigators working in rodent spermatogonial renewal and proliferation [25, 26] and is depicted in Figure 17.4. Confusion remains in the literature because the As, Apr, and Aal were called undifferentiated spermatogonia [23]. Clearly, because the Apr and Aal are committed to differentiate toward sperm (i.e., there is no turning back), this term seems incorrect because these cells are already functionally differentiated. Indeed, de Rooij and Russell suggested that the term undifferentiated be dropped when describing these spermatogonial subtypes [27]. In summary, all mammalian species have two principal categories of spermatogonia: a spermatogonial stem cell and spermatogonia that are undergoing various phases of differentiation. These latter cells may be referred to as transient differentiating cells (Fig. 17.4), a term that has been used to describe similar categories of cells in other stem cell renewing systems.

V. RECEPTOR SIGNALING ON TYPE A SPERMATOGONIA IN MAMMALS Signals from the somatic Sertoli cells are believed to regulate germ cell proliferation and differentiation.

FIGURE 17.4 Spermatogonial renewal and differentiation in rodents. One proposed pattern of spermatogonial renewal and differentiation in rodents is depicted in this diagram. In all species there is a spermatogonial stem cell; this may be called the Astem. This cell renews itself and also gives rise to more differentiated spermatogonia, referred to as transient differentiating spermatogonia. In adult mice and rats, nine such spermatogonial subtypes, Apr to type B, have been described, as shown in the diagram. The type B spermatogonia, the most differentiated spermatogonia in this class of cells, divide by mitosis to give rise to preleptotene spermatocytes (Pl); these cells enter the first meiotic prophase to give rise to leptotene spermatocytes (L); the leptotene cells differentiate through zygotene (Z), pachytene (P), and then become secondary spermatocytes (II). These divide to become round spermatids that proceed through spermiogenesis to form a fully mature spermatid in the seminiferous epithelium; upon release into the tubule lumen the late spermatid is called a spermatozoon.

Chapter 17 The Role of the Sertoli Cell in Spermatogonial Stem Cell Fate

FIGURE 17.5 Illustration of c-kit/PI3-K signaling in type A spermatogonia. This model demonstrates that c-kit/PI3-K promotes cell cycle progression via the AKT/p70S6K/cyclin D3 pathway.

Studies on growth factor–receptor systems on type A spermatogonia seemed to remain dormant until the early 1990s when Yoshinaga et al. [28] reported that in the mouse, the type A1 to A4 spermatogonia possess the c-kit receptor and are therefore dependent on stem cell factor, but the Astem (As) are stem cell factor independent. Once the c-kit receptor was identified in the germ cell population, it was hypothesized and later demonstrated that stem cell factor (SCF) is produced by Sertoli cells [29, 30]. In the past few years, a number of reports have highlighted the importance of stem cell factor/c-kit in spermatogenesis [31–33] (Fig. 17.5). In one study, addition of SCF to cultured type A spermatogonia (A1 to A4) from prepubertal mice stimulated their entry into mitosis and significantly reduced apoptosis in these cells [34]. The presence or absence of c-kit in type A spermatogonia may possibly be used as a marker for differentiation [35, 36]. We demonstrated that SCF activates phosphatidylinositol

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3’-kinase (PI3-K) and p70 S6 kinase (p70S6K) and that rapamycin, an inhibitor of FRAP/mTOR, completely inhibited BrdU incorporation induced by SCF in primary cultures of spermatogonia [37]. SCF induced cyclin D3 expression and phosphorylation of the retinoblastoma protein. Furthermore, AKT but not protein kinase C zeta (PKC-ζ), is used by SCF/c-kit/PI3-K to activate p70S6K. Dominant negative AKT-K179M completely abolished p70S6K phosphorylation induced by the constitutively active PI3-K catalytic subunit p110. Constitutively active v-AKT highly phosphorylated p70S6K, which was totally inhibited by rapamycin. Thus, SCF/c-kit uses a rapamycin-sensitive PI3-K/ AKT/p70S6K/cyclin D3 pathway to promote spermatogonial cell proliferation. Two other reports in in vivo models were published in the same year that emphasized the importance of the PI3-K pathway in spermatogonial proliferation and differentiation [38, 39]. A point mutation in the kit receptor that disrupted PI3-K binding of SCF to its receptor and reduced PI3-K–dependent activation of AKT by 90% resulted in decreased fertility in males without affecting other important kit responses. These latter reports demonstrated that the kit/SCF receptorinduced activation of PI3-K is essential for male fertility. Thus, a single signaling pathway downstream from the binding of SCF to the kit receptor can be disrupted and lead to infertility in the male. These appear to be the first reports of the importance of a Sertoli cell product in spermatogonial proliferation/differentiation that resulted in male infertility. From the above brief summary, it is clear that the c-kit receptor and its Sertoli cell produced ligand, SCF (kit ligand, steel factor, mast cell growth factor), play a significant role in postnatal development of the testis, particularly in the regulation of the more differentiated type A1 to A4 spermatogonial population and male fertility. Meng et al. described a novel growth factor/receptor system in the testis [36] consisting of the ligand, glial cell line–derived neurotrophic factor (GDNF), and a receptor signaling complex that includes the Ret receptor tyrosine kinase and GDNF Family Receptor-α1 (GFR-α1). When occupied by GDNF, GFR-α1 dimerizes and forms a complex with GDNF and Ret [40], and then Ret is activated and mediates the intracellular response [41–43]. GDNF is produced by Sertoli cells [36, 44, 45]. Other studies by Meng et al. have linked GDNF to the proliferation of Asingle spermatogonia [36], because transgenic mice overexpressing this factor show an accumulation of Asingle spermatogonia in the seminiferous epithelium [11]. They also showed that transgenic mice overexpressing GDNF exhibit testicular tumors identical to classic seminoma in men [46]. Moreover, GDNF and its receptors are strongly

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FIGURE 17.6 Regulation of spermatogonial stem cell fate. The spermatogonial stem cell can be directed to one of three fates by the Sertoli cells: renewal, differentiation, and death. It is believed that the Sertoli cells release growth factors (e.g., GDNF) that activate receptors on the surface of the spermatogonia.

expressed in human seminomas [44]. Thus, the germ cell that is at the origin of seminoma might be an Asingle spermatogonia, and a deregulation of the GDNF/ GFR-α1/Ret system in that cell could be part of the etiology of this neoplasm. In our own recent work in collaboration with MarieClaude Hofmann (unpublished), we used immunomagnetic bead procedures and isolated a GFR-α1 population of type A spermatogonia from mouse testes; most of these cells were the type As and Apr spermatogonia and they proliferated in response to GDNF. Furthermore, gene expression profiles in the GFR-α1 group of cells revealed many similar genes when compared to hematopoietic stem cells. Figure 17.6 depicts a Sertoli cell and a spermatogonial stem cell resting on the basement membrane of the seminiferous epithelium and the three possible fates of the stem cell in response to growth factors produced by the Sertoli cells. The detailed molecular mechanisms by which GDNF promotes self-renewal and/or differentiation in germ line stem cells are still unknown. The action of GDNF also seems to be potentiated by several other growth factors. In neurons, GDNF works in synergy with TGFβ to promote survival [47, 48]. Other reports have demonstrated the importance of a combination of growth factors, including GDNF, bFGF, EGF, and LIF, for the proliferation of gonocytes in vitro [49, 50]. Although it is likely that the proliferation and survival effects of activated Ret are mediated in germ cells through the same pathways as in nerve cells, it is also

well known that the same receptor systems can work differently in different cell types. The GDNF/GFR-α1/Ret signaling processes in highly purified populations of spermatogonial stem cells require further elucidation. The BMP4 signal transduction pathway is another novel mechanism in mouse type A spermatogonia that has been described recently by Pellegrini et al. involving BMP4/ALK3/SMAD5 [51]. BMP4 is produced by Sertoli cells and its receptor ALK3 is present on type A spermatogonia; addition of BMP4 to cultured spermatogonia leads to increased proliferation and differentiation. The bone morphogenetic proteins (BMPs) are a large group of proteins within the TGFβ family and include BMP2, BMP4, BMP7, and growth and differentiation factor 5 (GDF5), and the Drosophila orthologues decapentaplegic (Dpp) and 60A. BMPs play crucial roles such as providing instructive signals during embryogenesis and maintaining and repairing bone and other tissues in the adult [52]. In the regulation of the establishment of the mammalian germ cell lineage, it was revealed that BMP4 from the Dpp class and BMP8b from the 60A class, both of which are expressed in the extraembryonic ectoderm of pregastrula and gastrula embryos, induce the formation of primordial germ cells, the precursors of spermatogonia and oogonia, from the proximal region of E6.25 epiblasts [11]. Targeted inactivation of either of these two genes resulted in a failure to form primordial germ cells (PGCs) [53, 54]. Coculture of epiblast masses of E6.0–6.25 mouse embryos with

Chapter 17 The Role of the Sertoli Cell in Spermatogonial Stem Cell Fate

COS cells expressing BMP4 and BMP8b together caused the formation of PGCs in vitro, whereas this formation was not achieved with COS cells expressing either BMP4 or BMP8b [54]. It is apparent that the formation of PGCs requires the presence of a mixture of BMP4 and BMP8b in the form of heterodimers. Notably, increasing evidence reveals that BMP4 also is involved in the recruitment of stem cells to their niche [55] and in regulating the development of the male germ line stem cell [51, 55]. In the early postnatal mouse testis, BMP4 is secreted by Sertoli cells, while its receptor is expressed on spermatogonial stem cells [51]. The interaction of BMP4 and its receptor ALK3 leads to a signal transduction pathway that induces the expression of c-kit and allows for spermatogonial stem cell differentiation. A combination of evidence showing that c-kit is upregulated by BMP4 and other studies on the downstream molecules in the BMP signal pathway indicated that the BMP4-triggered intracellular signal network regulates the fate of spermatogonial stem cells [51, 56]. BMP4 generates a similar intracellular pathway as do other members of the TGFβ family [52]. The basic signaling engine of the TGFβ family consists of two receptor serine/threonine protein kinases (receptor II and I) and a family of proteins known as the Smads. The TGFβ ligands first bind to the type II receptor, which then recruits a type I receptor. After the formation of a ligand/type II receptor/type I receptor complex, the type II receptor phosphorylates serine and threonine residues within the intracellular GS (glycine-serine-rich) domain of the type I receptor kinase. This kinase in turn phosphorylates particular Smad members called receptor-regulated Smads (R-Smads), which include two classes: Smad2 and Smad3 for activin/TGFβ signals, and Smad1, Smad5, and Smad8 primarily for BMP signals. The phosphorylated R-Smads are released from the receptor and form a complex with the Co-Smad, Smad4. The R-Smad/Smad4 complex translocates into the nucleus and interacts with transcription factor complexes on the promoters of target genes. Additional reports, however, demonstrated that the intracellular regulation of this type of signal transduction pathway is quite complicated because of its crosstalk with other signaling pathways, which include Ca2+/calmodulin signaling, Wnt signaling, MAPK pathway, JAK-STAT pathway, and the mTOR pathway [57]. The upregulation of c-kit by BMP4 in spermatogonial stem cells is probably mediated by the Smad5/Smad4 pathway [51]. The expression of c-kit results in the formation of a group of spermatogonial cells that is more sensitive to differentiation-stimulating factors, such as stem cell factor, and the high spermatogonial proliferation/ differentiation rate may be the result of a combination of BMP4 signaling and SCF/c-kit signaling. In addition,

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the mammalian target of rapamycin (mTOR) acts as the key downstream kinase of SCF/c-kit signaling in the regulation of spermatogonial differentiation-division [58]; mTOR also is one target of the BMP4 signal pathway in the regulation of the early differentiation in the central nervous system stem cells [59]. Thus, it may be a reasonable hypothesis that SCF/c-kit signaling crosstalks with the BMP4 pathway at the mTOR kinase in the regulation of early differentiation of spermatogonia. Other receptor/ligand systems have been described in the testis that are likely important in spermatogonial stem cell interactions. Leptin is a circulating hormone produced by adipocytes that is believed to influence body weight through appetite regulation and control of reproduction, most likely through an effect on the central nervous system (Fig. 17.7). Leptin receptors are expressed in rat Leydig cells and leptin appears to have an inhibitory effect on hCG-stimulated testosterone production by adult rat Leydig cells in culture [60, 61]. Based on recent published reports on the role of leptin in stem cell proliferation [62, 63], we decided to examine its role in the testis [64]. Initially, we identified the leptin receptor in the testis, and in 5-day-old rats, it was mainly found in type A spermatogonia. We then demonstrated that leptin induced phosphorylation of STAT3 in the adult and 5-day-old testis, suggesting a role for leptin in stem cell renewal and/or differentiation. We suggested that leptin prevents spermatogonial differentiation through STAT3, thus allowing the stem cells to undergo renewal, as described in other systems [65, 66]. Several other reports have described leptin in the testis but its role in spermatogenesis remains an enigma [67, 68]. Members of the Notch gene family have been shown to play an important role in the control of cell fate, proliferation, and survival in many developmental systems [69–74] in both invertebrate and vertebrate species. It has also been demonstrated that the Notch signaling system is essential for gametogenesis in the adult germ line of C. elegans [75, 76]. Other work from several laboratories noted that some of the Notch system genes have been found to be expressed in rodent testis [77–81], but the cellular localization and the possible role of Notch in spermatogenesis have not been addressed. We hypothesized that the fate of the male germ line stem cells may be mediated through the Notch signaling pathway. We therefore sought to determine whether the components of the Notch pathway are expressed in the mouse testis [82]. Western blot analysis revealed the expression of Notch receptors (Notch 1–3), Notch ligands (Jagged 1–2 and Delta 1), and presenilin (PS1) in neonatal mouse testis. We then examined their cellular localization by immunohistochemical analysis of cocultures of spermatogonia and Sertoli cells. The three Notch receptors, Notch 1–3, were

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FIGURE 17.7 Leptin and spermatogonia. The hypothesized leptin actions on male germ cell renewal and differentiation is depicted in this schematic drawing. For spermatogonial stem cells, leptin may act through STAT3 to prevent differentiation, thus allowing the cells to undergo stem cell renewal. On the other hand, in spermatocytes, leptin directs the cells to full maturation into spermatids. Negative feedback regulators from spermatids or other germ cells may contribute to the absence of Ob-R from germ cells in stages other than IX and X of the spermatogenic cycle. (From El-Hefnawy et al. [64].)

found to be expressed in spermatogonia. Sertoli cells expressed only Notch 2 receptor. Among the Notch ligands, Delta 1 and Jagged 1 were localized exclusively in spermatogonia and Sertoli cells, respectively. Presenilin 1 was apparent in both spermatogonia and Sertoli cells. The presence of Notch receptors and Notch ligands in spermatogonia and Sertoli cells indicates that these cells are capable of both responding to and eliciting Notch signaling during the process of spermatogenesis. This research suggests that Notch signaling is required for successive cell fate determination events that occur during spermatogenesis (Fig. 17.8). An important area of future work in the mammalian testis will be investigations on Notch signaling, especially with regard to stem renewal versus stem cell differentiation.

VI. SIGNALING IN THE GERM LINE IN CAENORHABDITIS ELEGANS AND DROSOPHILA MELANOGASTER In invertebrates such as C. elegans and Drosophila, the signaling pathways that control adult germ line cell fates, that is, renewal or differentiation, have been well characterized. It was found that signals originating from

the somatic cells target their receptors on germ line stem cells to generate pathways that regulate proliferation or differentiation/meiosis. Abnormalities in these signal paths often result in either the overproliferation or the loss of germ line stem cells with a sterile phenotype. In C. elegans, the germ line stem cells interact with a somatic DTC at the end of the elongate gonad, with mitotically dividing cells at the distal zone and maturing gametes at the proximal zone. DTC produces a ligand named lag-2, and the germ line stem cell expresses glp-1, a receptor in the Notch family [83]. Loss of glp-1 signaling results in a proliferation defect and early entry of the germ cells to meiosis, whereas constitutive activity of this receptor allows germ cells to remain mitotic and leads to tumor formation [84]. The interaction of lag-2 and glp-1 generates a similar signal pathway as do other members of the Notch family. Notch is a single spanning transmembrane protein including many repeats of a protein module resembling epidermal growth factor (EGF) and three membrane-proximal Lin12/Notch/Glp-1 (LNG) repeats. The intracellular domain has four distinct regions, the RAM domain, the ankyrin repeats, a transcriptional activator domain (TAD), and the PEST (proline-, glutamate-, serine-, threonine-rich) sequence. In response to the binding of ligands, Notch receptors undergo two proteolytic cleavages. The first cleavage releases a membrane-tethered form of the Notch intracellular domain, which is then subjected to a

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FIGURE 17.8 The Notch receptor system in Sertoli cells and spermatogonia. A schematic representation of a hypothetic model for the role of the Notch system in the fate of type A spermatogonial stem cells (SPGA). The Sertoli cells produce the ligand Jagged 1, whereas the type A spermatogonia have the Notch receptor. In addition, the spermatogonia also produce the ligand Delta 1. (From Dirami et al. [82].)

second cleavage producing the soluble intracellular domain of Notch. The latter is translocated to the nucleus where it binds a transcription factor of the Su(H)/CBF1 via its RAM domain and ankyrin repeats regulating gene expression [74]. In C. elegans, glp-1 and lag-1 are homologues of Notch and Su(H), respectively. The known mechanism of the glp-1 signal pathway that promotes continued mitoses of germ line stem cells is repression of the expression of the gld-1 and gld-2 genes; the latter both promote commitment to the meiotic cell cycle [85]. In addition, a group of RNA-binding proteins, FBF-1 and FBF-2 (FBFs), also regulates stem cell fate. FBF-1 and FBF-2 sustain the mitotic cell cycle and prevent meiotic entry by downregulating the expression of gld-1 and gld-2, targeting their 3’UTR elements. Double mutants of FBF-1 and FBF-2 lead to the loss of germ line stem cells [15]. Whether FBFs are regulated by glp-1/Notch signaling remains unknown. In D. melanogaster, male germ line stem cells lie at the apical tip of the testis, surrounding a cluster of nondividing somatic cells called the hub. Each germ line stem cell is enveloped by a pair of cyst progenitor cells, the somatic stem cells, which also contact the hub [3]. The germ line stem cell and its two cyst progenitor cells divide asymmetrically. Thus, one daughter germ cell maintains contact with the hub, whereas the other displayed away from the hub becomes a gonialblast, undergoing differentiation [86, 87]. It was found that a

hub-secreted ligand, unpaired (Upd), activates the Janus kinase-signal transducers and activators of transcription (JAK-Stat) pathway within the hub-contacting daughter cell to maintain its stem cell identity [17, 18]. The inactivation of either Upd or JAK-Stat function leads to loss of maintenance of germ line stem cells, whereas ectopic expression of Upd causes unrestricted stem cell amplification. The gonialblast initiates four rounds of mitotic amplification divisions, then undergoes spermatocyte growth, meiosis, and terminal differentiation. This proliferation/differentiation transition of the germ cells is controlled by somatic cyst cells. Excess numbers of stem cells and gonialblasts in the testes carry a deficiency of either raf activity or the epidermal growth factor receptor (EGFR) in somatic cells [88, 89]. Given the fact that raf is the downstream kinase of EGFR, it is likely that the somatic cell needs the EGFR-raf-Erk pathway to be functional in promoting the proliferation/ differentiation transition in germ cells. The origination of the ligand for the EGFR is not clear yet.

VII. FATE OF GERM LINE STEM CELLS: RENEWAL VERSUS DIFFERENTIATION Stem cells can renew, differentiate, or die, and the regulation of this delicate balance in mammals has eluded biologists for many years. A clear understanding

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of this phenomenon is critical for many biological processes such as cancer, developmental biology, and cell death in the aging process. It is generally accepted that extrinsic and intrinsic factors must determine whether, on division, a stem cell will retain stem cell properties or instead give rise to differentiated cells. The extrinsic factors may come from the niche cells, the surrounding basement membrane, or from the circulation; intrinsic factors may also play an important role. Thus, what “keeps a stem a stem” is one of the most important unanswered questions in biology today and this topic was reviewed by Watt and Hogan in a recent article [90]. It was reported earlier in skin that stem cells have integrin receptors and that their loss is associated with differentiation [91]. Shinohara et al. reported that β1 and α6 integrins are markers on spermatogonial stem cells and they used this to isolate these cells for transplant [92], although Sertoli cells and other type A spermatogonia also have integrin receptors. Ohta et al. demonstrated that c-kit may be important for spermatogonial differentiation, but not for renewal [33]. Mice overexpressing GDNF showed an accumulation of undifferentiated spermatogonia, whereas GDNF-null allele mice showed a depletion of stem cell reserves [36]. LIF was first identified by its ability to maintain ES cells in an undifferentiated state [93]. It also allows primordial germ cells to proliferate and avoid apoptosis. More recent work identified this ligand in the testis [94]. Cell fate decisions are now believed to also be regulated by the Notch system [69–71], and we have identified this receptor and its ligands in the testis [21]. Leptin is believed to play a role in cell fate decisions in hematopoietic cells [62, 63] and in other systems [15] and this ligand may also be important in determining spermatogonial cell fate [25]. The regulation of spermatogonial cell fate in mammals is a topic in its infancy and a great deal of additional research is required. The transition from mitosis to meiosis has been studied extensively in C. elegans. The DTC regulates the mitotic divisions in the germ line stem cells by GLP-1/Notch signaling [15]. Decisions between mitosis and meiosis in the stem cells are controlled by a number of RNA regulatory proteins. Mitosis appears to be regulated by FBF-1 and FBF-2, whereas GLD-1, GLD-2, GLD-3, and NOS-3 promote commitment to meiosis [15, 75]. In Drosophila, it has been demonstrated that the germ line stem cell divides in an asymmetric manner, with one daughter cell remaining as a stem cell and the other daughter cell becoming a differentiated cell, the gonialblast [18]. The proximity to the somatic hub cells seems to be important for the retention of stem cell identity. In a recent series of experiments by Yamashita et al. [95], it was demonstrated that when a germ line

stem cell divides, the orientation of the mitotic spindle in the dividing stem cell determines which of the daughter cells remains a stem and which one differentiates. During interphase, the single centrosome was always located next to the hub cells, and after centrosomal duplication one centrosome remained attached to the hub and the other one migrated to the opposite pole of the nucleus. Thus, throughout mitosis the cells themselves maintain the mitotic spindle perpendicular to the hub–germ cell interface; the daughter cell that remains in contact with the hub cells remains a stem cell, whereas the cell that is further away from the hub differentiates. It was demonstrated in the germ line stem cells that this spindle orientation is established by using Apc2, a Drosophila homologue of APC (Adenomatous Polyposis Coli tumor suppressor protein), to anchor the centrosome next to the hub. Apc2 itself is localized near the hub–germ cell interface by binding germ cell Armadillio (Arm, fly β-catenin), which in turn is connected with the same molecule in the hub using DE-cadherin [95]. Immunofluorescence analysis of the region between the hub cells and the germ line stem cells revealed the presence of DE-cadherin (fly epithelial cadherin) and Armadillio. High levels of DE-cadherin and Arm were not found around the rest of the germ line stem cells.

VIII. SUMMARY AND CONCLUSIONS In the 1950s and 1960s, a great deal of research on spermatogonial stem cells was carried out by Clermont and colleagues in rodents, monkeys, and humans [20–22, 96–100]. During the same years, the Sertoli cell was recognized to be of paramount importance in germ cell regulation. Today, 40 to 50 years later, and after the publication of many thousands of papers on the topic in mammals, some of the basic questions remain unanswered. For example, what factor or cocktail of factors produced by the Sertoli cells allows for cell fate decisions in the spermatogonial stem cells? What genes and proteins are uniquely expressed in spermatogonial stem cells? What signal transduction mechanisms are active during stem cell renewal versus stem cell differentiation? Furthermore, a major stumbling block to our research on cell fate decisions in the testis is our inability to obtain long-term cultures of spermatogonial stem cells, even though Sertoli cell-produced media are added to the stem cells in culture. The recent development of a number of spermatogonial cell lines may allow for further progress in this area [101, 102]. In collaboration with Marie-Claude Hofmann (unpublished), we developed

Chapter 17 The Role of the Sertoli Cell in Spermatogonial Stem Cell Fate

another spermatogonial stem cell line that retains many characteristics of germ line stem cells such as the presence of GFR-α1 and Oct-4. Furthermore, a number of the genes present in this germ line are similar to stem cell genes found in hematopoietic stem cells. The cells also respond to Sertoli cell-produced GDNF with increased proliferation. Finally, perhaps one of the most important and potential uses of spermatogonial stem cells lies in the field of germ line genetic engineering. The idea is to change the human genome in a heritable way, that is, to fix all the known errant genes in the sperm. These would be permanent fixes to be passed from one generation to another. Eventually it should be possible to introduce corrected genes into the dividing spermatogonial stem cells, in vivo, and then allow them to differentiate into transgenic sperm. Germ cell gene therapy is one of the last major frontiers in medicine to be conquered and a thorough knowledge of the interactions between the supporting Sertoli cell and the spermatogonial stem cell will be required before this becomes a reality.

Acknowledgments Supported in part by National Institutes of Health grants HD33728 and HD36483.

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C H A P T E R

18 Sertoli Cell–Somatic Cell Interactions MICHAEL K. SKINNER Center for Reproductive Biology, Center for Integrated Biotechnology, School of Molecular Biosciences, Washington State University, Pullman, Washington

interstitial space are the Leydig cells that are responsible for testosterone production (Fig. 18.1). Other somatic cells in the interstitium are the testicular macrophages and vascular endothelial cells. The germ cell populations within the seminiferous tubule are spermatogonia, spermatocytes, spermatides, and the mature spermatozoa (Fig. 18.1). The process of germ cell development (i.e., spermatogenesis) is in large part supported by the Sertoli cells on both a nutritional and structural basis. Therefore, the regulation of Sertoli cell function is one of the most critical elements influencing spermatogenesis, testis function, and male reproduction. The critical regulatory steps within the testis that influence Sertoli cells and the process of spermatogenesis are cell–cell interactions between the various testicular cell types.

I. INTRODUCTION II. SERTOLI CELL–LEYDIG CELL INTERACTIONS III. SERTOLI CELL–PERITUBULAR CELL INTERACTIONS IV. OTHER SOMATIC CELL–SERTOLI CELL INTERACTIONS V. SUMMARY References

I. INTRODUCTION This chapter will review the current understanding of Sertoli cell–somatic cell interactions in the testis. Several other chapters are dedicated to Sertoli cell–germ cell interactions, so this topic will not be emphasized here. The literature during the past decade will also be emphasized with earlier literature being primarily cited in the form of previous reviews.

B. Cell–Cell Interactions The evolution of multicellular organisms was prompted by the ability of different cell types to interact and develop higher order integrated functions. For this reason no cells in a multicellular organism or tissue are autonomous. Therefore, the different cell types within an organ (e.g., testis) form a functional unit with a network of cellular interactions essential for tissue function and the regulation of individual cell types. The testis provides an excellent example of how a network of cell–cell interactions controls cellular functions [1]. A large number of reviews have addressed different aspects of cell–cell interactions in the testis [1–12]. The majority have focused on Sertoli cell–germ cell interactions due to the critical function of the testis

A. Cell Biology The testis is composed of a number of somatic cell types that create the morphological characteristics and endocrinology of the organ. The primary function of the testis is to support and control the process of germ cell development (i.e., spermatogenesis). The Sertoli cell is the primary somatic cell that forms the seminiferous tubule and provides the cytoarchitectural support for the developing spermatogenic cells (Fig. 18.1). The peritubular myoid cell is the mesenchymal-derived cell that surrounds the tubule and is separated from the Sertoli cell by a basement membrane. Within the SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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FIGURE 18.1

Michael K. Skinner

Cell biology of testis.

being linked to spermatogenesis [2]. Others have focused on somatic cell interactions [4, 6, 10] and specific cell types or factors [9, 12]. The primary cellular interactions in the testis to consider are between the somatic cells and germ cells (Fig. 18.2). Somatic cell–germ cell interactions are primarily mediated by the Sertoli cell. The only somatic cell type in direct contact with the spermatogenic cells is the Sertoli cell. In addition, the blood–testis barrier created by tight junctional contacts between Sertoli cells results in a serum-free microenvironment within the tubule that other cell secretory components cannot penetrate [13–15]. Therefore, Sertoli cells are the primary cell supporting the spermatogenic process and interacting with germ cells (Fig. 18.2). Sertoli cell– spermatogenic cell interactions involve structural and environmental elements to support the complex cytoarchitecture between the cells [16–18]. Nutritional and regulatory substances are also required to provide nutrient support and regulatory control of the spermatogenic cells [1–3]. Several other chapters in this book review the details of Sertoli cell–germ cell interactions (Chapters 17 and 23) so the current chapter will focus on somatic cell interactions with the Sertoli cell. The majority of cellular interactions identified have the primary function of directly or indirectly influencing spermatogenesis through the Sertoli cell.

Somatic cell interactions with Sertoli cells primarily involve the Leydig cells and peritubular myoid cells (Fig. 18.2). The interactions between both of these cell types and Sertoli cells will be reviewed. Other cell types also exist in the testis that have not been extensively investigated but are likely to have critical cellular interactions. Therefore, a network of cell–cell interactions is essential for the maintenance and control of testis function [1–12]. To help clarify the variety of cellular interactions, several categories have been developed to classify different types of cell–cell interactions (Table 18.1) [1]. This categorization separates into three classes the functionally distinct types of cell–cell interactions. The first is termed environmental interactions. These types of interactions are mediated through the extracellular environment of the cell to influence the cytoarchitecture of the cell. An environmental cell–cell interaction is influenced by an extracellular matrix or cell adhesion molecules. The primary functional effect is an influence on cell shape, contact, and tissue organization. The second type of cell–cell interaction is a nutritional interaction. These interactions involve the delivery of essential nutrients between cells via energy metabolites and vitamins. The functional effect is to support cell viability and metabolism. The final category is termed a regulatory interaction (Table 18.1). This involves the

FIGURE 18.2 Cell–cell interactions in the testis.

TABLE 18.1 Categorization of Cell–Cell Interactions Classification

Definition

Examples/mediators

Environmental

Interactions that influence the extracellular environment of the cell to affect cell contacts and cytoarchitecture

Extracellular matrix; cell adhesion molecules

Nutritional

Interactions involved in the delivery of essential nutrients between cells

Transfer of energy metabolites, metals, or vitamins

Regulatory

Agents provided by a cell that, through a signal transduction event, regulate another cell’s function on a molecular level

Paracrine/autocrine factors; growth factors; differentiation factors; cytokines

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Chapter 18 Sertoli Cell–Somatic Cell Interactions

production of a regulatory substance that is secreted and then, through a receptor-mediated signal transduction event, influences cellular functions on a molecular level. Examples of these regulatory substances are growth factors and cytokines [1]. This categorization of cell–cell interactions helps classify the different cellular interactions to be discussed as environmental, nutritional, or regulatory interactions (Table 18.1).

II. SERTOLI CELL–LEYDIG CELL INTERACTIONS The Leydig cell [19] located in the interstitium of the testis was one of the first somatic cells identified as critical for Sertoli cell functions [1]. Environmental cell–cell interactions do not exist between Leydig cells and Sertoli cells. These cells are not in direct contact with each other nor do they have an extracellular matrix in common. The inability of these cells to physically interact indicates that direct environmental interactions between Sertoli cells and Leydig cells are not possible in vivo. Nutritional cell–cell interactions also are negligible between Leydig cells and Sertoli cells. The blood–testis barrier prevents Leydig cell products from entering the seminiferous tubule. Both Sertoli cells and Leydig cells are in contact with the nutrient supply from vasculature in the interstitial space. Therefore, neither cell type is dependent on each other for nutrient supply or transport. Junctional contacts are not present between the cells to transfer metabolites either. Due to the separation and distinct cellular localization of Leydig cells and Sertoli cells (Figs. 18.1 and 18.2), only regulatory interactions occur between the cells.

Regulatory cell–cell interactions are critical between Leydig cells and Sertoli cells. The identification of androgen (i.e., testosterone) production by Leydig cells [20–26] and the ability of androgens to maintain the process of spermatogenesis [26–29] have led to observations that an essential regulatory interaction between Leydig cells and Sertoli cells is mediated by androgens. The gonadotropin leuteinizing hormone (LH) acts on Leydig cells to increase testosterone production and then acts on Sertoli cells to influence spermatogenesis [1]. This is a well-established regulatory cell–cell interaction between Leydig cells and Sertoli cells that is known to be essential for testis function. Although the androgen receptor is present in Sertoli cells, in vitro analysis has demonstrated minimal genes directly regulated by androgen. The specific mechanism of how androgen regulates Sertoli cell function remains to be elucidated. As discussed later, peritubular myoid cells also contain the androgen receptor and may help mediate androgen actions on the Sertoli cell [1]. Therefore, the regulatory interaction between Leydig cells and Sertoli cells mediated by androgens is well established, but the specific mechanism of how androgens act on Sertoli cells remains to be elucidated. In addition to the steroidal substances produced by Leydig cells, a number of protein factors also are produced and can regulate Sertoli cell function in vitro [1]. Factors shown to be produced by Leydig cells that can potentially influence Sertoli cells include renin [30, 31], prodymorphin [32], oxytocin [33], pro-opiomelanocortin (POMC) peptides (β endorphin, αMSH, ACTH) [34–37], and growth hormone releasing hormone (GHRH) [38] (Table 18.2). The majority of these peptide factors have been shown to be produced by Leydig cells and have negligible effects on Sertoli cells in comparison to

TABLE 18.2 Sertoli Cell–Leydig Cell Regulatory Interactions Potential paracrine factor

Site production

Site action

Actions/proposed function

Androgen

Leydig

Sertoli

Regulate/maintain function and differentiation

POMC peptides β endorphin, MSH, ACTH

Leydig

Sertoli

Decrease FSH actions; increase FSH actions

GNRH-like factor

Sertoli

Leydig

Decrease steroidogenesis

Estrogen

Sertoli

Leydig

Decrease steroidogenesis

IGF-1

Sertoli

Leydig

Increase steroidogenesis

TGFα

Sertoli

Leydig

Decrease steroidogenesis; increase growth

TGFβ

Sertoli

Leydig

Increase steroidogenesis

IL-1

Sertoli

Leydig

Decrease steroidogenesis

Inhibin

Sertoli

Leydig

Increase steroidogenesis

MIS

Sertoli

Leydig

Cell proliferation

SCF

Sertoli

Leydig

Cell proliferation

Dhh

Sertoli

Leydig

Cell proliferation

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Michael K. Skinner

follicle-stimulating-hormone (FSH) actions. The in vivo function for these factors remains to be elucidated. A number of studies have demonstrated that Sertoli cells appear to directly regulate Leydig cell function [1]. One approach that has been used is to selectively destroy Leydig cells and determine how Sertoli cells may influence their regeneration [39]. More direct studies have shown that Sertoli cell–secreted products can regulate Leydig cell steroidogenesis and function [40–42]. Recently, using patients with acquired hypogonadotropic hypogonadism and recombinant LH and FSH, observations support a role for local Sertoli cell–derived paracrine factors to influence Leydig cells [43]. Another recent in vivo experiment used the FSH receptor knockout mouse model and demonstrated that Leydig cell hormone responsiveness and function are impaired in the absence of normal Sertoli cells [44]. These two in vivo experiments provide further support that Sertoli cell products directly feed back and regulate Leydig cell function. The factors that mediate this response remain to be fully elucidated. A number of regulatory factors have been shown to be produced by Sertoli cells that can act on Leydig cells and influence cellular function (Table 18.2) [1]. Estrogen produced by Sertoli cells has been shown to suppress androgen production by Leydig cells in vivo [45, 46], but the physiological relevance of this needs to be determined. In addition, mature adult animals have negligible estrogen production by Sertoli cells. Protein regulatory factors include gonadotropin releasing hormone (GnRH) [47–49], which has been shown to be expressed by and can act on Leydig cells in vitro. Insulin growth factor-1 (IGF-1) is produced by both cell types and can influence the functions of both cell types [50], but the high circulating levels of IGF-1 lead us to question the role of local production. Transforming growth factors alpha (TGFα) and beta (TGFβ) are also produced by Sertoli cells and can influence Leydig cells [1, 9, 51]. However, Leydig cell production of TGFβ isoforms and other cells in the interstitium questions the specificity of Sertoli cell–Leydig cell TGFα or TGFβ mediated interactions. The interleukins have also been suggested as paracrine factors, but the specifics of cellular localization and action again question the specificity of such an interaction. Inhibin production by Sertoli cells has also been shown to influence Leydig cell function as a paracrine factor [1, 52–54]. Several recently identified protein factors shown to be produced by Sertoli cells that potentially affect Leydig cells have expanded the list in Table 18.2. Stem cell factor/kit ligand has been shown to be produced by Sertoli cells in vitro and can influence Leydig cell function [55]. Müllerian-inhibiting substance (MIS) is also produced by Sertoli cells, and

observations in MIS knockout animals demonstrate that MIS can influence Leydig cell proliferation [56]. In contrast, MIS-overexpressing animals have Leydig cell hyperplasia, supporting a role for MIS mediated Sertoli cell–Leydig cell interactions. This is one of the few Sertoli cell factors shown in vivo to influence Leydig cells. Another interesting factor is desert hedgehog (Dhh), which has been shown to be expressed by Sertoli cells and acts on Leydig cells [57]. Whether this is only an early embryonic development phenotype or also an adult cell–cell interaction remains to be assessed. Additional protein factors have been postulated to mediate Sertoli cell–Leydig cell interactions, but require further investigation [58–60]. The majority of these factors have been shown to be expressed and act on Leydig cells in vitro, however, the physiological significance of the factors in vivo remains to be elucidated.

III. SERTOLI CELL–PERITUBULAR CELL INTERACTIONS The peritubular myoid cell is the mesenchymal/ stromal cell population that surrounds the seminiferous tubules and is adjacent to the basal surface of the Sertoli cell (Fig. 18.1) [1, 12, 61–64]. The peritubular cells and Sertoli cells are separated by a basement membrane and the two cell types make up the somatic elements of the seminiferous tubule. Peritubular cells have been shown to exist in all mammalian species examined [1, 12], with some differences in the number of layers of peritubular cells. Early in embryonic development at the time of testis determination, the peritubular cells are derived from mesenchymal cells within the mesonephros adjacent to the developing gonad. Mesonephros cells migrate into the gonad and promote cord formation, with Sertoli and primordial germ cell aggregates, to become the outer layer of cells and precursor peritubular cells. The developing peritubular cells appear to provide structural integrity to the developing testis cords and have a regulatory role in testis development. At the onset of puberty, androgen promotes the formation of peritubular myoid cells to develop smooth muscle characteristics [65, 66]. In the adult the peritubular cells have smooth muscle characteristics and are involved in contracting the seminiferous tubules and promoting movement of spermatozoa within the lumen of the tubules into the rat testis [1, 12, 66–68]. The contractility of the peritubular cells has been observed and postulated to be involved in spermatogenic cell movement within the tubule. The contractility of the peritubular cells appears to be developmentally regulated and stage specific [68].

Chapter 18 Sertoli Cell–Somatic Cell Interactions

Regulatory factors described later appear to influence this process. The integral association of Sertoli cells and peritubular myoid cells has promoted the evolution of critical cell–cell interactions between the cell populations. This is the classic example of a mesenchymal cell–epithelial cell interaction found to be important for the development of all tissues examined [69, 70]. The cellular interactions between peritubular cells and Sertoli cells are mediated by a variety of secretory products including extracellular matrix components and growth factors [1, 12]. Environmental cell–cell interactions are critical between peritubular cells and Sertoli cells [1, 12]. This environmental interaction is mediated by the extracellular matrix (i.e., basement membrane) separating the two cell populations. This basement membrane contributes to the structural integrity of the tubule and acts as a partial component (i.e., prefilter) to help develop the blood–testis barrier [71]. The extracellular matrix is produced cooperatively by both peritubular cells and Sertoli cells [1, 12, 72]. Each cell population produces different components of the extracellular matrix (Table 18.3). Sertoli cells have been shown to produce laminin, collagen I, collagen IV, and chondroitin proteoglycans [1, 12]. Peritubular cells produced fibronectin, collagen I, and chondroitin proteoglycans [1, 12]. The coculture of peritubular cells and Sertoli cells has been shown to influence the expression of these extracellular matrix components by the other cell type, suggesting a regulatory interaction between the cells in the control of extracellular matrix production [73–75]. The regulation of extracellular matrix degradation is also critical in the dynamic interactions between peritubular cells and Sertoli cells via the basement membrane. The expression of collagenases and glycosidases by the two cell populations [76, 77] suggests that interactions between the cell types is required [1, 12]. Abnormal expression and/or degradation of the extracellular matrix may be a factor in some forms of human male infertility [78]. In addition to the classic

TABLE 18.3 Extracellular Matrix Components Produced Sertoli cell

Peritubular cell

Laminin

Fibronectin

Collagen I

Collagen I

Collagen IV

Chondroitin proteoglycans

Chondroitin proteoglycan

Cell chondroitin proteoglycan

Heparin/chondroitin proteoglycan Cellular heparin proteoglycan

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extracellular matrix components, other extracellular factors such as cell adhesion molecules (e.g., cadherins) also are likely involved in environmental peritubular and Sertoli cell interactions [79]. Regulatory cell–cell interactions between peritubular cells and Sertoli cells are critical in the regulation of Sertoli cell function [1, 12]. Initial observations utilized cocultures of peritubular cells and Sertoli cells to demonstrate the ability of peritubular cells to influence Sertoli cell functions [80–83]. This influence has been confirmed more recently by demonstrating that the presence of peritubular cells in cocultures influences Sertoli glycosaminoglycan synthesis [84] and androgen binding protein (ABP) production [85]. Cocultures of the cells also influence cell proliferation and hormone responsiveness of the cells [86]. These observations have been extended by using peritubular cell conditioned medium to influence Sertoli cell functions in vitro [1, 12]. Serum-free conditioned medium from peritubular cells was found to stimulate the expression of a number of Sertoli cell gene products [87–89]. Subsequently a peritubular cell line was generated that produced conditioned medium with similar activity [90] and the same investigators found that stromal cell lines from several tissues also produced conditioned medium that can stimulate Sertoli cells [91]. The bioactive component within the peritubular cell conditioned medium was isolated and termed PModS [92, 93]. The PModS activity was found to influence a number of Sertoli cell functions [92–97] and utilized signal transduction pathways distinct from other hormones (e.g., FSH) known to influence Sertoli cells [98, 99]. The PModS activity has been isolated [93] but has not been fully characterized. Rigorous biochemical separation of individual peptides alters bioactivity, suggesting that a complex of multiple proteins is required for full bioactivity and this has complicated purification and cloning procedures. Verhoeven et al. [100] have shown that the PModS bioactivity cannot be attributed to other paracrine factors such as IGF-1, bFGF, EGF, TGFβ, NGF, PDGF heregulins, or neu differentiation factors [91, 100, 101]. PModS appears to be unique and remains to be characterized on a molecular level. The bioactivity associated with PModS suggests that it is a major regulatory factor stimulating Sertoli cell differentiation and mediating peritubular cell–Sertoli cell interactions. Peritubular cells contain high levels of androgen receptor [102–104] and their development is dependent on androgen [65, 66]. Peritubular cells respond to androgens in vitro and can modulate Sertoli cell functions [87, 105–107]. The production of PModS appears to be stimulated by androgens and in part mediates

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androgen effects on Sertoli cells via the peritubular cells [1, 12, 108]. Further analysis of androgen mediated peritubular cell–Sertoli cell interactions is needed to understand how androgens control Sertoli cell function and spermatogenesis. A number of other potential paracrine factors have been shown to be produced by peritubular cells that can influence Sertoli cell function [1, 12]. Peritubular cells express the epidermal growth factor (EGF) family member TGFα [109, 110] and Sertoli cells do have low levels of the EGF receptor [110, 111]. TGFα does not have major effects on Sertoli cell functions [109, 110], but does influence specific functions such as lactate production [112]. Sertoli cells also produce TGFα [110] and the peritubular cells are stimulated to proliferate in response to TGFα/EGF. Therefore, the importance and specificity of TGFα mediated peritubular cell–Sertoli cell interactions remains to be determined. Peritubular cells also produce TGFβ [113–116], which can inhibit the actions of growth factors such as TGFα. Sertoli cells also produce TGFβ isoforms [115–117] such that the specificity of the cell–cell interaction is difficult to assess. Sertoli cells do not respond to TGFβ with any major functional changes measured, but peritubular cell proliferation is inhibited by TGFβ. Another growth factor produced by both cell types that can act on both cell types is IGF-1 [118, 119]. The degree to which IGF-1 acts as a paracrine factor between the cells is unclear. Recently, peritubular cells have been shown to express activin, which can act on Sertoli cells to influence prepubertal cell proliferation [120]. This is a novel site for the expression of activin and it was found that Sertoli cells produced negligible levels. Therefore, activin may be a paracrine factor mediating peritubular cell–Sertoli cell interactions. Peritubular cells also produce heregulins or neu differentiation factors (NDF), NDFα, and NDFβ, which can act through receptors on Sertoli cells [101]. Specific actions on Sertoli cells involve slight stimulation in the expression of gene products such as transferrin, but not as dramatic as FSH or PModS [101]. Peritubular cells recently have been shown to produce leukemia inhibitory factor (LIF) that can act on early-stage spermatogenic cells (e.g., spermatogonia) and potentially on Sertoli cells [121]. Specific actions on Sertoli cells remain to be examined [121]. All of the factors just described are summarized in Table 18.4. They have the potential to be produced by peritubular cells and to modulate Sertoli cell functions. However, many do not have major effects on Sertoli cells, but yet to be identified specific functions may be the primary targets of action. The physiological significance of these factors in vivo remains to be determined.

TABLE 18.4 Major Sertoli Cell and Peritubular Cell Paracrine Regulatory Products Potential paracrine factor

Site production

Action / proposed function

Site action

PModS

Peritubular

Sertoli

Paracrine regulatory agent

TGFα

Peritubular

Sertoli

Growth stimulation/ EGF-like

TGFβ

Both

Both

Growth inhibition

IGF-I

Both

Both

Maintenance cell growth/ differentiation

NDFα/NDFβ

Peritubular

Sertoli

Increase differentiation

LIF

Peritubular

Gonia/ Sertoli

Growth stimulation

Activin

Peritubular

Sertoli

Growth stimulation

NT3

Sertoli

Peritubular

Embryonic chemotactic factor

bFGF

Sertoli

Peritubular

Growth stimulation

The ability of Sertoli cells to regulate peritubular cells was postulated from the observation that androgens alone could not promote peritubular cell differentiation, but gonadotropins acting indirectly through Sertoli cells were also required [67]. Several paracrine factors discussed are produced by both Sertoli cells and peritubular cells, which can act on peritubular cells including TGFα, TGFβ, and IGF-1 [110, 115–120]. The degree to which these factors act as paracrine mediators of Sertoli cell–peritubular cell interactions versus autocrine actions on each other remains to be determined. During early embryonic development, upon male sex determination, the Sertoli cells produce neurotrophin 3 (NT-3), which can act as a chemotactic factor and promote migration of the precursor peritubular cells into the testis and promote cord formation for testis sex differentiation [122]. This is perhaps the first Sertoli cell–peritubular cell interaction during development [122], but this NT-3 paracrine interaction does not appear to be critical in the adult testis. Other factors produced by Sertoli cells that have the capacity to influence peritubular cells include basic fibroblast growth factor (bFGF), which is produced by Sertoli cells in response to FSH and can promote the proliferation of peritubular cells [123]. As shown in Table 18.4, several factors may act as potential paracrine factors to mediate Sertoli cell–peritubular cell interactions that require further investigation to assess the importance of these cell–cell interactions.

Chapter 18 Sertoli Cell–Somatic Cell Interactions

Upon differentiation at the onset of puberty by androgens and gonadotropin actions on Sertoli cells, peritubular cells develop smooth muscle characteristics to become peritubular myoid cells [66, 67]. These cells have the ability to contract and move spermatozoa through the seminiferous tubules to the rete [67, 68]. Factors that may be involved in the contraction of peritubular cells have been identified. The peritubular cells have receptors and can respond to endothelins (ET) [124–126], vasopressin [127] and platelet-derived growth factor beta (PDGFβ) [128] to induce contractility of the seminiferous tubules. These factors are likely important for testis function, but the site of production of these factors remains to be elucidated. Sertoli cells have not been shown to be the site for production of these factors. As discussed later, other testicular cells in the interstitium may be involved (e.g., endothelial cells), but this remains to be investigated.

IV. OTHER SOMATIC CELL–SERTOLI CELL INTERACTIONS Other testicular somatic cells that may interact with Sertoli cells are primarily located within the interstitium. A cell population that makes up approximately 20% of the interstitial cell population is the testicular macrophage [129]. These testicular macrophages appear to be unique and have been shown to interact with Leydig cells to influence steroidogenesis and Leydig cell function [129–132]. Factors mediating the macrophage–Leydig cell interaction include steroids [132] and tumor necrosis factor (TNF) [129, 131]. No major factors produced by macrophages have been shown to influence Sertoli cell functions. However, macrophage migration inhibitory factor (MIF) was found to influence peritubular cell calcium mobilization [133]. The ability of MIF to act on peritubular cells and indirectly affect Sertoli cells is a possibility that needs to be examined. Another cell type in the interstitium is lymphocytes, which may have a role in the immunology of the testis [134], but factors that specifically influence Sertoli cell function have not been identified. Vascular and lymphatic endothelial cells are also relatively abundant in the interstitium of the testis [135]. As previously reviewed [1], the vasculature associates with the outer wall of the seminiferous tubules associated with peritubular cells and the lymphatic endothelial cells can envelop Leydig cells. Specific products of these endothelin cell populations that affect Sertoli cells have not been identified. However, it is anticipated that endothelial cell products such as endothelials may

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influence peritubular cells. Sertoli cell products that could influence endothelial cells such as bFGF [123] remain to be investigated. It is likely these other testicular cell populations have a role in testis cell–cell interactions, but such a role remains to be elucidated.

V. SUMMARY In considering the categories of cell–cell interactions, the only major environmental somatic cell–Sertoli cell interactions are between peritubular cells and Sertoli cells. This environmental interaction is mediated by a basement membrane between the cells that is produced cooperatively by both cell types. This environmental interaction is critical for maintaining the structural integrity of the seminiferous tubules and polarized epithelial morphology of the Sertoli cells. Other somatic cells in the testis (e.g., Leydig cell) are not in direct contact with Sertoli cells and, hence, there are no environmental interactions between the cells. Because the blood–testis barrier encompasses the developing spermatogenic cells and Sertoli cells, no major nutritional interactions are required between the somatic cells. Nutritional interactions are essential between Sertoli cells and germ cells, but because the basal surface of the Sertoli cells and the other testicular somatic cells are in contact with serum derived nutrients, no major somatic cell–Sertoli cell nutritional interactions are required. Regulatory cell–cell interactions between somatic cells and Sertoli cells are prevalent and required for normal testis function. These regulatory interactions are generally mediated by secreted factors that, through receptor mediated events, influence cellular function on a molecular level. The number of regulatory agents identified to potentially mediate somatic cell–Sertoli cell interactions has increased during the past decade [1, 12] and are summarized in Tables 18.2 and 18.4. The focus of the current review was on peritubular cell–Sertoli cell interactions and Leydig cell–Sertoli cell interactions. The majority of research has been on these somatic cell interactions. Although other somatic cells (e.g., macrophages and endothelial cells) are present in the interstitium, specific interactions with Sertoli cells have not been rigorously investigated. Further characterization of the network of regulatory cell–cell interactions is required to understand the development and control of testis function. The list of potential paracrine regulatory factors has increased as shown in Tables 18.2 and 18.4. All of these factors have been shown to be produced by a specific testicular cell type, to have receptors, and to act potentially on another cell type. Some of the factors do have

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major actions on the target cell to influence cell growth or differentiation. Others have negligible effects on the target cell considering the specific functions investigated. The importance of these specific regulatory factors remains to be determined and needs to be the primary focus of further analysis of the factor or cell–cell interaction. Relatively few factors have been shown to be critical on a physiological level. Examples are the production of MIS and inhibin by Sertoli cells and actions on adult Leydig cells. Through transgenic and knockout mouse models, the importance of these specific cell–cell interactions and paracrine factors has been established. Future analysis of specific regulatory factors needs to be extended to assess the physiological role of the cell–cell interaction of interest. The classic endocrine concept and the control of tissue function need to be extended to effects on local cell–cell interactions. We now know that the ability of an endocrine agent to act on a specific cell type promotes a cascade of local cell–cell interactions. Understanding the local network of cell–cell interactions and the ability of endocrine agents to modulate them is essential to elucidate the endocrinology of a tissue. The two primary endocrine agents to influence testis function are the gonadotropins LH and FSH (Fig. 18.3). LH acts on Leydig cells to promote the production of androgen, which then acts on Sertoli cells and peritubular cells. The degree to which direct versus peritubular cell– mediated androgen actions have on Sertoli cell function is currently unclear. Very few genes have been shown to be directly responsive to androgens in Sertoli cells, such that direct molecular actions of androgens in Sertoli cells are uncertain at this time. Peritubular cell secreting products (e.g., PModS) are androgen responsive and have dramatic effects on the Sertoli cell, but the molecular characteristics of this factor remain to be elucidated. Therefore, the actions of androgens in the testis likely involve both Leydig cell–Sertoli cell and peritubular cell–Sertoli cell interactions,

but the specific action of androgens requires further investigations. FSH acts on Sertoli cells and also likely promotes cell–cell interactions with other cells in the testis such as Leydig cells (Fig. 18.3). Although the ability of Sertoli cells to respond to FSH and influence Leydig cell function (e.g., steroidogenesis) is well established, the specific factors that modulate this interaction require further investigation. Elucidation of the cell–cell interactions in the testis and the ability of the endocrine system (e.g., gonadotropins) to regulate these interactions will provide the next phase of our understanding of male reproductive endocrinology and the regulation of testis biology. As previously discussed the majority of research on cell–cell interactions in the testis has been focused on Sertoli cell–germ cell interactions. Due to the complex architectural support and unique microenvironment within the seminiferous tubule, the spermatogenic cells are dependent on normal and optimal Sertoli cell function. In the mammalian testis, the existence of the blood–testis barrier and germ cell syncytium indicates the complexity and integrated dependence the germ cells have on the Sertoli cell. Any factor that regulates or influences Sertoli cell function indirectly controls spermatogenesis and male fertility. Therefore, local somatic cell interactions within the testis have a critical role in the regulation of testis function and male reproduction. Both Leydig cell and peritubular cell interactions with Sertoli cells are now known to be essential for Sertoli cell development and function. The complex network of testis cell–cell interactions is starting to be elucidated, but significant future research is needed to identify and characterize the specific paracrine factors and the actions of these factors on the various cell types. This will be a fruitful area of research for the future and assist in the identification of pharmaceutical agents to treat testis pathophysiologies such as male infertility and develop male contraceptive agents.

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FIGURE 18.3 Endocrine cellular interactions.

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44. Krishnamurthy, H., Kats, R., Danilovich, N., Javeshghani, D., and Sairam, M. R. (2001). Intercellular communication between Sertoli cells and Leydig cells in the absence of follicle-stimulating hormone-receptor signaling. Biol. Reprod. 65(4), 1201–1207. 45. Dorrington, J. H., and Armstrong, D. T. (1975). Follicle-stimulating hormone stimulates estradiol-17beta synthesis in cultured Sertoli cells. Proc. Natl. Acad. Sci. USA 72(7), 2677–2681. 46. Rommerts, F. F., and Brinkman, A. O. (1981). Modulation of steroidogenic activities in testis Leydig cells. Mol. Cell Endocrinol. 21(1), 15–28. 47. Sharpe, R. M., and Fraser, H. M. (1980). Leydig cell receptors for luteinizing hormone releasing hormone and its agonists and their modulation by administration or deprivation of the releasing hormone. Biochem. Biophys. Res. Commun. 95(1), 256–262. 48. Sharpe, R. M., and Cooper, I. (1987). Comparison of the effects on purified Leydig cells of four hormones (oxytocin, vasopressin, opiates and LHRH) with suggested paracrine roles in the testis. J. Endocrinol. 113(1), 89–96. 49. Clayton, R. N., Katikineni, M., Chan, V., Dufau, M. L., and Catt, K. J. (1980). Direct inhibition of testicular function by gonadotropin-releasing hormone: Mediation by specific gonadotropin-releasing hormone receptors in interstitial cells. Proc. Natl. Acad. Sci. USA 77(8), 4459–4463. 50. Spiteri-Grech, J., and Nieschlag, E. (1992). The role of growth hormone and insulin-like growth factor I in the regulation of male reproductive function. Horm. Res. 38(Suppl 1), 22–27. 51. Gautier, C., Levacher, C., Saez, J. M., and Habert, R. (1997). Expression and regulation of transforming growth factor beta1 mRNA and protein in rat fetal testis in vitro. Biochem. Biophys. Res. Commun. 236(1), 135–139. 52. Hsueh, A. J., Dahl, K. D., Vaughan, J., Tucker, E., Rivier, J., Bardin, C. W., and Vale, W. (1987). Heterodimers and homodimers of inhibin subunits have different paracrine action in the modulation of luteinizing hormone-stimulated androgen biosynthesis. Proc. Natl. Acad. Sci. USA 84(14), 5082–5086. 53. Lin, T., Calkins, J. K., Morris, P. L., Vale, W., and Bardin, C. W. (1989). Regulation of Leydig cell function in primary culture by inhibin and activin. Endocrinology 125(4), 2134–2140. 54. Drummond, A. E., Risbridger, G. P., and de Kretser, D. M. (1989). The involvement of Leydig cells in the regulation of inhibin secretion by the testis. Endocrinology 125(1), 510–515. 55. Munsie, M., Schlatt, S., deKretser, D. M., and Loveland, K. L. (1997). Expression of stem cell factor in the postnatal rat testis. Mol. Reprod. Dev. 47(1), 19–25. 56. Behringer, R. R. (1994). The in vivo roles of mullerian-inhibiting substance. Curr. Top. Dev. Biol. 29, 171–187. 57. Bitgood, M. J., Shen, L., and McMahon, A. P. (1996). Sertoli cell signaling by Desert hedgehog regulates the male germline. Curr. Biol. 6(3), 298–304. 58. Loveland, K. L., Hedger, M. P., Risbridger, G., Herszfeld, D., and De Kretser, D. M. (1993). Identification of receptor tyrosine kinases in the rat testis. Mol. Reprod. Dev. 36(4), 440–447. 59. Wollina, U., Schreiber, G., Gornig, M., Feldrappe, S., Burchert, M., and Gabius, H. J. (1999). Sertoli cell expression of galectin-1 and -3 and accessible binding sites in normal human testis and Sertoli cell only-syndrome. Histol. Histopathol. 14(3), 779–784. 60. Habasque, C., Satie, A. P., Aubry, F., Jegou, B., and Samson, M. (2003). Expression of fractalkine in the rat testis: Molecular cloning of a novel alternative transcript of its gene that is differentially regulated by pro-inflammatory cytokines. Mol. Hum. Reprod. 9(8), 449–455. 61. Clermont, Y., and Percy, B. (1957). Quantitative study of the cell population of the seminiferous tubules in immature rats. Am. J. Anat. 100, 241–260.

62. Lacy, D., and Rotblat, J. (1960). Study of normal and irradiated boundary tissue of the seminiferous tubules of the rat. Exp. Cell Res. 21, 49–70. 63. Leeson, C. R., and Leeson, T. S. (1963). The postnatal development and differentiation of the boundary tissue of the seminiferous tubule of the rat. Anat. Rec. 147, 243–260. 64. Ross, M. (1967). The fine structure and development of the peritubular contractile cell component in the seminiferous tubules of the moist. Am. J. Anat. 121, 523–528. 65. Bressler, R. S., and Ross, M. H. (1972). Differentiation of peritubular myoid cells of the testis: Effects of intratesticular implantation of newborn mouse testes into normal and hypophysectomized adults. Biol. Reprod. 6(1), 148–159. 66. Hovatta, O. (1972). Effect of androgens and antiandrogens on the development of the myoid cells of the rat seminiferous tubules. Z. Zellforsch. Mikrosk. Anat. 131(3), 299–308. 67. Ross, M. H., and Long, I. R. (1966). Contractile cells in human seminiferous tubules. Science 153(741), 1271–1273. 68. Suvanto, O., and Kormano, M. (1970). The relation between in vitro contractions of the rat seminiferous tubules and the cyclic stage of the seminiferous epithelium. J. Reprod. Fertil. 21(2), 227–232. 69. Grobstein, C. (1967). Mechanisms of organogenetic tissue interaction. Natl. Cancer Inst. Monogr. 26, 279–299. 70. Cunha, G. R., Chung, L. W., Shannon, J. M., Taguchi, O., and Fujii, H. (1983). Hormone-induced morphogenesis and growth: Role of mesenchymal–epithelial interactions. Recent Prog. Horm. Res. 39, 559–598. 71. Dym, M., and Fawcett, D. W. (1970). The blood–testis barrier in the rat and the physiological compartmentation of the seminiferous epithelium. Biol. Reprod. 3(3), 308–326. 72. Skinner, M. K., Tung, P. S., and Fritz, I. B. (1985). Cooperativity between Sertoli cells and testicular peritubular cells in the production and deposition of extracellular matrix components. J. Cell Biol. 100(6), 1941–1947. 73. Raychoudhury, S. S., Blackshaw, A. W., and Irving, M. G. (1993). Rat Sertoli cell extracellular matrix regulates glycosaminoglycan synthesis by peritubular myoid cells in vitro. Mol. Reprod. Dev. 35, 151–158. 74. Raychoudhury, S. S., Blackshaw, A. W., and Irving, M. G. (1993). Hormonal modulation of the interactions of cultured rat testicular Sertoli and peritubular myoid cells. Effects on glycosaminoglycan synthesis. J. Androl. 14, 9–16. 75. Raychoudhury, S. S., Irving, M. G., Thompson, E. W., and Blackshaw, A. W. (1992). Collagen biosynthesis in cultured rat testicular Sertoli and peritubular myoid cells. Life Sci. 51(20), 1585–1596. 76. Ailenberg, M., Stetler-Stevenson, W. G., and Fritz, I. B. (1991). Secretion of latent type IV procollagenase and active type IV collagenase by testicular cells in culture. Biochem. J. 279 (Part 1), 75–80. 77. Sang, Q. X., Stetler-Stevenson, W. G., Liotta, L. A., and Byers, S. W. (1990). Identification of type IV collagenase in rat testicular cell culture: Influence of peritubular–Sertoli cell interactions. Biol. Reprod. 43(6), 956–964. 78. Gulkesen, K. H., Erdogru, T., Sargin, C. F., and Karpuzoglu, G. (2002). Expression of extracellular matrix proteins and vimentin in testes of azoospermic man: An immunohistochemical and morphometric study. Asian J. Androl. 4(1), 55–60. 79. Johnson, K. J., and Boekelheide, K. (2002). Dynamic testicular adhesion junctions are immunologically unique. II. Localization of classic cadherins in rat testis. Biol. Reprod. 66(4), 992–1000. 80. Tung, P. S., and Fritz, I. B. (1980). Interactions of Sertoli cells with myoid cells in vitro. Biol. Reprod. 23(1), 207–217.

Chapter 18 Sertoli Cell–Somatic Cell Interactions 81. Hutson, J. C., and Stocco, D. M. (1981). Peritubular cell influence on the efficiency of androgen-binding protein secretion by Sertoli cells in culture. Endocrinology 108(4), 1362–1368. 82. Holmes, S. D., Lipshultz, L. I., and Smith, R. G. (1984). Regulation of transferrin secretion by human Sertoli cells cultured in the presence or absence of human peritubular cells. J. Clin. Endocrinol. Metab. 59(6), 1058–1062. 83. Ueda, H., Tres, L. L., and Kierszenbaum, A. L. (1988). Culture patterns and sorting of rat Sertoli cell secretory proteins. J. Cell Sci. 89(Part 2), 175–188. 84. Raychoudhury, S. S., Blackshaw, A. W., and Irving, M. G. (1993). Hormonal modulation of the interactions of cultured rat testicular Sertoli and peritubular myoid cells. Effects on glycosaminoglycan synthesis. J. Androl. 14(1), 9–16. 85. Thompson, E. W., Blackshaw, A. W., and Raychoudhury, S. S. (1995). Secreted products and extracellular matrix from testicular peritubular myoid cells influence androgen-binding protein secretion by Sertoli cells in culture. J. Androl. 16(1), 28–35. 86. Schlatt, S., de Kretser, D. M., and Loveland, K. L. (1996). Discriminative analysis of rat Sertoli and peritubular cells and their proliferation in vitro: Evidence for follicle-stimulating hormone-mediated contact inhibition of Sertoli cell mitosis. Biol. Reprod. 55(2), 227–235. 87. Skinner, M. K., and Fritz, I. B. (1985). Structural characterization of proteoglycans produced by testicular peritubular cells and Sertoli cells. J. Biol. Chem. 260(21), 11874–11883. 88. Skinner, M. K., and Fritz, I. B. (1986). Identification of a nonmitogenic paracrine factor involved in mesenchymal–epithelial cell interactions between testicular peritubular cells and Sertoli cells. Mol. Cell Endocrinol. 44(1), 85–97. 89. Hutson, J. C., Yee, J. B., and Yee, J. A. (1987). Peritubular cells influence Sertoli cells at the level of translation. Mol. Cell Endocrinol. 52(1–2), 11–15. 90. Hoeben, E., Briers, T., Vanderstichele, H., De Smet, W., Heyns, W., Deboel, L., Vanderhoydonck, F., and Verhoeven, G. (1995). Characterization of newly established testicular peritubular and prostatic stromal cell lines: Potential use in the study of mesenchymal–epithelial interactions. Endocrinology 136(7), 2862–2873. 91. Hoeben, E., Deboel, L., Rombauts, L., Heyns, W., and Verhoeven, G. (1994). Different cells and cell lines produce factors that modulate Sertoli cell function. Mol. Cell Endocrinol. 101(1–2), 263–275. 92. Skinner, M. K., and Fritz, I. B. (1985). Testicular peritubular cells secrete a protein under androgen control that modulates Sertoli cell functions. Proc. Natl. Acad. Sci. USA 82(1), 114–118. 93. Skinner, M. K., Fetterolf, P. M., and Anthony, C. T. (1988). Purification of a paracrine factor, P-Mod-S, produced by testicular peritubular cells that modulates Sertoli cell function. J. Biol. Chem. 263(6), 2884–2890. 94. Norton, J. N., and Skinner, M. K. (1989). Regulation of Sertoli cell function and differentiation through the actions of a testicular paracrine factor P-Mod-S. Endocrinology 124(6), 2711–2719. 95. Rosselli, M., and Skinner, M. K. (1992). Developmental regulation of Sertoli cell aromatase activity and plasminogen activator production by hormones, retinoids and the testicular paracrine factor, PModS. Biol. Reprod. 46(4), 586–594. 96. Anthony, C. T., Rosselli, M. and Skinner, M. K. (1991). Actions of the testicular paracrine factor (P-Mod-S) on Sertoli cell transferrin secretion throughout pubertal development. Endocrinology 129(1), 353–360. 97. Mullaney, B. P., Rosselli, M., and Skinner, M. K. (1994). Developmental regulation of Sertoli cell lactate production by

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C H A P T E R

19 Sertoli Cell Lines KENNETH P. ROBERTS Department of Urologic Surgery, University of Minnesota, Minneapolis, Minnesota

cell lines were developed in the early 1980s and include the TM4 cell line, which will be discussed at length in this chapter. Several cell lines were established and reported on in the 1990s, only a few of which are well studied. In this chapter, the various Sertoli cell lines created will be reviewed with their published strengths and weaknesses. The utility of these cell lines, as well as their limitations, should become clear as a result of this discussion.

I. INTRODUCTION II. SOURCES OF SERTOLI CELLS AND METHODS OF IMMORTALIZATION III. ESTABLISHED SERTOLI CELL LINES IV. SUMMARY References

I. INTRODUCTION Effective cell lines can greatly facilitate research on Sertoli cell function by providing a readily available supply of cells with consistent and predictable properties. An ideal Sertoli cell line would facilitate the study of germ cell–Sertoli cell interaction, gene regulation, protein secretion and processing (glycosylation, proteolytic cleavage, etc.), protein production and isolation, and germ cell development including stem cell propagation. Unfortunately, no one cell line exists that is perfectly suited to all of these types of investigations. In reality, some loss of specialization always occurs when terminally differentiated, nondividing cells are immortalized. Generally, terminally differentiated cells, such as Sertoli cells, cease dividing as they come to full differentiation. When such cells are induced to divide, a necessary state for cell lines, some of the differentiated function of the cell is invariably lost. Fortunately, a cell line can be useful without retaining every feature of specialized function of the cell type from which it is derived. The challenge is to create a cell line that retains the features of the parent cell type that are required to perform the desired study. To date, many different Sertoli cell lines have been created and used in various studies. The first Sertoli SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

II. SOURCES OF SERTOLI CELLS AND METHODS OF IMMORTALIZATION With the exception of a series of lines from sheep, the SM lines, all other Sertoli cell lines developed have been derived from the mouse or rat. This is somewhat representative of the emphasis in Sertoli cell biology, in which the bulk of the research has been carried out in cells isolated from mice or rats. Ten of these cell lines were derived from immature animals and seven from adult animals. Of the cell lines derived from adult animals, two were isolated from tumors (TR-ST and 45T-1). Most of the research on primary cultures of Sertoli cells has been carried out on cells isolated from immature animals, in part because these cells are more readily purified from contaminating germ cells. Cell lines from immature Sertoli cells have, in most cases, been established for the same reason. Several methods of immortalization have also been used to establish Sertoli cell lines. The first Sertoli cell line reported, the TM4 cell line, was established by passing cultures of immature mouse Sertoli cells over

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Kenneth P. Roberts

many generations until an immortal line was obtained. In one case, the TR-ST cell line, the Sertoli cell line was isolated from a spontaneously arising Sertoli cell tumor. In all other cases, a transforming antigen was used to immortalize the cells. Several methods of transforming antigen delivery have been employed in immortalizing Sertoli cells. Direct viral infection of Sertoli cells has proven effective, even with Sertoli cells from adult cells, which do not divide appreciably in culture. Transfection of primary cultures with a vector driving expression of viral transforming antigens has also been used. By far the most common method of transforming antigen delivery has been to create transgenic animals with a transgene driving ubiquitous or Sertoli cell-specific expression of the transforming antigen. The characteristics of each

Sertoli cell line and the method of immortalization are summarized in Table 19.1.

III. ESTABLISHED SERTOLI CELL LINES A. TM4 Cells By far the most widely studied Sertoli cell line is the TM4 cell line. It was established by Mather in 1980 from primary cultures of Sertoli cells isolated from 11to 13-day-old BALB/c mice [1]. These cells were propagated in serum-containing medium and clones selected over time. Unlike most cell lines established recently, no transforming tumor antigen was used to establish these cells, and the cells are not tumorigenic

TABLE 19.1 Summary of Sertoli Cell Line Origins and Properties Cell line

Species

Maturity

Primary uses

Immortalization method

Reference

TM4

Mouse

11–13 day

Gene expression and regulation, endocrine response, toxicology, second messengers, cell cycle, phagocytosis, immune regulation, viral replication

Propagation of primary cultures

S14-1

Mouse

20 day

Characterization only

SV40 virus infection of primary cultures

111

93RS2

Rat

16 day

Characterization only

SV40 virus infection of primary cultures

116

ASC-17D

Rat

Adult

Cell adhesion, toxicology

SV40 virus infection of primary cultures

103

WL/ES

Mouse

15–19 day

Characterization only

Retroviral infection of primary cultures using the HPV 16 genes

114

SG5-2

Mouse

6 day

Characterization only

Transfection of primary cultures with inducible SV40 T antigen

115

SF-7

Mouse

10 day

Germ cell coculture, gene regulation

Transfection of primary cultures with inducible SV40 T antigen

97

SerW3

Rat

17 day

Characterization only

Transfection of primary cultures with inducible SV40 T antigen

117

SM

Ovine

Prepubertal

Characterization only

Transfection of primary cultures with inducible SV40 T antigen

113

MSC-1

Mouse

Adult

Gene regulation, toxicology

Transgenic Sertoli cell–specific expression of SV40 T antigen

68

SMAT-1

Mouse

6.5 day

Characterization only

Transgenic Sertoli cell–specific expression of SV40 T antigen

112

1

TTE-3

Mouse

Adult

Toxicology

Transgenic expression of SV40 T antigen

106

SK11

Mouse

10 day

Characterization only

Transgenic expression of SV40 T antigen

109

15P-1

Mouse

Adult

Germ cell coculture, gene regulation, phagocytosis

Transgenic expression of polyoma T antigen; lines isolated from cultured cells

82,83

42GPA-9

Mouse

Adult

Toxicology

Transgenic expression of polyoma T antigen; lines isolated from cultured cells

82,92

45T-1

Mouse

Adult

Gene regulation

Transgenic expression of polyoma T antigen; lines isolated from tumor

82

TR-ST

Rat

Adult

Toxicology

Spontaneous Sertoli cell tumor

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Chapter 19 Sertoli Cell Lines

when injected subcutaneously into syngeneic nude mice [1]. However, characteristic of transformed cells, recent cytogenetic analysis of TM4 cells shows that they contain an increased and heterogeneous chromosome number, averaging 85 to 95 per cell, as well as stable translocations between chromosomes [2]. Initial characterization studies provided evidence that TM4 cells possess Sertoli cell-like characteristics such as secretion of transferrin and plasminogen activator and expression of the androgen receptor [3]. Very importantly, the TM4 cell line was shown in early studies to be responsive to FSH in both growth and production of cAMP [1]. The production of cAMP in response to FSH was small compared to the responsiveness of primary cultures of Sertoli cells, leading to the suggestion that TM4 cells shared the characteristics of immature Sertoli cells rather than Sertoli cells isolated from pubertal or mature animal [1]. Numerous subsequent studies have confirmed the Sertoli cell-like nature of TM4 cells by comparing their behavior with primary cultures of Sertoli cells. Similar to freshly isolated Sertoli cells, TM4 cells form aggregates when plated on monolayers of primary cultures of myoid cells or a myoid cell line (TR-M) [4]. TM4 cells have been used as a feeder layer for the culture of primordial germ cells isolated from embryos and have been shown, like primary cultures of Sertoli cells, to inhibit gonocyte proliferation [5, 6]. Spermatogonial stem cells, which can be maintained in culture with good viability for 7 days, are rapidly lost when cultured on feeder layers of TM4 cells [7]. These inhibitory effects on maintenance of early germ cells have been attributed to the facilitation of germ cell differentiation that occurs when these cells are grown in contact with Sertoli cells and attests to the fact that TM4 cells interact with early germ cells in a manner similar to that of Sertoli cells. Gene expression studies that have been conducted in both TM4 cells and primary cultures of Sertoli cells show numerous similarities between the two at the molecular level. In situ hybridization analysis of testis sections shows that Sertoli cells express the protein tyrosine phosphatase PTP-RL10 and the protein tyrosine kinase c-Src. The messenger RNAs for both of these enzymes are also found in TM4 cells [8]. Entactin, a basement membrane protein, is made by Sertoli cells and TM4 cells, as well as peritubular cells and TM3 cells, a Leydig cell line [9]. TM4 cells and primary cultures of rat Sertoli cells express the receptor tyrosine kinases Rse and Mer, as well as the ligand for these receptors, Gas6. This receptor system is involved in the regulation of cell growth. Forskolin increases the expression of Gas6 in both primary culture of Sertoli cells and TM4 cells, indicating that Gas6 may regulate, in an autocrine fashion, the proliferation of Sertoli cells [10].

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Several studies using TM4 cells as a model of Sertoli cell function followed the early characterization studies, showing that TM4 cells retained Sertoli cell–like characteristics. The growth-promoting activity of ceruloplasmin was studied using TM4 cells [11]. TM4 cells were shown to respond to seminiferous growth factor (SGF) by increased cell proliferation and an increase in production of sulfated glycoprotein-1 (SGP-1). SGF also altered the secreted protein profile of TM4 cells [12, 13]. TM4 cells are not stimulated to proliferate by secreted proteins from primary cultures of Sertoli cells isolated from immature rats, and adenosine was shown to inhibit TM4 cell proliferation [14, 15]. TM4 cells were shown to secrete a protein that inhibits the hCG-stimulated production of testosterone, and cAMP, from purified Leydig cells [16]. In each of these studies, TM4 cells were used to model Sertoli cell function. TM4 cells have been used to study the formation of cell–cell contacts and the involvement of NCAM, cadherins, and F-actin in their formation, and to synthesize connexin 43, a gap junction protein that is localized to occluding junctions in seminiferous tubules [17, 18]. TM4 cells have also been used, along with many other cell types, to investigate phosphatase activity [19]. 1. Endocrinology of TM4 Cells Because spermatogenesis is an endocrine-regulated process, with many of the endocrine-dependent processes in the testis mediated by Sertoli cells, the endocrine responsiveness of Sertoli cells lines is very important. In the initial characterization of TM4 cells, this line was shown to respond to FSH with increased growth and increased production of cAMP [1]. Sertoli cells and TM4 cells both respond to FSH (or cAMP analogues) with disruption of actin filaments [20]. Additional studies have shown that monolayers of TM4 cells respond to FSH by undergoing a reversible Cl−-dependent depolarization [21]. TM4 cells have also been shown to express both androgen and estrogen receptors, as do primary cultures of Sertoli cells [22]. TM4 cells do not make androgen binding protein (ABP), but ABP has been studied in these cells after stable transfection with an expression vector containing the ABP cDNA [23]. It is well known that vitamin A is important for Sertoli cell function and that vitamin A deficiency causes male infertility. An early study with TM4 cells showed that both retinoic acid and retinol stimulate cell proliferation and increase cell volume [24]. This study also demonstrated that retinoic acid inhibits the growth stimulatory effect of FSH on TM4 cells. TM4 cells have also been shown to synthesize and secrete

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a retinol-binding protein (RBP) that is distinct from either serum or cellular RBP [25]. TM4 cells, like primary cultures of Sertoli cells, secrete a factor with activin-like activity, and they have been shown by radioligand binding assays to have specific binding sites for inhibin A [26, 27]. Western blot analysis and immunocytochemical localization demonstrate that TM4 cells express the epidermal growth factor (EGF) receptor, which has been localized to the Sertoli cells in the intact testis [28]. TM4 cells have been shown to contain vitamin D receptors, as do primary cultures of Sertoli cells, and when treated with vitamin D, TM4 cells mobilize Ca2+ [29]. Calcitonin stimulates an increase in cAMP levels in TM4 cells in a manner similar to that of primary cultures of Sertoli cells [30]. 2. Gene Regulation Studies in TM4 Cells Since their characterization, TM4 cells have been used to examine the molecular mechanism of Sertoli cell gene regulation [31–33]. The earliest such study investigated the regulation of the glucose-regulated protein 78 (GRP78) gene [31]. This study demonstrated that the GRP78 gene is regulated by both testosterone and Ca2+ in TM4 cells, and some of the DNA-binding proteins involved with this regulation were demonstrated. Sertoli cells are known to express the receptor for nerve growth factor (NGF) and the NGF system plays a role in spermatogenesis [34]. The low-affinity nerve growth factor receptor (LNGFR) gene is expressed in TM4 cells and the level of mRNA for LNGFR is increased modestly (2-fold) by retinoic acid and dramatically (10-fold) by testosterone. Transfection studies using 4.8 kb of flanking region fused to the CAT gene were able to locate the regions responsible for regulation by these two factors in TM4 cells [35]. TM4 cells and primary cultures of Sertoli cells both show an increase in inhibin βB subunit mRNA when treated with cAMP. Transfection assays in TM4 cells with a luciferase reporter construct and 1.5 kb of flanking region showed a cAMP-dependent increase in reporter activity [36]. Similar transfection studies in TM4 cells have investigated the regulation of the inhibin α gene and its regulation by cAMP [37]. The Fas system is thought to play a role in regulating immune responses in the testis, and Sertoli cells express the Fas ligand constitutively. Transfection of TM4 cells with deleted and mutated constructs from the 5’ flanking region of the Fas ligand gene demonstrated that the SP1 transcription factor is responsible for the constitutive expression of the gene in TM4 cells and, presumably, Sertoli cells [38].

In studies involving the transfection of an androgenresponsive promoter construct, TM4 cells have been used to show that the Wilms’ tumor suppressor gene product (WT1) is an inhibitor of androgen receptor mediated transcription [39]. The expression of the mer gene, encoding a receptor tyrosine kinase, increases with age in the testis during pubertal development. Transfection studies into TM4 cells using luciferase reporter constructs revealed the regions responsible for the transcriptional activity. DNase footprinting and gel shift assays demonstrated that the transcription factors Sp1, Sp3, and E2F are involved with this regulation [40]. Triiodothyronine (T3) is known to downregulate aromatase activity, an event known to correspond to the terminal differentiation of Sertoli cells. Using TM4 cells, transfection studies using the 5’ flanking region of the aromatase gene linked to the luciferase reporter gene have shown the regulatory region of the gene responsible for T3 inhibition of expression. Gel shift studies, using nuclear extracts from TM4 cells with normal and mutated oligonucleotides from the aromatase regulatory region, show that the T3/thyroid hormone receptor complex binds to the SF1 motif in the aromatase gene, thereby suppressing its transcription [41]. 3. Toxicology Studies in TM4 Cells TM4 cells have been used extensively for the study of toxicants on testicular cells. Gossypol is a polyphenol found in the cottonseed plant that is known to cause testicular dysfunction and infertility, and its use has been proposed as a male contraceptive [42]. However, the mechanism by which gossypol affects the testis is still not fully understood. Several studies have used TM4 cells to test the effects of gossypol on Sertoli cells. A study using both primary cultures of Sertoli cells and TM4 cells has shown both to be growth sensitive to gossypol [43]. In subsequent work, gossypol was shown to damage the mitochondria of TM4 cells and its cell specificity was likely due to the propensity of Sertoli cells to take up gossypol [44–47]. Although gossypol appeared to affect TM4 cells via its effect on the mitochondria, the mechanism by which gossypol inhibits TM4 proliferation probably does not involve inhibition of cellular energetics [48]. Alkylphenols, compounds with estrogenic properties, are thought to disrupt testicular function and to decrease male fertility. Many of these agents have been shown to inhibit testicular Ca2+ ATPase. Studies with TM4 cells show that these compounds (nonylphenol, octylphenol, bisphenol A, and butylated hydroxytoluene) increase cytoplasmic calcium,

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likely by mobilizing intracellular stores, causing apoptosis [49]. This effect could be reversed with a caspase inhibitor, supporting an apoptotic mechanism of alkylphenol action in the testis. Similarly, TM4 cells have been used, along with other testicular cell lines, to study the effects of genistein on cell proliferation and apoptosis [50, 51]. These studies were extended to show that the mechanism of genistein action involves the activation of caspase-3 apoptosis [52]. Humic acid (HA), which has a dramatic detrimental effect on the testis, has been studied using TM4 cells and data from these studies suggest that HA acts by inhibiting Sertoli cell growth [53]. TM4 cells have been used to study the effects of trinitrofluorenone on DNA synthesis, cytotoxicity, and induction of sister chromatid exchange [54]. TM4 cells have also been used to study the effects of methoxyacetic acid, a substance that causes germ cell loss in the testis [55]. TM4 cells were used in microarray experiments to show the toxic effect on gene expression caused by chromium(III) chloride [56]. The results show that several genes are affected by chromium(III) chloride, and that transcription factor Bach2 is particularly affected. A crucial property of the large T antigen, a protein that facilitates transformation in cells, is the binding of this molecule to the Rb tumor suppressor gene. Rodier et al. have shown, using TM4 cells transfected with the polyoma virus large T antigen, that the T antigen inhibits apoptosis induced by taxol, tumor necrosis factor alpha (TNG-α), or the FasR agonists [57]. This effect has implications for the responsiveness of cells transformed by the large T antigen to undergo a normal apoptotic response to cells stressed by exposure to toxicants. 4. Other Studies Using TM4 Cells TM4 cells have been used in studies of second messenger pathways involving inositol 1,4,5-trisphosphate (IP-3) and nitric oxide synthase (NOS). Sertoli cells and TM4 both respond to ATP by activating the IP-3 second messenger signal cascade [58]. TM4 cells also activate this second messenger pathway in response to endothelin-1, whereas primary cultures of Sertoli cells do not. TNFα was shown to act synergistically with interferon γ to increase the production of nitric oxide in TM4 cells [59]. This effect was inhibited by aspirin [60]. Cell cycle regulatory proteins Cdk5, cyclin E, and cyclin D1 have been studied in TM4 cells [61]. Although Cdk5 has been shown to be an important regulator of proliferation in many cell types, this does not appear to be the case in Sertoli cells. The expression of Cdk5 in TM4 cells is not affected by FSH or EGF,

suggesting that this protein regulates some constitutive function of the cell rather than proliferation [62]. TM4 cells have been used to study phagocytosis and certain proteins involved in this process. Tokuda et al. have shown that phagocytosis in TM4 cells is Ca2+ dependent and that phagocytotic activity can be increased by insulin and EGF [63]. The iron transport protein Nramp2 (DMT-1) has been shown to be present in endosomes of TM4 cells [64]. This protein is responsible for the transport of iron across the endosomal membrane. TM4 cells have been used to show that cellular factors determine permissiveness for viral (minute virus of mice) replication [65]. Finally, TM4 cells have been used to investigate the expression of major histocompatibility complex (MHC) and signaling pathways triggered by cross-linking of the CD95 ligand [66, 67].

B. MSC-1 Cells The mouse Sertoli cell 1 (MSC-1) cell line was derived from a mouse carrying a transgene encoding both the small and large T antigens from the temperaturesensitive SV40 virus ts1609 [68]. Sertoli cell–specific expression of the transgene was driven by the Müllerianinhibiting substance (MIS) gene regulatory sequences. These transgenic mice develop testicular tumors with Sertoli cell–like cellular histology but are fertile as young adults, indicating that expression of the T antigen is not incompatible with spermatogenesis. The MSC-1 cell line was cloned from one of the tumors isolated from a mature mouse. MSC-1 cells retain the expression of characteristic Sertoli cell proteins such as transferrin, SGP-2 (clusterin), and inhibin βB. They do not express the FSH receptor, a Sertoli cell–specific gene product, but they are responsive to stimulation by cAMP analogues, indicating that the second messenger system used by the FSH receptor remains intact in MSC-1 cells. In a separate study designed to characterize the MSC-1 cell line, these cells were shown to have similar morphology to primary cultures of mouse Sertoli cells, to express mRNA for ABP, as well as SGP-1, SGP-2, and transferrin, but not the FSH receptor mRNA [69]. This and a later study from the same laboratory showed that SGP-2 is regulated in MSC-1 cells by heat stress in an apparent Sertoli cell–specific manner [70]. Proliferation of MSC-1 cells is significantly reduced when the cells are by incubated with cAMP. The effect of cAMP on the regulation of protein kinase A subunits (RII-α, RII-β, C-α, C-β) has also been studied in MSC-1 cells [71]. The results of this study show that the RII-β subunit of PKA is regulated by cAMP.

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In an attempt to use MSC-1 cells as a model for FSH action in Sertoli cells, Eskola et al. stably transfected MSC-1 cells with a construct encoding the FSH receptor driven by the SV40 promoter/enhancer [72]. This modified MSC-1 cell line expressed the FSH receptor and responded to FSH stimulation with increased synthesis of cAMP. Enhancement of the cAMP production by pertussis toxin and inhibition of cAMP production by TPA demonstrated that the G-protein signaling pathway in these cells mirrored that in Sertoli cells within seminiferous tubules. FSH stimulation of these modified MSC-1 cells resulted in suppression of proliferation, a feature characteristic of maturing Sertoli cells, and an effect consistent with that seen after treating MSC-1 cells with cAMP [69]. At the same time, FSH treatment suppressed the levels of inhibin α mRNA, a response characteristic of immature Sertoli cells. The ability of these cells to respond to FSH has allowed them to be used to characterize the potency of recombinant FSH. Recombinant human FSH was shown to be 10-fold more potent than rat FSH when used to stimulate cAMP production in FSH receptorexpressing MSC-1 cells [73]. Other work using transient transfection of the FSH receptor cDNA into MSC-1 cells showed that G-protein coupled receptor kinase 2 is involved in FSH receptor phosphorylation and internalization [74]. The immortalization of Sertoli cells is often associated with suppression of expression of the FSH receptor gene. Griswold and Kim have demonstrated that methylation of cytosine residues in the core promoter region of the FSH receptor gene is characteristic of cells not expressing the FSH receptor [75]. In DNA isolated from primary cultures of Sertoli cells expressing the FSH receptor, the core promoter region is found to be unmethylated. Moreover, when MSC-1 cells are treated with 5-aza-2’-deoxycytidine to induce demethylation, expression of the FSH receptor gene is activated. This study illustrates a mechanism by which FSH receptor expression can be suppressed when Sertoli cell lines are immortalized. The fact that MSC-1 cells are capable of expressing the FSH receptor when the gene is unmethylated demonstrates that the cellular machinery required for FSH receptor expression is present in these cells. MSC-1 cells have been used to investigate the regulatory elements critical for the expression of the FSH receptor in Sertoli cells [76]. Transfection of various segments of the regulatory region fused to a reporter gene localized several promoter elements required for optimal expression of the FSH receptor and confirmed the presence of a critical E-box element. Gel shift assays demonstrated that upstream stimulatory factors (USF) 1 and 2 are components of the regulatory complexes

binding to the E-box regulatory element. Interestingly, extracts from primary cultures of Sertoli cells had a second E-box-binding complex not found in MSC-1 cells, demonstrating a molecular difference in the FSH receptor regulatory machinery between MSC-1 cells and normal Sertoli cells. In addition to the FSH receptor gene, MSC-1 cells have been used as a model to study the mechanisms regulating Sertoli cell expression of several other genes. The regulatory elements that are responsible for Sertoli cell–specific expression of the ABP gene were deciphered using DNase 1 footprinting, gel mobility shift assays, and transient transfection in MSC-1 cells. Observations suggest that the ABP gene is not regulated directly by FSH or androgens in Sertoli cells [77, 78]. Transient transfection studies in MSC-1 cells show that GATA-binding proteins are involved in the regulation of the inhibin/activin βB gene and that the binding sites for the GATA, sex determining factor and steroidogenic factor 1 regulatory elements found in the 5’ flanking region of the testin gene are responsible for transcription of this gene in Sertoli cells [79, 80]. Finally, high levels of promoter activity were found for the MSC-1 retrovirus in transiently transfected MSC-1 cells [81]. Expression of this retrovirus in vivo is restricted to Sertoli cells isolated from rats of 20 and 27 days of age. Although few toxicology studies have been published using MSC-1 cells, methoxyacetic acid (MMA), a chemical that causes germ cell loss via apoptosis in the testis, has been shown to decrease the production of ABP and ABP mRNA in MSC-1 cells, suggesting that at least part of the effect of MMA on germ cells is mediated via effects on Sertoli cells [55].

C. 15P-1 Cells Sertoli cell line 15P-1 was established from Sertoli cells of a transgenic mouse expressing the polyoma virus large T antigen [82]. This cell line was shown to express the Wilm’s tumor and Steel genes, consistent with a Sertoli cell origin for these cells [83]. 15P-1 cells have been used extensively to study differentiation of germ cells in culture. When 15P-1 cells were cocultured with a mixture of testicular cells isolated from a transgenic mouse expressing the lacZ gene under control of the protamine regulatory region (Prm-1), the number of blue-staining advanced spermatids increased dramatically during the culture period, suggesting that ongoing germ cell differentiation was supported by this cell line. Flow cytometric and 3H-thymidine uptake studies confirmed that meiosis was supported in this coculture system. These experiments demonstrate that immortalized Sertoli cells can

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retain the necessary phenotypic functions to support at least limited germ cell differentiation in vitro. In subsequent studies 15P-1 cells have been used as a model to study Sertoli cell phagocytic activity [84]. In an assay involving the uptake of fluorescent latex beads, a culture of 15P-1 cells contains a small fraction of cells that actively phagocytose the beads. However, when germ cells are added to the cultures, in particular pachytene spermatocytes, all of the 15P-1 cells become phagocytic. This activation of phagocytosis required contact with the germ cells, but phagocytic activity was sustained by factor(s) secreted into the medium. Later stages of meiotic germ cells inhibited the phagocytic response. 15P-1 cells have also been used to investigate the localization of the endosomal protein Nramp2 in phagosomes [64]. The molecular mechanisms of FSH receptor gene expression have been extensively studied in 15P-1 cells. Deletion analysis of the promoter region fused to the luciferase gene localized a strong promoter 3’ of the transcriptional start site that was further mapped by DNase 1 footprinting [85]. Gel mobility shift assays revealed that upstream regulatory factors 1 and 2 bound to this proximal promoter. These transcription factors were also shown to be involved with binding to the E box in studies performed with MSC-1 cells [76]. Mutation of the E box resulted in loss of activity from the proximal promoter in 15P-1 cells as well as elimination of activity from a minimal promoter construct containing only the E-box element. A second region with DNA binding activity was localized using DNase1 footprinting just 5’ to the transcription start site [86]. Binding of nuclear protein from 15P-1 cells to this region could be competed away with sequences encoding binding sites for Krupple-like transcription factors, ZNF202-like factor and Ras-responsive element binding protein-like factor. Cotransfection studies confirmed the potential involvement of these two transcription factors in regulation of FSH receptor gene expression, with Ras-responsive element binding protein 1 activating transcription and ZNF202 repressing transcription [87]. Peptides with antibiotic activity have been found in Sertoli cells and also in 15P-1 cells [88]. This activity was increased with 15P-1 cells that were cocultured with round spermatids and nerve growth factor. Expression of the antibiotic genes cryptdin and cryptdinrelated genes was confirmed and cryptdins 1, 2, 3, and 6 were found in the culture medium. The ability to genetically manipulate cell lines allows for genetic selection strategies to be employed that would not be possible with primary cultures of Sertoli cells. Taking advantage of the known gene activation response of 15P-1 cells to coculture with germ

cells, a gene trap strategy was devised to identify genes involved in this cell–cell interaction. A promoterless construct encoding the (β)geo drug resistance transgene was randomly integrated into the genome of 15P-1 cells and 2000 clones were selected via the drug resistance encoded in the transgene [89]. These clones were then screened for those that upregulate expression of the β-galactosidase gene in the presence of germ cells or NGF. From this screen a series of genes was discovered that are regulated by germ cells and/or by stimulation with NGF. One of the upregulated genes was Fra1, a constituent of the AP1 regulatory complex. Primary cultures of Sertoli cells were shown to induce Fra1 in a manner similar to 15P-1 cells, confirming the physiological nature of this response. The 15P-1 cells were then used to show that the effects of NGF were mediated by the TrkA receptor and the ERK1-ERK-1 kinase kinase pathway. Successful implementation of sophisticated experimental strategies such as this one relies completely on the availability of cell lines.

D. 45T-1 Cells The 45T-1 cell line was derived from the same transgenic mouse as the 15P-1 cell line, but this cell line was isolated from a testicular tumor and not from cultured Sertoli cells [82]. In culture, the tumor cells from which the 45T-1 cells were derived grow in close contact with other cells derived from germ cells [82]. Subsequently, the 45T-1 cells have been used to investigate the role inhibin plays in the regulation of cell proliferation in the testis [90]. The tumors and the cell line both display reduced levels of inhibin production. When transfected with a construct directing expression of inhibin, the growth of these cells was reduced, suggesting that inhibin may regulate Sertoli cell growth and reduction in inhibin production may lead to Sertoli cell tumor formation. The role of TNFα in the development of testicular differentiation was investigated using 45T-1 cells. When treated with TNFα, 45T-1 cells form structures resembling spermatic cords and, in conjunction with these morphological changes, upregulate the synthesis of laminin [91]. The same effect was achieved in coculture with germ cells, suggesting that TNFα plays a role in the development of the in vivo organization of the seminiferous tubule.

E. 42GPA-9 Cells The 42GPA-9 cell line was established from the same transgenic mice expressing the polyoma virus large T antigen used to establish the 15P-1 and 45T-1

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cell lines [82]. These cells were isolated from the testes of adult mice (6 months old) and subsequently cloned by serial dilution to establish a pure clonal cell line [92]. The 42GPA-9 cells display characteristics of Sertoli cells including expression of transferrin, SGP-2, the c-kit ligand, and plasminogen activator. The expression of plasminogen activator and plasminogen activator inhibitor 1 were altered by stimulation of 42GPA-9 cells with LPS or TPA, compounds that effect phagocytosis in Sertoli cells [93]. FSH receptor mRNA could also be detected in these cells and cAMP production was stimulated by treatment with FSH. EM ultrastructural analysis demonstrated Sertoli cell–like morphology of these cells, including the formation of tight junctional complexes [94]. Western blot analysis confirmed that 42GPA-9 cells express connexin 43 (Cx43), and immunofluorescent studies show localization of this protein to gap junctional complexes. The gap junctional complexes in 42GPA-9 cells are disrupted when the cells are treated with the reproductive toxin lindane [95]. This toxin appears to affect the localization of proteins Cx43 and zonula occludins 1 (ZO-1) to the gap junctions, causing them to accumulate in the cytoplasm. This effect of lindane was shown to involve activation of the ERK pathway in 42GPA-9 cells [96].

F. SF-7 Cells The SF-7 cell line was produced by transfection of testicular cells isolated from 10-day-old mice with the SV40 large T antigen [97]. These cells exhibit Sertoli cell–like morphology when cultured at high density and express the FSH receptor, albeit at lower levels than freshly isolated Sertoli cells. When cocultured with other testicular somatic cell lines, structures resembling two-dimensional spermatic cords can be formed. These cords can support the survival of pachytene spermatocytes for more than 1 week, suggesting they retain significant Sertoli cell-like function. In a recent study it was shown that spermatogonial stem cells are rapidly lost when cocultured with SF-7 cells [7]. These stem cells can be maintained in culture alone with up to 25% viability for 7 days. The loss of stem cells during coculture is likely due to the facilitation of germ cell differentiation that occurs when these cells are grown in contact with Sertoli cells or Sertoli-like cell lines. SF-7 cells alone have also been shown to form hollow tubules in culture on Matrigel. The basement membrane components laminin and collagen IV, along with hepatocyte growth factor, are required for the remodeling of these cells into tubules [98]. SF-7 cells have been used to study the transcriptional regulation of the prosaposin and stem cell factor

(SCF) genes [99, 100]. In Sertoli cells, SCF is regulated by FSH via the second messenger cAMP. SCF-7 cells were transfected with vectors containing the SCF gene regulatory region fused to the luciferase reporter gene sequence [100]. Expression from a cAMP responsive element was demonstrated using a construct containing only 162 bp of SCF 5’ flanking sequence. A distal regulatory element suppressed basal expression but did not suppress cAMP activation of transcription. Interestingly, FSH had no effect on expression from these constructs in SF-7 cells even though they appeared to express the FSH receptor.

G. TR-ST Cells The TR-ST cell line was established from cells of a spontaneous testicular tumor in a rat found to have abnormally high circulating levels of ABP [3]. This cell line was reported to have a growth response to FSH and secrete plasminogen activator and ABP. Like primary cultures of Sertoli cells, the rat TR-ST cell line expresses both androgen and estrogen receptors [22]. The TR-ST cell line has been used along with other cell lines to characterize the growth-promoting effect of ceruloplasmin [11]. This cell line will form aggregates when plated on monolayers of primary cultures of myoid cells or a myoid cell line (TR-M) [4]. TR-ST cells phosphorylate specific proteins in response to coculture with germ cells [101]. Taken together, these data show that TR-St cells have numerous Sertoli cell–like features. TR-ST cells have been used as a model system to investigate the effect of gossypol on testicular cells [43]. Gossypol inhibits proliferation and causes several morphological changes in TR-ST cells, but does not kill the cell [102]. The effects of gossypol include altering the function of the mitochondria, as evidenced by aberrant uptake of rhodamine 123. TR-ST cells are much more sensitive than other non–Sertoli cell lines to these effects of gossypol.

H. ASC-17D Cells The ASC-17D cell line was created by immortalization of Sertoli cells isolated from adult rats with the temperature-sensitive SV40 virus, tsA255 [103]. These cells were shown to express cytokeratin, vimentin, and SGP-1, consistent with a Sertoli cell origin for these cells. When propagating at the permissive temperature for the SV40 large T antigen, these cells made very little transferrin or SGP-2, two secreted proteins characteristic of Sertoli cells. However, when the cells were cultured at 39°C, the nonpermissive temperature, proliferation of the cells ceased and expression of both transferrin and SGP-2 was increased. ASC-17D cells

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did not respond directly to FSH, but the expression of both transferrin and SGP-2 were increased by treatment of the cells with cAMP at the nonpermissive temperature. Because the expression of SGP-2 is increased in many cell types under stress conditions and during apoptosis, it is possible that culturing Sertoli cell lines transformed with the temperature-sensitive SV40 virus at the nonpermissive temperature could induce SGP-2 expression due to stress and not differentiation. The ASC-17D cell line was shown to increase expression of SGP-2 at 39°C without evidence of apoptosis when evaluated for DNA laddering [103]. The ASC-17D cell line has been used to study the role of clusterin (SGP-2) in cell adhesion [104]. The expression of clusterin was increased transiently early after plating when the ASC-17D cells were attaching to the culture dish. Inhibition of clusterin synthesis using an antisense oligonucleotide decreased the efficiency of attachment of the ASC-17D cells to the culture plate and resulted in significant apoptosis of the cells. This effect was more pronounced when clusterin synthesis was inhibited prior to plating. The ASC-17D cell line has also been used to study the toxic effect of dinitrobenzene, dinitrotoluene, and cadmium chloride on Sertoli cells [105].

I. TTE-3 Cells TTE-3 cells were established from sexually mature transgenic mice expressing the temperature-sensitive large T antigen, tsA58 [106]. These cells exhibited many Sertoli cell–like characteristics including expression of transferrin, SGP-2, inhibin α, Steel factor, and the FSH receptor. The expression of transferrin and SGP-2 was increased when the cells were cultured at the nonpermissive temperature while expression of other Sertoli cell genes remained essentially unchanged. These cells synthesize ZO-1, a protein associated with tight junctions, and this protein was localized by immunocytochemistry to the periphery of the TTE-3 cells. This expression and staining pattern was markedly increased when the cells were cultured at the nonpermissive temperature of 39°C. TTE-3 cells have been used in subsequent toxicology studies where microarray analysis was performed after treatment with bisphenol A (BPA) [107]. Several genes were shown to be elevated by treatment with this toxicant, including chop-10, fra-2, c-myc, and ornithine decarboxylase, with chop-10 affected most dramatically [108].

J. Other Sertoli Cell Lines A number of cell lines have been established and their characterization described in the literature but

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with few or no subsequent studies published using the cells. The following cell lines fall into this category. 1. SK11 Cells A series of cell lines have been cloned from Sertoli cells isolated from sexually immature (10-day-old) H-2Kb-tsA58 transgenic mice expressing the temperature-sensitive SV40 large T antigen [109]. The SK11 cell line was representative of these cell lines and displayed many Sertoli cell–like characteristics, including low levels of FSH receptor expression. Most of the Sertoli cell marker genes expressed by these cells were present at both the permissive and nonpermissive temperatures. SGP-1 and GATA-1 expression were significantly increased in expression at the nonpermissive temperature. In this initial characterization study, the expression of ABP or aromatase could not be demonstrated. However, a second study has shown that these cells do in fact synthesize aromatase when cultured with forskolin at the nonpermissive temperature [110]. 2. S14-1 Cells S14-1 cells were derived by infection of mouse Sertoli cells isolated from 20-day-old mice with the temperature-sensitive strain of the SV40 virus, tsA255. The express purpose of establishing these cells, in addition to obtaining a cell line with Sertoli cell characteristics in vitro, was to produce a cell type that could be used as a model of Sertoli cell malignancy in vivo [111]. These cells were shown to secrete transferrin and to express SGP-2 mRNA, both characteristic of primary cultures of Sertoli cells. Both of these functions were increased in culture when the temperature was raised to 40°C. These cells were also shown to have tumorigenic properties and, when injected into the peritoneum, to form tumors with features of primitive seminiferous tubules. Electron micrographic analysis of these tumors demonstrated cell-to-cell junctions and interdigitated processes of plasma membrane characteristic of Sertoli cells within seminiferous tubules. 3. SMAT-1 Cells SMAT-1 cells were derived from Sertoli cells isolated from the testis of 6.5-day-old transgenic mice expressing the SV40 large T antigen from the anti-Müllerian hormone (AMH) promoter [112]. These cells display many of the characteristics of immature mouse Sertoli cells, including the expression of WT-1, ABP, SCF, SGP-2, inhibin α, and inhibin βB mRNAs. The feature that separates this Sertoli cell line from others is the

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demonstrated expression of AMH and the type II AMH receptor. 4. SM Cell Lines The unique feature of the SM cell lines is that they were established from Sertoli cells isolated from the testes of prepubertal lambs. Sertoli cells were immortalized by transfection with the SV40 large T antigen with expression of the tumor antigen driven by the vimentin promoter sequence [113]. These cell lines, named SM1–7, display Sertoli cell–like morphology in culture and all express mRNA for SGP-2, inhibin α, WT-1, transferrin, and Sox9. 5. WL and ES Cell Lines A series of cell lines has been established from testicular cells of 15- to 19-day-old wild-type and α-ERKO knockout mice deficient in estrogen receptor α by retroviral infection using the HPV 16 genes [114]. The objective of this study was not to establish Sertoli cell lines specifically, but to establish testicular cell lines for the study of cellular processes dependent on particular estrogen receptor mediated mechanisms. Successful cloning of cell lines that express one, both, or neither estrogen receptor was accomplished and some of these cell lines, including WS4, WL3, and ES4, have Sertoli cell–like features. However, each of these cell lines also has characteristics of peritubular cells as well. Nevertheless, these cells may prove useful in deciphering some of the relative roles of the estrogen receptor types in testicular function. 6. SG5-2 Cells A series of cell lines was created with the goal of establishing an in vitro Sertoli cell culture system that could be used to support and study germ cell development. These cell lines were created by transfection of Sertoli cells derived from 6-day-old mice with SV40 large T antigen driven by the ponasterone A inducible promoter [115]. This construct system allows the large T antigen expression to be shut down by removal of the inducing factor ponasterone A. However, longterm passage of these cell lines ultimately resulted in the constitutive expression of the large T antigen. The SG5-2 cell line was representative of the most Sertoli cell–like of the sublines derived from this cloning experiment. The SG5-2 cell line has Sertoli cell–like morphology, is negative for Leydig and peritubular cell markers, and expresses many genes characteristic of Sertoli cells including stem cell factor, SGP-2, TGFβ, inhibin α, GATA-1, and steroidogenic factor 1.

None of the cell lines established, including SG5-2 cells, appeared to express the androgen receptor or the FSH receptor RNA. When used in coculture experiments, these cells supported the early stages of spermatogonial development and the survival of type A spermatogonia for a minimum of 7 days. 7. 93RS2 Cells The Sertoli cell line 93RS2 was established by immortalizing Sertoli cells from prepubertal (16-day-old) rats with the temperature-sensitive strain of the SV40 virus tsA255 [116]. This is the same immortalization method used to establish the adult rat Sertoli cell line ASC-17D with similar results. These cells propagate rapidly at the permissive temperature of 32°C but much more slowly when switched to the nonpermissive temperature of 40°C. The 93RS2 cells express the Sertoli cell markers transferrin, SCF, SGP-1, and SGP-2. Like ASC-17D cells, the level of SGP-2 mRNA is induced at the nonpermissive temperature, as is the level of mRNA for SCF. Transferrin mRNA levels are stimulated by testosterone at the nonpermissive temperature. Although the parent cells displayed some FSH responsiveness, the 93RS2 cells do not make FSH receptor, nor do they produce mRNA for ABP. 8. SerW3 Cells The SerW3 Sertoli cell line was established by transfection of Sertoli cells isolated from immature (17day-old) Wistar rats with a plasmid expressing the SV40 large and small T antigen [117]. From the transformants, the SerW3 cells were selected based on their morphological similarities to Sertoli cells. These cells were shown to form tight junctions with adjacent cells in culture and to have nuclei characteristic of Sertoli cells. SerW3 cells show a distinct sensitivity to cisplatin, as do Sertoli cells, and these cells secrete transferrin. This cell line was not responsive to FSH.

IV. SUMMARY As this brief review demonstrates, many various Sertoli cell lines are available for research. Some of these cell lines retain very specialized functional properties of Sertoli cells such as expression of the FSH receptor and the ability to support germ cell differentiation in coculture experiments. Several of these cell lines have been used extensively for studies of the molecular mechanisms governing Sertoli cell gene regulation. Some have also proven useful for studies of Sertoli cell response to known testicular toxins.

Chapter 19 Sertoli Cell Lines

The study of cellular functions such as phagocytosis has been facilitated by some of these cell lines. Although no one of these cell lines perfectly reflects all the attributes of native Sertoli cells, any particular aspect of Sertoli cell function is likely represented by one or more of these cell lines. Nevertheless, the primary culture of freshly isolated Sertoli cells remains the “gold standard” to which all experiments with Sertoli cell lines must still be compared.

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98. van der Wee, K., and Hofmann, M. C. (1999). An in vitro tubule assay identifies HGF as a morphogen for the formation of seminiferous tubules in the postnatal mouse testis. Exp. Cell Res. 252, 175–185. 99. Sun, Y., Jin, P., and Grabowski, G. A. (1997). The mouse prosaposin locus: Promoter organization. DNA Cell Biol. 16, 23–34. 100. Taylor, W. E., Najmabadi, H., Strathearn, M., Jou, N. T., Liebling, M., Rajavashisth, T., Chanani, N., Phung, L., and Bhasin, S. (1996). Human stem cell factor promoter deoxyribonucleic acid sequence and regulation by cyclic 3’,5’adenosine monophosphate in a Sertoli cell line. Endocrinology 137, 5407–5414. 101. Ireland, M. E., and Welsh, M. J. (1987). Germ cell stimulation of Sertoli cell protein phosphorylation. Endocrinology 120, 1317–1326. 102. Tanphaichitr, N., Chen, L. B., and Bellve, A. R. (1984). Direct effect of gossypol on TR-ST cells: Perturbation of rhodamine 123 accumulation in mitochondria. Biol. Reprod. 31, 1049–1060. 103. Roberts, K. P., Banerjee, P. P., Tindall, J. W., and Zirkin, B. R. (1995). Immortalization and characterization of a Sertoli cell line from the adult rat. Biol. Reprod. 53, 1446–1453. 104. Kang, S. W., Lim, S. W., Choi, S. H., Shin, K. H., Chun, B. G., Park, I. S., and Min, B. H. (2000). Antisense oligonucleotide of clusterin mRNA induces apoptotic cell death and prevents adhesion of rat ASC-17D Sertoli cells. Mol. Cells 10, 193–198. 105. Sorenson, D. R., and Brabec, M. (2003). The response of adult rat Sertoli cells, immortalized by a temperature-sensitive mutant of SV40, to 1,2-dinitrobenzene, 1,3-dinitrobenzene, 2,4-dinitrotoluene, 3,4-dinitrotoluene, and cadmium. Cell Biol. Toxicol. 19, 107–119. 106. Tabuchi, Y., Ohta, S., Yanai, N., Obinata, M., Kondo, T., Fuse, H., and Asano, S. (2002). Development of the conditionally immortalized testicular Sertoli cell line TTE3 expressing Sertoli cell specific genes from mice transgenic for temperature sensitive simian virus 40 large T antigen gene. J. Urol. 167, 1538–1545. 107. Tabuchi, Y., Zhao, Q. L., and Kondo, T. (2002). DNA microarray analysis of differentially expressed genes responsive to

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bisphenol A, an alkylphenol derivative, in an in vitro mouse Sertoli cell model. Jpn. J. Pharmacol. 89, 413–416. Tabuchi, Y., and Kondo, T. (2003). cDNA microarray analysis reveals chop-10 plays a key role in Sertoli cell injury induced by bisphenol A. Biochem. Biophys. Res. Commun. 305, 54–61. Walther, N., Jansen, M., Ergun, S., Kascheike, B., and Ivell, R. (1996). Sertoli cell lines established from H-2Kb-tsA58 transgenic mice differentially regulate the expression of cell-specific genes. Exp. Cell Res. 225, 411–421. Walther, N., Jansen, M., Ergun, S., Kascheike, B., Tillmann, G., and Ivell, R. (1997). Sertoli cell-specific gene expression in conditionally immortalized cell lines. Adv. Exp. Med. Biol. 424, 139–142. Boekelheide, K., Lee, J. W., Hall, S. J., Rhind, N. R., and Zaret, K. S. (1993). A tumorigenic murine Sertoli cell line that is temperature-sensitive for differentiation. Am. J. Pathol. 143, 1159–1168. Dutertre, M., Rey, R., Porteu, A., Josso, N., and Picard, J. Y. (1997). A mouse Sertoli cell line expressing anti-Müllerian hormone and its type II receptor. Mol. Cell Endocrinol. 136, 57–65. Merhi, R. A., Guillaud, L., Delouis, C., and Cotinot, C. (2001). Establishment and characterization of immortalized ovine Sertoli cell lines. In Vitro Cell Dev. Biol. Anim. 37, 581–588. Mueller, S. O., and Korach, K. S. (2001). Immortalized testis cell lines from estrogen receptor (ER) alpha knock-out and wildtype mice expressing functional ERalpha or ERbeta. J. Androl. 22, 652–664. Hofmann, M. C., Van Der Wee, K. S., Dargart, J. L., Dirami, G., Dettin, L., and Dym, M. (2003). Establishment and characterization of neonatal mouse Sertoli cell lines. J. Androl. 24, 120–130. Jiang, C., Hall, S. J., and Boekelheide, K. (1997). Development and characterization of a prepubertal rat Sertoli cell line, 93RS2. J. Androl. 18, 393–399. Pognan, F., Masson, M. T., Lagelle, F., and Charuel, C. (1997). Establishment of a rat Sertoli cell line that displays the morphological and some of the functional characteristics of the native cell. Cell Biol. Toxicol. 13, 453–463.

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VII SERTOLI CELL PATHOPHYSIOLOGY

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C H A P T E R

20 Sertoli Cell Toxicants KIM BOEKELHEIDE

KAMIN J. JOHNSON

Department of Pathology and Laboratory Medicine, Brown University, Providence, Rhode Island

Division of Biological Sciences, CIIT Centers for Health Research, Research Triangle Park, North Carolina

JOHN H. RICHBURG Division of Pharmacology and Toxicology, College of Pharmacy, The University of Texas at Austin, Austin, Texas

I. II. III. IV. V. VI. VII.

model systems that provide important mechanistic information are included. Section II, Definition, describes the fundamental characteristics that identify Sertoli cell toxicants. A short Mechanisms section provides a simple framework for conceptualizing a few of the means by which toxicants might result in Sertoli cell–specific injury. Section IV, Manifestations, includes descriptions and illustrations of the morphological, biochemical, and molecular alterations that result from toxicant-induced Sertoli cell injury. The Literature Review section summarizes the available information concerning the chemistry, metabolism, toxicology, pathology, and mechanism of action of Sertoli cell toxicants. A section entitled Reversibility is included with a brief introduction of the important issue of germ cell repopulation following exposure to Sertoli cell toxicants. Finally, the text ends with a section that summarizes the current state-of-knowledge in this field.

INTRODUCTION DEFINITION MECHANISMS MANIFESTATIONS LITERATURE REVIEW REVERSIBILITY CONCLUSIONS References

I. INTRODUCTION A major function of the Sertoli cell is to create an environment appropriate for germ cell proliferation and maturation. Consequently, alterations in Sertoli cell function can result in germ cell loss and subsequent infertility. In this chapter, we will discuss toxicants that specifically target the Sertoli cell, resulting in germ cell damage and disruption of functional spermatogenesis. The reader should view these toxicants as potential tools to better delineate those Sertoli cell functions critical to germ cell viability. The success of this chapter will be measured by its ability to stimulate new thinking about potential mechanisms of toxicant action while serving as a practical laboratory manual for the identification and understanding of Sertoli cell toxicants. The focus is on xenobiotics of environmental and industrial relevance and their in vivo effects, though model compounds and SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

II. DEFINITION The term toxicant encompasses any toxic agent of natural, biological, or synthetic origin and is used throughout this chapter in preference to toxin which, though often misapplied, is limited by definition to toxic agents of biological origin. The identification of the Sertoli cell as the target for a testicular toxicant is a difficult task. The difficulty resides in the complex

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interdependence of cells within the testis and our rudimentary understanding of the molecular and cellular events that underlie the symbiotic interactions of these cells. In the absence of a more complete understanding of testicular homeostasis, toxicant-induced perturbations in germ cell production are assumed to result from dysfunction of the first cell that manifests alterations during an in vivo exposure. Therefore, a testicular toxicant producing its earliest detected alterations in Sertoli cells is considered a Sertoli cell toxicant. The accurate designation of the Sertoli cell as the target of a testicular toxicant depends on the quantity and quality of available information. Theoretically, any testicular toxicant may produce subtle alterations in germ cell function that trigger secondary manifestations of Sertoli cell dysfunction as the earliest observed sign of testicular injury. Such an unrecognized germ cell toxicant is considered a Sertoli cell toxicant until the knowledge base is improved by further experimentation. Alternatively, any testicular toxicant producing subtle, undetected alterations in Sertoli cell function followed by germ cell abnormalities as the earliest sign of testicular injury is incorrectly classified as a germ cell toxicant. The frequency of incorrect assignment of toxicant to target cell is unknown; however, given the central role of the Sertoli cell in germ cell maintenance, one would predict that cryptic Sertoli cell toxicants are more often misidentified as germ cell toxicants than vice versa. Only toxicants that fulfill three criteria⎯production of early and specific Sertoli cell alterations following an in vivo exposure⎯are considered in this review. Early is a relative term which dictates that the toxicantinduced Sertoli cell alterations occur before observable changes in other testicular cell types. The time frame for the manifestation of Sertoli cell abnormalities after an exposure may vary considerably among toxicants. For example, Sertoli cell vacuolation is the earliest morphological response of a testis exposed to either di-(n-pentyl)phthalate or 2,5-hexanedione; however, vacuolation occurs within 3 hr of an appropriate phthalate exposure and not until 3 weeks after a continuous exposure to 2,5-hexanedione. The biological relevance of the toxic response is ensured by the selective induction of Sertoli cell abnormalities in a setting in which many cell types could be injured; thus, the focus in this review is on those toxicants for which testicular abnormalities are a prominent and specific consequence of an in vivo exposure. Using these criteria, Sertoli cell toxicants are considered as molecular probes for mechanistic investigations that elucidate those structural and functional characteristics that make the Sertoli cell unique.

III. MECHANISMS This section highlights the modes by which toxicants perturb Sertoli cell function. Sertoli cell toxicants likely target characteristics unique to Sertoli cells, resulting in germ cell apoptosis; some of these characteristics are shown in Figure 20.1. To be considered mechanistically important, observed alterations must be early, primary events in the toxic response rather than secondary alterations. Secondary alterations are termed manifestations of the toxic response and are considered later. A significant limitation to mechanistic studies of Sertoli cell toxicants is the lack of a comprehensive understanding of Sertoli cell biology. Because of this, no Sertoli cell toxicants of environmental and occupational significance have a completely understood mechanism of action.

A. Altered Germ Cell Attachment All germ cells show adhesive contact with Sertoli cells that is mediated through adhesion junctions as well as a carbohydrate-based system [1, 2]. Sertoli–germ cell adhesion junctions are attractive mechanistic targets

FIGURE 20.1 This schematic drawing illustrates the various mechanisms discussed in the text by which toxicants might produce Sertoli cell dysfunction and testicular injury. (Reproduced with permission of Cache River Press.)

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because of their unique ultrastructural morphology and dynamic molecular composition as well as the specificity of germ cell adhesion to Sertoli cells [1, 3, 4]. Genetic deletion of a Sertoli cell adhesion protein (nectin-2) that produces defects in germ cell maturation highlights the importance of Sertoli–germ cell adhesion in spermatogenesis [5, 6]. Loss of germ cell attachment to the seminiferous epithelium and the appearance of germ cells in the lumen is observed with several Sertoli cell toxicants, including nitroaromatics [7] and phthalates [8, 9], and is interpreted as a loss of Sertoli–germ cell adhesion. In addition, germ cell detachment from underlying Sertoli cells is observed in cocultures exposed to Sertoli cell toxicants [10]. Because intercellular adhesion generates cell survival signals in other tissues [11], loss of Sertoli–germ cell adhesion may precede germ cell apoptosis and be an early and specific effect of Sertoli cell toxicants rather than a secondary manifestation of exposure.

B. Blood–Testis Barrier Defect Formation and maintenance of the blood–testis barrier is a major Sertoli cell function. The blood–testis barrier is composed of a morphologically unique tight junctional complex [12], making it an appealing mechanistic target. Knockout of the Sertoli cell tight junction protein claudin-11 produces a restricted phenotype that includes male sterility [13]. Claudin-11 deficient mice exhibit extensive sloughing of Sertoli and germ cell aggregates into the tubule lumen, reminiscent of some Sertoli cell toxicants. To date, very little is known regarding alterations in tight junction–associated protein or mRNA expression following exposure to a known Sertoli cell toxicant. Two Sertoli cell toxicants, cisplatin [14] and 1-(2,4-dichlorobenzyl)-1H-indazole3-carboxylic acid [15], produce blood–testis barrier defects, though these results are supported by single studies using a single technique. A thorough evaluation of blood–testis barrier integrity, including localization and expression of tight junctional proteins, should be considered for any Sertoli cell toxicant producing cell sloughing or early degeneration of adluminal germ cells.

C. Insufficient Apical Cytoskeletal Support Sertoli cell cytoskeletal components play a central role in maintaining the structure and function in this elongate, highly asymmetric cell type. A structural role for Sertoli cell microtubules is suggested by studies of the microtubule disrupting agent, colchicine,

which results in sloughing of the apical seminiferous epithelium [16]. Presumably, colchicine-induced loss of cytoplasmic microtubules leaves the thin apical projections of the Sertoli cell without sufficient mechanical support, causing breakage of the seminiferous epithelium between cohorts of germ cells [16]. Intermediate filaments also perform a germ cell anchoring role. Within Sertoli cells, bundles of intermediate filaments project from a perinuclear region to Sertoli–germ cell adhesion junctions [17]. Alterations in microtubules or intermediate filaments lead to disturbed structural support of the adluminal compartment and cell sloughing. Such a mechanism is important for postnatal phthalateinduced injury, because a disruption of Sertoli cell intermediate filaments is an early response in this model [18].

D. Metabolic Insult Tight junctions between adjacent Sertoli cells create two separate compartments within the seminiferous epithelium: a basal compartment below the tight junction and an adluminal compartment above. Sertoli cells secrete hormonal and nutritive factors into the adluminal compartment to create a specialized microenvironment to foster the development and viability of germ cells that reside there. One important factor that Sertoli cells secrete into the adluminal compartment is lactate [19, 20]. In general, germ cells, particularly postmeiotic germ cells, utilize lactate exclusively as their source for energy metabolism because they are unable to utilize glucose [21–23]. As a result, germ cells in the adluminal compartment require a constant supply of Sertoli cell– produced lactate and possibly pyruvate to promote their viability, possibly by inhibiting their ability to undergo apoptosis [21, 24, 25]. Therefore, metabolic dysfunction leading to altered nutrient supply is one mechanism by which Sertoli cell toxicants could result in germ cell death and testicular atrophy. Toxicant-induced alterations in metabolism do occur, because Sertoli cells in culture exposed to 1,3-dinitrobenzene or mono-(2-ethylhexyl) phthalate increase their output of lactate in a dosedependent manner [26]. It is likely that this is a response of Sertoli cells to toxicant-induced injury; therefore, toxicant-induced Sertoli cell metabolic alterations are useful as an in vitro screening tool [26] and also may play an etiological role in some forms of testicular injury.

E. Microtubule-Dependent Transport Defects Secretion of seminiferous tubule fluid is a Sertoli cell function [27]. Recent studies have modified long-standing hypotheses about the composition and

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formation of the seminiferous tubule fluid [28, 29]. Though the overall mechanism of seminiferous tubule fluid formation and secretion is ill understood, microtubule-dependent transport is clearly critical to this process. This assertion is based on morphological alterations following colchicine exposure [30], the requirement for microtubule-dependent transport for apical protein secretion in other polar cells [31], and a detailed analysis of the secretory consequences of disruption of microtubule-dependent transport in isolated seminiferous tubules [32]. Because the viability of adluminal germ cells depends on the provision of nutritional and hormonal factors in the seminiferous tubule fluid, disruption of the secretory process is one mechanism by which Sertoli cell dysfunction could result in germ cell apoptosis and testicular atrophy. This mechanism of action was proposed for 2,5-hexanedione [33], a Sertoli cell toxicant that alters the assembly properties of testicular microtubules during early exposure [34].

F. Receptor/Second Messenger Alterations The Sertoli cell is responsive to many different peptide hormones and steroids that influence its function. The most widely characterized hormones to which the Sertoli cell responds are follicle-stimulating hormone (FSH), estrogens, androgens, and thyroid hormone. Each of these hormones binds to receptors to activate pathways within the Sertoli cell. A description of the physiological actions of many of these agents is detailed in a series of earlier chapters in this book. Alterations in the function of these receptors, their expression, and/or elements of the intracellular pathways they control are other targets to consider in the evaluation of toxicant-induced Sertoli cell dysfunction. For example, exposure of Sertoli cells in vitro to mono-(2-ethylhexyl) phthalate reduces the FSHdependent increases in intracellular cAMP [35]. In addition, we have an increased appreciation of the role of hormones in reproductive tract development. Studies of the effects of “endocrine-disrupting chemicals” and the availability of animal models with altered hormonal responsiveness drive this new insight. The observation of altered male reproductive tract development in alligators living in a DDEcontaminated lake [36, 37] and the creation of the ERKO (estrogen receptor α knockout) mouse [38] are hallmark events for this field, and the literature in this area is expanding rapidly. An apparent increase in the prevalence of human male reproductive tract abnormalities during the past half-century led Skakkebaek to articulate “testicular

dysgenesis syndrome” as an explanation for this phenomenon. According to Skakkebaek [39, 40], the observed abnormalities—including malformed external genitalia (hypospadias), undescended testis (cryptorchidism), spermatogenic defects, and testis germ cell cancer—result from alterations in the in utero and perinatal hormonal milieu, causing disruption of sensitive male reproductive tract developmental events. The Sertoli cell is a possible target cell initiating this developmental disruption [41]. As discussed in Section V.D, phthalates are ubiquitous environmental contaminants and endocrine-disrupting chemicals. Rats exposed to di-(n-butyl)phthalate during a critical in utero window of male reproductive tract development have altered gene expression and a number of reproductive tract abnormalities associated with testicular dysgenesis syndrome, including underdeveloped or absent reproductive organs, hypospadias, cryptorchidism, and decreased sperm production [42, 43]. Sertoli cells clearly are important as potential targets for disruption by endocrine active compounds. Both estrogens and thyroid hormone are key modulators of Sertoli cell proliferation, which normally occurs during a fixed developmental window encompassing late in utero gestation to about 2 weeks postnatal in the rodent [41]. The goitrogen, 6-propyl-2-thiouracil (PTU), induces neonatal hypothyroidism when administered to lactating dams. Recovery from this condition results in testicular enlargement and also leads to an increase in sperm production in rats [44, 45] and in mice [46]. The testicular enlargement observed in PTU-treated rats is attributed to both an increase in the seminiferous tubule components and interstitial components of the testis [47]. Neonatal hypothyroidism prolongs the period of maturation and proliferation of neonatal Sertoli cells, causing an increase in the number of Sertoli cells [47, 48]. In adulthood, after recovery, the number of germ cells [47] and Leydig cells [49] also is increased. Such an effect is induced only if PTU treatment is initiated during the neonatal period (days 4–8) when Sertoli cells are proliferating most rapidly [50]. Importantly, many toxicants affect thyroid function and thyroid hormone production and potentially impact Sertoli cell proliferation during this sensitive time period. A variety of estrogenic compounds have been investigated for their effects on Sertoli cell proliferation. Gestational and lactational exposure of rats and mice to the powerful synthetic estrogen diethylstilbestrol results in reduced Sertoli cell numbers, impaired spermatogenesis, and small testes in adulthood [51]. Such perinatal estrogenic exposures result in persistent alterations in gene expression, hormone secretion, and seminiferous tubule fluid dynamics [52].

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IV. MANIFESTATIONS By definition, toxicants producing early and selective injury to the Sertoli cell following in vivo exposure are classified as Sertoli cell toxicants. Specific injury to Sertoli cells often is revealed through characteristic alterations in the testis. The goal of this section is to illustrate and describe the testicular manifestations that occur secondarily to Sertoli cell injury. Simple, yet powerful, morphological and biochemical indices are widely used as tools to distinguish Sertoli cell toxicants from other testicular toxicants. These classical approaches continue to provide critical data concerning the pathogenesis of toxicant-induced testicular injury. In addition, a number of newly described molecular and cellular experimental approaches are being employed to provide a more sophisticated and dynamic understanding of the etiological events that account for the mechanisms of Sertoli cell injury.

A. Vacuolation Vacuolation is a common and easily identified morphological alteration produced by a variety of xenobiotics at different times during the evolution of a testicular injury [53–56]. Vacuolation is also the earliest morphological sign of testicular injury and the cardinal response seen with many Sertoli cell toxicants. At the light microscopic level, two rather distinct forms of basal Sertoli cell vacuolation are observed, characterized by the presence of either multiple, small vacuoles or a single, large vacuole. By electron microscopy, both patterns of vacuolations result from swelling and coalescence of intracytoplasmic membrane-bound organelles, such as endoplasmic reticulum and vesicles [8, 57]. The presence of multiple, small vacuoles in the basal Sertoli cell cytoplasm is a prominent feature of the early response to phthalate exposure in young rats [8] and accompanies displacement of spermatogonia toward the lumen (Fig. 20.2). The occurrence of single, large vacuoles within the basal cytoplasm of the Sertoli cell is described in normal adult rat seminiferous epithelium in stages associated with meiosis [57]; however, the incidence and character of vacuolation are altered by exposure to certain Sertoli cell toxicants. Following 2,5-hexanedione exposure, both the number and size of Sertoli cell vacuoles observed in a testicular cross section are increased manyfold [57–59] and multiple vacuoles often are observed within the same seminiferous tubule (Fig. 20.3). Multiple testicular toxicants produce vacuolation of the seminiferous epithelium, including indenopyridines [60], nitrofurazone [61], and dibromoacetic acid [62], helping to identify these compounds as likely Sertoli cell toxicants.

FIGURE 20.2 Phthalate-induced vacuolation of the Sertoli cell cytoplasm. Upper panel: The normal appearance of seminiferous tubules from a 28-day-old rat (×280). Lower panel: This testis cross section is from a 28-day-old rat treated by gavage with 2.2 g/kg of di-(n-pentyl)phthalate and then euthanized 6 hr later (×280). The spermatogonia are displaced toward the lumen due to vacuolation and expansion of the basal Sertoli cell cytoplasm (stars). (Note: Unless otherwise indicated, all photomicrographs in this chapter are of Formalin-fixed testes embedded in glycol methacrylate, sectioned at 2.0–2.5 μm, and stained with periodic acid–Schiff’s reagent and hematoxylin.) (Reproduced with permission of Cache River Press.)

B. Apical Sloughing and Shedding Apical sloughing is defined as toxicant-induced detachment of fragments of the seminiferous epithelium leading to the appearance of aggregates of cellular material in the lumen. Sloughed cellular aggregates contain both germ cells and whole or fragmented Sertoli cells. Intratesticular injection of colchicine produces extensive apical sloughing [16]. In its most dramatic form, colchicine-induced apical sloughing is seen as cleavage of the seminiferous epithelium between cohorts of germ cells (Fig. 20.4). Electron microscopy shows that the sloughed seminiferous epithelium following colchicine exposure consists of clusters of germ cells adherent to fragments of membrane-bound apical Sertoli cell cytoplasm [16]. Possibly occurring as a separate process, shedding of germ cells into the lumen is defined as release of germ cells from the seminiferous

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FIGURE 20.3 This stage XII–XIII seminiferous tubule illustrates the multiple, large, basal Sertoli cell vacuoles (V) seen with 2,5-hexanedione exposure. A young adult rat was treated with 1% 2,5-hexanedione in the drinking water for 25 days (×235). (Reproduced with permission of Cache River Press.)

epithelium without attached fragments of Sertoli cells. In most instances, detailed ultrastructural studies are not performed to distinguish apical sloughing from shedding; however, many Sertoli cell toxicants, including 2,5-hexanedione [59], 1,3-dinitrobenzene [63], and the phthalates [8, 9], result in the appearance of germ cells in the lumen. A useful approach to detecting apical sloughing or shedding is the appearance of immature germ cells in the epididymis [64, 65]. As a word of caution, note that germ cell detachment from the seminiferous epithelium is probably a common artifact in the normal testis that is sectioned or manipulated prior to fixation [66]. Determining true germ cell detachment requires gentle tissue processing and appropriate controls.

C. Sertoli Cell Apoptosis Apoptosis is a term used to describe a distinctive form of cell death characterized by specific morphological and biochemical alterations that ultimately lead to the efficient elimination of the cell without instigating an inflammatory response. Very few reports of Sertoli cell apoptosis exist in the literature. In general, Sertoli cell apoptosis is limited largely to a period in testicular development in which the Sertoli cells are actively dividing by mitosis. In many species, Sertoli cells proliferate throughout embryonic and early postnatal testicular development. In rodents, Sertoli

FIGURE 20.4 Intratesticular colchicine injection (40 mg/testis) results in apical sloughing due to cleavage (asterisks) between spermatocyte and spermatid cohorts in this stage IV seminiferous tubule. This photomicrograph was taken 3 hr after injection (×420). (Reproduced with permission of Cache River Press.)

cell mitosis occurs until postnatal day 15–20 [67–69]. Exposure of 3- or 4-day-old rats, but not adult rats, to 5 Gy of ionizing radiation induces apoptosis of Sertoli cells, which peaks between 4 and 8 hr after exposure [70]. Sertoli cell cultures prepared from 10-day-old rats undergo apoptosis if they are maintained in suspension and not allowed to adhere to the culture dish substratum [71]. Nonylphenol, a biodegradation product of the nonionic detergent nonylphenol polyethoxylate, induces apoptosis in vitro of Sertoli cells prepared from 20-day-old Sprague-Dawley rats [72]. Collectively, these data demonstrate that proliferating Sertoli cells exhibit an increased apoptotic sensitivity following injury compared to quiescent Sertoli cells. However, more recent reports utilizing Bcl-w deficient mice indicate that Sertoli cell apoptosis also

Chapter 20 Sertoli Cell Toxicants

occurs in adult testis [73]. Bcl-w is a member of the anti-apoptotic Bcl-2 family of proteins. Male mice deficient in the expression of Bcl-w show a progressive loss of both Sertoli and germ cells of the testis [74]. The loss of Sertoli cells by apoptosis in the testes of Bcl-w deficient mice occurs between postnatal days 20 and 34, a time period immediately after the cells exit the proliferative phase [74]. The mechanism by which the antiapoptotic Bcl-2 family members mediate their effect is through binding and inactivation of the pro-apoptotic Bcl-2 family members [75, 76]. Bcl-w mediates the survival of postmitotic Sertoli cells via inhibition of the pro-apoptotic Bcl-2 family member Bax [73]. The observation of Sertoli cell apoptosis in adult Bcl-w–deficient mice indicates that even adult Sertoli cells undergo apoptosis if critical survival pathways are disrupted.

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a sensitive germ cell population [18]. Labeling fragmented DNA using terminal deoxynucleotidyl trans-ferasemediated digoxigenin-UTP nick end labeling (TUNEL) easily detects germ cell apoptosis (Fig. 20.5). In general, spermatogonia are the most resistant germ cell population following exposures to Sertoli cell toxicants, although prolonged Sertoli cell dysfunction often leads to extensive spermatogonial apoptosis [59, 78]. Postexposure fertility studies provide additional information about sensitive germ cell populations by pinpointing periods of reduced fertilizing ability, which reflect germ cell depletion [79]. As reported for 1,3-dinitrobenzene, quantitatively testing the fertilizing capacity of spermatozoa removed from the cauda epididymis enhances the sensitivity of a postexposure fertility study [80].

D. Germ Cell Apoptosis

E. Decreased Seminiferous Tubule Fluid Secretion

Apoptosis of testicular germ cells occurs in the testis under physiological conditions and increases after physical or chemical-induced testicular injury (for a review, see [77]). As an endpoint, germ cell apoptosis is easy to detect and quantify and provides information of potential mechanistic relevance, such as the germ cell developmental stage that is most sensitive to toxicant-induced Sertoli cell dysfunction. Studies demonstrating Sertoli cell-mediated selective apoptosis of stage XII–II pachytene spermatocytes following acute phthalate exposure illustrate the usefulness of detecting

One final common pathway by which Sertoli cell dysfunction results in germ cell apoptosis is decreased seminiferous tubule fluid secretion. The normal Sertoli cell–dependent secretion of seminiferous tubule fluid is measured easily by ligating the efferent ducts [27]. After approximately 16 hr, the increased weight of a ligated testis is compared with its contralateral unligated testis to determine the rate of seminiferous tubule fluid secretion. This technique demonstrates decreased seminiferous tubule fluid secretion following phthalate [10], 2,5-hexanedione [33], and indenopyridine

FIGURE 20.5 Identification of apoptotic germ cells. TUNEL detects DNA fragmentation characteristic of cells undergoing apoptosis. In thin paraffin cross sections, TUNEL-positive germ cells are identified easily by prominent staining (arrows).

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[60] exposure. Because seminiferous tubule fluid secretion is such a fundamental property of Sertoli cells and because the efferent duct technique is simple, straightforward and easily quantified, this measurement is useful in the investigation of any suspected Sertoli cell toxicant. The combination of morphological assessment and efferent duct ligation during a time course of exposure may provide mechanistic information, such as the observation that bulk germ cell apoptosis immediately follows failure of seminiferous tubule fluid secretion during 2,5-hexanedione exposure [33]. A refinement of the in vivo ligation technique for measuring seminiferous tubule fluid secretion uses intact seminiferous tubules in vitro (Fig. 20.6). After placing an oil droplet within the lumen, video microscopy is used to follow its movement, a reflection of the amount of seminiferous tubule fluid formed [32]. With this technique, seminiferous tubule fluid formation is energy dependent, requires intact microtubules, and is sensitive to brefeldin A, an inhibitor of the intracellular membrane transport system [32]. This in vitro technique was used to document decreased seminiferous tubule fluid formation following 2,5-hexanedione exposure [32] and exposure to carbendazim, the active metabolite of the fungicide benomyl [81].

F. Changes in Distribution, Quantity, or Biochemical Properties of Testicular Components Clues to the mechanism of action of a Sertoli cell toxicant may come from studying the early alterations in distribution, quantity, or biochemical properties of

testicular components. In this section, selected toxicantinduced alterations in testicular components are described that demonstrate both the potential power and limitations of these approaches: changes that are relatively easy to observe and quantitate may be mechanistically irrelevant or significant, a distinction that can be difficult to make without extensive additional experimentation. For example, the loss of a coherent microtubule organization within Sertoli cells following intratesticular colchicine injection is easily appreciated by immunostaining a testicular cross section with antitubulin antibody: the normal spoke-like arrangement of microtubules in the supranuclear Sertoli cell cytoplasm is obviously disrupted by colchicine exposure (Fig. 20.7). Because colchicine is so selective in its action on microtubules, a pathogenetic sequence involving colchicine-induced depolymerization of microtubules followed by apical sloughing and further degeneration of the seminiferous epithelium is both likely and logical [16]. For agents with nonspecific reactivity or for which target sites are unknown, the mechanistic interpretation of observed testicular alterations is more difficult. 2,5-Hexanedione exposure alters the assembly properties of testicular tubulin [58, 82]. The presence of a decreased time to achieve the maximal velocity of assembly and more rapid assembly characterizes the microtubule assembly altera-tions associated with 2,5-hexanedione exposure. The observed 2,5-hexanedione-induced alterations in microtubule assembly may be of mechanistic significance because they occur before other signs of injury and, therefore, could represent a critical pathogenetic step [33, 34, 59].

FIGURE 20.6 Measurement of seminiferous tubule fluid secretion via the luminal oil droplet procedure. Seminiferous tubules of defined stage are dissected from rat testes and mounted in a perfusion chamber bathed with Krebs-Henseleit bicarbonate buffer (pH 7.4, 32°C) as previously described [32]. An oil droplet is microinjected into the tubule, and the rate of movement of the oil droplet along the tubule is measured as an indirect indicator of seminiferous tubule fluid production. (A) A screen capture from a video recording of the microscope field of view showing an oil droplet in an isolated seminiferous tubule soon after its introduction. (B) A display of the same tubule and droplet after 30 min. The drop has moved 6.3 μm/min. Bar = 500 μm. (Reproduced with permission of Cache River Press.)

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exposure most likely represents loss of this cell population via sloughing and apoptosis [86]. Another approach is to analyze alterations in enzymes and proteins that are produced by specific cell types within the testis as markers of both cell injury and shifts in cell populations after toxicant exposure. Many of these markers were first employed to measure the changes in testicular cell populations that occur during development [87, 88]. During the early phases of exposure to a Sertoli cell toxicant, one might expect that Sertoli cell–associated enzymes, such as β-glucuronidase and γ-glutamyl transpeptidase, would decrease in activity due to functional deficits of this target cell type. Later in the time course of exposure, the activities of Sertoli cell enzymes might increase because of germ cell depletion and the relative enhancement of the Sertoli cell enzyme contribution to the testicular protein extract. Just such a biphasic pattern of enzyme activity has been reported during exposure to 2,5-hexanedione [89]. Other marker enzymes studied following exposure to Sertoli cell toxicants generally alter their activities in a manner predicted by the dynamics of their cell populations of origin [89–91]; however, for many of these enzyme and protein markers, definitive histochemical studies remain to be performed to verify the specificity of their localization. Increasingly, mRNA expression analysis is used to examine cell-specific effects during toxicant exposure. For example, changes in Sertoli cell–expressed genes following exposure to 2-methoxyacetic acid suggest that Sertoli cells may be important direct targets of this testicular toxicant [92, 93].

G. Interstitial Release of Sertoli Cell Proteins FIGURE 20.7 Comparison of immunohistochemical staining for tubulin in control testis and colchicine-treated testis. Upper panel: The normal seminiferous tubule has an intense, well-organized, spoke-like arrangement of tubulin staining within Sertoli cells (×190). Lower panel: The immunoreactive tubulin in the colchicinetreated testis is less intense and dispersed throughout the Sertoli cell cytoplasm (×180). Colchicine (40 mg) was administered by intratesticular injection 3 hr before the rat was euthanized. The tissue was prepared and stained using a monoclonal antitubulin antibody and the immunoperoxidase technique as previously described [33]. (Reproduced with permission of Cache River Press.)

The element zinc is required for normal spermatogenesis [83], and rapid zinc depletion was initially suspected of being mechanistically involved in the pathogenesis of phthalate-induced testicular injury [84]. However, because zinc localizes predominantly to round spermatids [85], zinc depletion after phthalate

Proteins synthesized by Sertoli cells and selectively secreted into the seminiferous tubule fluid are released into the interstitium after toxicant exposure. The interstitial fluid concentration of inhibin increases severalfold within 1 day of a single exposure of adult rats to 25 mg/kg 1,3-dinitrobenzene [94]. The serum concentration of androgen binding protein doubles shortly after a single exposure of young rats to 2.0 g/kg di-(n-pentyl)phthalate and remains elevated for 3 weeks after dosing [95]. Interestingly, a single high dose of di-(n-pentyl)phthalate in young rats produces a transient acute inflammatory infiltrate in the testicular interstitium [8]. The acute inflammatory infiltrate, consisting of polymorphonuclear leukocytes, peaks within hours of di-(n-pentyl)phthalate exposure and is mostly absent 1 day after exposure [8]. Presumably, the phthalateinduced Sertoli cell injury produces interstitial release of chemotactic factors from the seminiferous tubules; such a release generates the inflammatory response. Multiple mechanisms for systemic release of Sertoli cell-produced

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proteins of the seminiferous tubule fluid must be considered, including breakdown of the blood–testis barrier, breaches of the integrity of downstream structures such as the epididymis and vas deferens, loss of polarity in the Sertoli cell transport and secretion pathways, and altered communication among testicular cells [95].

H. Decreased or Increased Sertoli Cell Number Developmental hormonal disruptions that affect Sertoli cells profoundly alter the number of Sertoli cells per testis in the adult. The most thoroughly studied example of this phenomenon is early postnatal thyroid hormone deficiency in the rat, which results in a delayed and protracted proliferation of Sertoli cells, ultimately causing a marked increase in the total number of Sertoli cells and increased testis size in the adult [47]. On the other hand, exposure of rodents to estrogenic compounds perinatally results in decreased numbers of Sertoli cells and small testes in adults [52]. The morphometric techniques needed to determine Sertoli cell numbers are developed to the point that this is a readily performed assay [96–98].

I. Experimental Approaches to Studying Sertoli Cell Dysfunction 1. In Vitro Techniques Working with Sertoli cells isolated from young animals and cultured in an artificial environment has distinct limitations [99]. Because any compound is toxic by appropriate manipulation of dose and time of exposure in culture, many “Sertoli cell toxicants” are described in vitro that have no relevance to the whole animal; thus, the definitional requirement was established at the outset of this chapter that Sertoli cell toxicants be selective for the Sertoli cell in an in vivo model. Once in vivo studies identify the Sertoli cell as the site of toxicity, in vitro approaches are extremely powerful in furthering our mechanistic understanding of Sertoli cell toxicants by simplifying the confounders of metabolism and multiple cell types inherent with in vivo studies. Thus, investigations of active metabolites [100] and mechanisms of action [35, 101] are advanced by in vitro techniques. In vitro approaches also are used for screening purposes, an approach best reserved for the study of congeners of known Sertoli cell toxicants [26, 94, 102–104].

Using this technique, testicular tissue from immature primates is placed subdermally in mice where the primate testicular tissue undergoes differentiation. One of the fascinating outcomes of this technique is that the transplanted primate testicular tissue progresses to maturity with a timing consistent with normal mouse development of approximately 35 days. Therefore, this transplantation technique allows an examination of primate Sertoli cell maturation in a reduced time frame. This technique is touted for its ability to generate functional sperm from the germ line ex vivo and new strategies for preserving fertility in young men undergoing chemotherapy. Moreover, application of this technique for the evaluation of toxicant-induced disruption of Sertoli cell maturation in primates may prove useful. 3. Germ Cell Transplants The ability to transplant spermatogonial germ cells into a recipient testis was developed through the pioneering work of Brinster in 1994 [106, 107]. In the technique, donor spermatogonial germ cells are transplanted into the seminiferous tubule lumen of infertile recipient animals where the cells migrate to the epithelial basement membrane and reestablish spermatogenesis. Since the original description of this technique, investigators have utilized this experimental method for the investigation of germ cell biology and testis physiology, explored its potential application as a male infertility therapy, and investigated it as an effective strategy for producing transgenic animals (for reviews, see [108, 109]). This technique may be useful for defining the cellular site of toxicant action in the testis. For many testicular toxicants, the primary cellular site of action is difficult to determine because both Sertoli cell and germ cell toxicants result in germ cell apoptosis. Germ cell transplantation into a toxicant-injured testis could reveal if the environment within the seminiferous tubule, produced by the Sertoli cell, is capable of supporting functional spermatogenesis indicating that Sertoli cells were not injured by the toxicant. Experiments using busulfan, a chemotherapeutic agent that results in the elimination of germ cells directly, have shown that Sertoli cells from treated young mice are capable of supporting spermatogonial transplants [106–108]. However, in adult mice the supportive environment and/or the effects of busulfan are different since transplantation into busulfan-treated adult mice is inefficient [110]. These examples suggest the use of this technique in assessing the cellular site of toxicant action.

2. Subcutaneous Testicular Xenotransplants

4. Sertoli Cell Transplants

The ectopic transplantation of testicular tissue from various species into mice has been described [105].

The testis has long been regarded as an “immune privileged site.” Numerous studies employ the

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transplantation of testicular material or isolated Sertoli cells into other organs in order to study the mechanisms of testis immune privilege. The exact mechanisms that explain this immunoprotection are not well understood but may be explained, in part, by the expression of the pro-apoptotic death receptor ligand, FasL, by Sertoli cells [111, 112]. With the advent of germ cell transplantation techniques and their possible utility as a therapeutic strategy for infertility, investigators have directed their attention to the transplantation of Sertoli cells into the testis to restore spermatogenesis in mice with dysfunctional Sertoli cells [113]. Proliferating Sertoli cells isolated from young mice are transplanted successfully into the testes of mice with dysfunctional Sertoli cells (e.g., Steel/Steel dickie mice) and establish a microenvironment within the seminiferous tubule that supports functional spermatogenesis [113]. As with germ cell transplants, the transplantation of Sertoli cells into a testis after toxicant-induced injury may aid in the determination of the cellular site of toxicant action.

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1. Chemistry and Metabolism Benomyl is converted rapidly to carbendazim with the release of n-butylisocyanate (Fig. 20.8). Subsequently, carbendazim is hydroxylated on its aromatic ring to yield 5-hydroxybenzimidazole [114]. Because of its antimitotic activity resulting from inhibition of microtubule function, carbendazim is being explored as a cancer chemotherapeutic agent with renewed interest in its pharmacokinetic profile [115–117]. When combined with past studies of carbendazim metabolism and distribution [118–120], a clear picture emerges. Carbendazim is distributed rapidly and widely among tissues following both intravenous and oral exposures, and urinary excretion is the primary route of elimination. 2. General Toxicity Carbendazim induces liver neoplasms in male and female mice. Upon exposure, carbendazim does not produce significant amounts of DNA damage [121];

V. LITERATURE REVIEW This section provides an in-depth appreciation of the chemistry and sources, metabolism and pharmacokinetics, toxicological and pathological manifestations, and mechanistic concepts for four established chemical types of Sertoli cell toxicants: carbendazim, γ-diketones, nitroaromatics, and phthalates. These environmental and occupational xenobiotics fulfill the aforementioned definitional criteria for Sertoli cell toxicants and have a large broad-based research literature to review. Additional compounds are discussed in a Section V.E, entitled Other Toxicants. These other agents consist of suspected Sertoli cell toxicants for which limited or questionable literature is available and compounds that are systemically cytotoxic but produce interesting alterations in Sertoli cells.

A. Carbendazim Carbendazim is the primary metabolite of the widely used fungicide benomyl. The cellular target for carbendazim is the microtubule; nonmammalian microtubule function is in general more sensitive to disruption by carbendazim than mammalian microtubule function. However, among mammalian organ systems, the testis has a particular sensitivity to disruption by carbendazim. During the last decade or so, Sertoli cell microtubules were the focus of mechanistic research explaining the molecular basis for carbendaziminduced testicular injury.

FIGURE 20.8 The fungicide benomyl is rapidly converted to the ultimate toxicant, carbendazim, releasing n-butylisocyanate. Hydroxylation on the aromatic ring of carbendazim to yield 5-hydroxybenzimidazole is a common pathway of further metabolism.

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however, carbendazim does produce kinetichorepositive micronuclei by an aneuploidogenic mechanism [122]. Recently, carbendazim was used by the Environmental Protection Agency’s Office of Pesticide Programs as a model compound for a mode of action evaluation of the association between aneuploidogens and carcinogenicity [123]. The major conclusions from this evaluation are that carbendazim (1) is not a mutagen, (2) is an aneuploidogen, (3) acts by a nonlinear dose–response mechanism, and (4) likely produces liver tumors in mice by inducing aneuploidy, a nonmutagenic mode of action. Because of its microtubule-disrupting properties, carbendazim is used as a model compound to interfere with microtubule-dependent meiotic events in the oocyte, resulting in very early pregnancy loss [124]. The mechanism of very early pregnancy loss relates to carbendazim-induced aneuploidy in oocytes and subsequent zygotic arrest or early developmental failure, rather than an effect on fertilization per se [125, 126]. An alteration in oocyte meiotic spindle microtubule stability and spindle pole integrity is the molecular basis for this effect [127]. Carbendazim also produces embryo lethality and teratogenic effects following in utero exposure in animals [128–130]. 3. Testicular Toxicity: General Features The testicular toxicity of benomyl is explained, in large part, by its metabolism to the ultimate toxicant, carbendazim [131]. Early studies demonstrate that subchronic or chronic exposures of animals to benomyl or carbendazim produce selective alterations in the male reproductive tract, including decreased fertility, decreased epididymal sperm counts, decreased caudal epididymal weights, and histopathological changes in the testis characteristic of hypospermatogenesis, without affecting copulatory behavior [132–135]. Depending on the dose and extent of exposure, the testicular effects are reversible [133] or irreversible [134]. Relatively high dose exposures produce a profound testicular atrophy characterized by an increased intratesticular testosterone concentration with normal serum testosterone levels and elevated levels of FSH and luteinizing hormone (LH) [136, 137]. Because of its selectivity for the testis, carbendazim became one of the standard compounds to use in evaluating protocols for the determination of toxicant-induced testicular injury [138–141]. A study of the age dependence of benomyl-induced testicular toxicity shows that prepubertal rats are less sensitive than adults [142], an observation explained by both a decreased distribution of carbendazim to the testes of young animals and an inherent resistance of the immature seminiferous

epithelium to the adverse effects of exposure [143]. The observed alterations in prepubertal rat testis include abnormal spermatids with nuclear enlargement or binucleation [144]. A study of the relative sensitivity of rat and mouse to carbendazim-induced testicular injury suggests an inherent resistance of the murine seminiferous epithelium to the adverse effects of microtubule disruption, including carbendazim exposure [145]. 4. Testicular Toxicity: Pathology Detailed histopathological analysis of the testicular effects of carbendazim exposure is facilitated by the use of single high doses. Oral dosing up to 800 mg/kg produces an early increase in testis weight followed by a dose-dependent decrease in testis weight, spermatid numbers, morphologically normal cauda epididymal sperm, and mean seminiferous tubule diameter at later time points [146–148]. Of particular interest are changes in the efferent ducts characterized by severe inflammation, epithelial disorganization, fibrosis, sperm granulomas, and occlusions. Carbendazim exposure produces direct effects on dividing germ cells and on spermiogenesis. Early after exposure, meiotic spermatocytes (rat stage XIV) die, leading to defects in the round spermatid population [149]. By electron microscopy, carbendaziminduced spermatid nuclear abnormalities occur in rat stages IX–XI, including nuclear distortions, nuclear invaginations, and abnormal positioning of the nuclear envelope [150]. In addition, discontinuous, multiple granular and fragmentary acrosomes are observed in rat stages VII–XI, poorly formed or absent ectoplasmic specializations are seen in Sertoli cells adjacent to acrosome-deficient spermatids, and irregular positioning of manchette microtubules is observed in step 9 through 11 spermatids [150]. A detailed ultrastructural and immunohistochemical analysis suggests that the acrosomal abnormalities result from impaired acrosomal development during early spermiogenesis with perturbation of the supply of material from the Golgi apparatus to the acrosome [151]. In support of the histopathological analyses, flow cytometry of testicular cells isolated following carbendazim exposure shows an altered ratio of germ cell types, abnormal sperm head morphology, and an altered sperm chromatin structure [152]. After carbendazim exposure, round spermatids have an increased frequency of aneuploidy, and epididymal sperm have an increased frequency of diploidy [153, 154]. Following single high-dose exposures to carbendazim, significant alterations are observed in Sertoli cell structure. Sloughing of the apical seminiferous

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epithelium is observed in all rat stages except stages III–IV [155]. Cleavage occurs in the apical Sertoli cell cytoplasm, resulting in a cluster of sloughed spermatids with intact ectoplasmic specializations attached to a portion of Sertoli cell cytoplasm. Intact microtubules are observed in the sloughed Sertoli cell cytoplasm but are reduced in the body of the Sertoli cell where aggregates of mitochondria occur along with swollen cisternae of endoplasmic reticulum [155]. Whereas the body of the Sertoli cell is columnar in control testis, it is conically shaped following carbendazim exposure [156]. Associated with this shape change are changes in the orientation and location of Sertoli cell nuclei [156]. A detailed ultrastructural examination shows that microtubules are reduced markedly in the bodies of Sertoli cells undergoing sloughing, indicating that the stage-specificity of sloughing is due to the stage-specific susceptibility of microtubules to carbendazim disruption [157]. Carbendazim-induced effects on the efferent ducts have a long-term impact on testicular function. In particular, the efferent ducts play an important role in hormone-dependent fluid resorption [158]. After a single high-dose exposure (400 mg/kg), reproductive tracts containing occluded efferent ducts are associated with decreased testicular weight and seminiferous tubular atrophy [146]. Postexposure, the number of patent efferent ducts is associated with the number of normal seminiferous tubules and testes with totally occluded efferent ducts have the most numerous atrophic seminiferous tubules, indicating that the long-term deleterious effects of carbendazim depend largely on efferent duct potency [159].

inhibitor of microtubule polymerization, for comparison, both carbendazim and colchicine cause extensive depolymerization of testicular microtubules, although the total tubulin content remains relatively constant after carbendazim exposure. Tyrosinated α-tubulin, a subpopulation of relatively labile tubulin subunits, is decreased in the microtubule pool after carbendazim exposure [167]. Based on studies of testicular microtubule polymerization in the presence and absence of microtubule associated proteins, carbendazim interferes with the initial events of microtubule polymerization, including GTP binding, an effect modulated by microtubule associated proteins [168]. 6. Future Directions The studies performed to date with carbendaziminduced testicular toxicity are very important from two perspectives. First, the observation of persistent testicular pathology ensuing from downstream occlusion of efferent ducts by fragments of sloughed seminiferous epithelium underscores the close interactions between these contiguous male reproductive tract structures in health and disease. Second, the focus on microtubules as a molecular target for testicular injury reinforces the importance and selective susceptibility of this cytoskeletal element in the Sertoli cell. These themes of the interplay between contiguous structures in the male reproductive tract and Sertoli cell microtubules as a molecular target for toxicant-induced testicular injury will continue as important research areas in the future.

B. γ-Diketones 5. Testicular Toxicity: Mechanisms The histopathological observation of sloughing of germ cells with attached fragments of Sertoli cells that have an altered microtubule cytoskeleton focuses attention on Sertoli cell microtubules as the molecular target for carbendazim-induced testicular injury. This focus on microtubules as a target originates with early studies in which carbendazim was used as a probe of nonmammalian eukaryotic microtubule structure and function [160]. The benzimidazoles bind to the β-tubulin subunit of the αβ-tubulin heterodimer, inhibiting microtubule polymerization [161, 162]. Mutations in the fungal β-tubulin benzimidazole binding site confer either resistance or supersensitivity to the microtubule polymerization inhibitory effects of carbendazim [163–166]. Detailed studies of the effects of carbendazim on polymerization of rat testis microtubules [167, 168] followed early in vitro studies of its effects on mammalian brain tubulin [169]. Using colchicine, a well-studied

Both n-hexane and methyl n-butyl ketone (2-hexanone) are commonly used solvents that have caused many episodes of clinically apparent peripheral polyneuropathy following both occupational exposure [170] and inhalation abuse [118]. Animal studies identify the γ-diketone, 2,5-hexanedione, as the nhexane and methyl n-butyl ketone metabolite responsible for neurotoxicity [171]. Interestingly, testicular atrophy occurs “before clinical signs of neuropathy and light microscopic evidence of axonal swelling” in rats exposed to 2,5-hexanedione [172]. However, no human cases of testicular injury by 2,5-hexanedione or its metabolic precursors are identified, probably because transient infertility and small gonads would be the only signs of male reproductive toxicity. This review, though focused on animal studies of 2,5-hexanedioneinduced testicular injury, draws on the vast research and clinical literature concerning γ-diketone neurotoxicity to discuss potential common mechanisms

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of selective injury of these two organ systems and to point to future directions of research. Two past reviews provide extensive background information concerning γ-diketone neurotoxicity [173, 174], and two recent reviews summarize the current state of knowledge about 2,5-hexanedione-induced testicular injury [175, 176]. 1. Chemistry, Metabolism, and Pharmacokinetics n-Hexane is converted into an interrelated family of aliphatic ketones and alcohols (Fig. 20.9) by oxidation at the ω-1 position by the hepatic microsomal mixed function oxidase system [177, 178]. Neurotoxicity depends on the ability of a precursor compound to be metabolized into a sufficiently high concentration of the active γ-diketone [171, 179–181]. Indeed, the neurotoxic potency of aliphatic precursors relates directly to the serum concentrations of 2,5-hexanedione that they generate [171]. Testicular injury also occurs after exposure to n-hexane and all of the aliphatic ketone and alcohol precursors of 2,5-hexanedione [171, 182]. 2,5-Hexanedione is widely distributed to tissues throughout the body and has a long in vivo residence time with a serum half-life of 1.5−3 hr and a clearance time of 16−24 hr, depending on species, dose, and route of exposure [171, 174, 177, 178, 183]. γ-Diketones and primary amines combine to form pyrroles in a reaction called the Paal-Knorr condensation [184, 185]. During γ-diketone exposure, pyrroles accumulate on tissue proteins [186]. Indeed, pyrrole formation is a required step in the induction of both the testicular and the nervous system injuries [187–189]. Following tissue pyrrole formation, subsequent oxidation and cross-linking events are required for the expression of neurotoxicity [190, 191]. A study of a series of aliphatic diketones supports the requirement for a crosslinking event underlying testicular injury as well [183]. 2. Testicular Toxicity: General Features The testicular toxicity of 2,5-hexanedione is evaluated after subcutaneous injection [192], gavage [171], inclusion in the drinking water [172], and intraperitoneal injection [177]. Because of its simplicity and reproducibility, inclusion of 2,5-hexanedione in the drinking water, usually as a 1% solution, is the most common method of exposure. Both the length and dose rate of exposure influence the extent of 2,5-hexanedioneinduced testicular injury [172, 193]. Rats are commonly exposed as young adults weighing approximately 200 g [59, 89]. Though neurotoxicity is caused by many different nonhexacarbon γ-diketone congeners and precursors,

FIGURE 20.9 n-Hexane is metabolized at the ω-1 position by the hepatic microsomal mixed function oxidase system into a series of related alcohols and ketones culminating in the ultimate testicular toxicant 2,5-hexanedione. 2,5-Hexanedione combines with a primary amine, such as a protein lysyl ε-amine, to form an heterocyclic aromatic compound called a pyrrole.

including 3,4-dimethyl-2,5-hexanedione [194, 195], 2,5-heptanedione [196], 3,6-octanedione [197], and 5-nonanone [180, 198], the spectrum of testicular sensitivity to these compounds is poorly defined. Of these compounds, 3,4-dimethyl-2,5-hexanedione lacks testicular toxicity at the doses and time points

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examined [82, 194, 195], whereas 5-nonanone produces testicular injury [198]. Because 2,5-hexanedione is neurotoxic, the concomitantly occurring testicular injury could be secondary to disruption of the hypothalamic–pituitary–gonadal axis. However, a primary testicular site of 2,5-hexanedione action is supported by the observation that plasma gonadotropin levels become elevated only after testicular alterations occur, whereas testosterone levels remain normal during and after exposure [78, 89]. 3. Testicular Toxicity: Pathology One unusual feature of 2,5-hexanedione-induced testicular injury is the long latency between initiation of a high-dose exposure and the appearance of histopathological alterations. In fact, a 2-week exposure to 1% 2,5-hexanedione in the drinking water is sufficient to produce a profound testicular injury even though morphological alterations in the seminiferous epithelium do not appear until 1−2 weeks after the exposure is terminated [59]. This delay in toxic effect is explained, in part, by the complex chemistry of the interaction of γ-diketones with tissue nucleophiles that requires sequential steps of pyrrole formation, oxidation, and cross-linking [183, 187–191]. The temporal sequence of biochemical and morphological alterations in Sertoli cells and germ cells are studied in detail for a 5- to 6-week exposure of young adult rats to 1% 2,5-hexanedione in the drinking water. Two weeks after initiating exposure, alterations are observed in the polymerization of purified rat testis tubulin, which is predominantly of Sertoli cell origin [34]. Three weeks after beginning exposure, the Sertoli cell cytoskeletal components of the tubulobulbar processes lose their immunoreactivity [33, 199], and the Sertoli cell-associated enzymes β-glucuronidase and γ-glutamyl transpeptidase are decreased significantly in activity [89]. The first morphological evidence of testicular injury occurs 3−4 weeks after initiating exposure and consists of the appearance of large, basal vacuoles in the Sertoli cell cytoplasm in stages XII, XIII, XIV, and I [57, 59] (Fig. 20.3). By electron microscopy, these vacuoles are described as swollen endoplasmic reticulum [57]. Hypercurvature of the elongate spermatid nucleus accompanies Sertoli cell vacuolation, but other major germ cell changes begin after cessation of seminiferous tubule fluid secretion [33]. Seminiferous tubule fluid secretion, a Sertoli cell function, is inhibited abruptly about 2 days after the appearance of Sertoli cell vacuolation [33]. About this same time, 4 weeks after beginning exposure, an alteration in the distribution of seminiferous tubule stages is observed with a decrease in stages VII and XIII and an increase

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in stages III, V, and VI [57]. In addition, 2,5-hexanedione exposure reduces the stage-dependent progression of labeled germ cells and increases the duration of the cycle of the seminiferous epithelium [200]. In the process of germ cell loss, the initial abnormalities consist of nuclear margination in round spermatids and giant cell formation [33]. A careful analysis of this process by morphology, DNA fragmentation detected by gel electrophoresis, and TUNEL staining identifies the germ cell loss as due to apoptosis [201]. TUNEL is quite sensitive and indicates an increased rate of germ cell apoptosis within 2 weeks of initiating 2,5-hexanedione exposure [201]. A potentially important pathway in signaling germ cell death following Sertoli cell injury is the Fas system [112, 202]. In this signaling system, FasL expressed by Sertoli cells activates Fas receptor expressed on germ cells to initiate apoptosis [112, 202]. Progressive loss of spermatocytes and spermatogonia and sloughing of seminiferous epithelium continues for several weeks after the initial germ cell manifestations of injury [59]. Alterations in lipid metabolism are noted 6 weeks after beginning exposure, a time of nearly complete germ cell depletion [203]. Finally, 12 weeks after beginning exposure, a profound testicular atrophy is present with only 1% of seminiferous tubules containing differentiating germ cells [59]. Germ cell repopulation of the seminiferous epithelium occurs in only a minority of exposed rats despite the presence of residual spermatogonia [59, 78]. The state of persistent testicular atrophy reached long after 2,5-hexanedione exposure ceased has been the focus of intense investigation. Interestingly, residual stem germ cells are present in the atrophic testis and proliferate with kinetics compatible with the cycle of the seminiferous epithelium [204]. A quantitative evaluation of the residual germ cell population combined with modeling of spermatogonial proliferation suggests that the stem cell population is reduced in size and that proliferation occurs to a limited extent before the germ cells die by apoptosis [205]. Because the calculated block occurs at the level of type A3 spermatogonia, the developmental point at which the stem cell factor/c-kit paracrine signaling system is required for germ cell survival, intratesticular infusions of soluble stem cell factor were tested as a therapeutic intervention [206]. The infusion of soluble stem cell factor causes a statistically significant increase in the number of labeled germ cells, indicating that stem cell factor promotes survival and/or stimulates proliferation of the residual germ cell population in 2,5-hexanedione– induced atrophic testes [206]. The focus on the stem cell factor/c-kit pathway is encouraged further by the observation of an altered ratio of soluble-to-membrane

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associated stem factor in atrophic testes [207]. The use of hormonal regimens that suppress testosterone synthesis reverses the 2,5-hexanedione-induced testicular atrophy and corrects the abnormal stem cell factor ratio [207]. 4. Testicular Toxicity: Mechanism The axonal lesions induced by 2,5-hexanedione are characterized by giant accumulations of neurofilaments as well as alterations in both fast and slow axonal transport [192, 208, 209]. Because of the dramatic morphological appearance of the neurofilamentous accumulations, attention focuses on alterations of these intermediate filaments as the mechanism of axonal injury [184, 186]. Experiments in cell culture model systems support an action of 2,5-hexanedione on intermediate filaments [210–212]. On the other hand, because axonal and Sertoli cell microtubules share common morphological and biochemical features [30, 213–216], targeting of microtubules by 2,5-hexanedione is proposed as one possible underlying mechanism of toxicity [58]. Indeed, a dramatic alteration is observed in the assembly of testis tubulin purified from 2,5-hexanedione exposed rats [34, 58, 193]. The assembly alteration is “taxollike” and characterized by precocious nucleation and abnormally rapid assembly. Tubulin treated with 2,5-hexanedione in vitro displays the same assembly alterations as that derived from the testes of 2,5hexanedione–treated rats [82]. The modified tubulin demonstrates altered control responses to the usual modulators of assembly, such as temperature, calcium, and guanosine 5’-triphosphate [82]. Because of enhanced nucleation of assembly, shorter and more abundant microtubules are formed. A cross-linking event that modifies the structure of the α-tubulin subunit may explain the 2,5-hexanedione-induced proassembly effect [217, 218]. Using sea urchin zygotes as an in vivo model system, mitotic spindle abnormalities are observed after microinjection of 2,5-hexanedionetreated tubulin [219]. Three separate in vivo approaches consistently support an association between altered testicular microtubule assembly and testicular injury: (1) the use of congeners with differential testicular and microtubule effects [58], (2) the determination of microtubule alterations throughout a time course of 2,5-hexanedione exposure and recovery [59], and (3) examination of the relationship between dose rate, testicular effects, and microtubule assembly alterations [193]. A hypothesis that ties together the morphological alterations produced by 2,5-hexanedione in the testis and the underlying microtubule abnormalities has

been proposed [33, 58, 213]. In this model, dysfunctional microtubules lead to the appearance of Sertoli cell vacuoles and, over time, to progressive failure of the Golgi-dependent secretory pathway, causing failure of seminiferous tubule fluid secretion and germ cell apoptosis. The rationale for this model rests with the known association of microtubules with the endoplasmic reticulum [220] and the Golgi complex [221, 222] and the known dynamic role of microtubules in protein secretion [31]. A substantial effort was expended to test this hypothesis, beginning with a better characterization of the components of secretory apparatus in the testis. Using an in vitro system [32], both colchicine, a classic inhibitor of microtubules, and brefeldin A, an inhibitor of protein secretion, significantly decrease seminiferous tubule fluid formation. Immunohistochemical analysis shows that the microtubule-dependent motor, cytoplasmic dynein (a minus-end-directed microtubule motor), is present in Sertoli cells during all stages of spermatogenesis with a peak in the apical cytoplasm during stages IX–XIV [223]. Kinesin (a plus-end-directed microtubule motor) is found in manchettes [223] and localizes to the Sertoli cell trans-Golgi network [224]. After 2,5-hexanedione exposure, apical cytoplasmic dynein immunofluorescence in Sertoli cells decreases and Golgi and kinesin immunostaining is disrupted [225]. Sertoli cells isolated from rat testes following 2,5-hexanedione exposure possess an atypical spindle shape and long cytoplasmic processes [226]. The cytoplasmic processes are rich in microtubules and vimentin, whereas actin stress fibers decrease in density in the Sertoli cells isolated from 2,5-hexanedione exposed rats compared to controls [226]. Interestingly, in vitro microtubule-dependent motility assays using kinesin as the motor show that 2,5-hexanedione–treated microtubules have reduced transport rates compared to control microtubules [227]. These results suggest that 2,5-hexanedione treatment directly alters the structure or conformation of the microtubule, producing both enhanced microtubule assembly and slowed kinesin-dependent transport. An alternative, molecular approach to testing the microtubule-dependent hypothesis of 2,5-hexanedione– induced testicular injury was explored recently. γ-Tubulin is a microtubule-nucleating protein that disrupts microtubule networks in cells following overexpression. Following rete injection and back-perfusion of the seminiferous tubules, adenoviruses selectively infect Sertoli cells [228]. By using adenoviruses engineered to express γ-tubulin, the effect of a selective alteration in Sertoli cell microtubule function was evaluated [229, 230]. γ-Tubulin overexpression causes alterations in Sertoli cell microtubule networks, and overexpressed γ-tubulin–enhanced green fluorescent

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fusion protein localizes to sites of elongate spermatid attachment in the seminiferous epithelium [229]. This Sertoli cell overexpression of γ-tubulin markedly alters the organization of the seminiferous epithelium, resulting in retention of elongate spermatids and residual bodies, and germ cell apoptosis [230]. This molecular approach to microtubule disruption highlights the important role of this Sertoli cell cytoskeletal element in germ cell viability. 5. Future Directions The hypothesized pathogenetic sequence for 2,5-hexanedione-induced testicular injury involves cross-linking of tubulin producing an altered microtubule assembly that inhibits microtubule-dependent transport in Sertoli cells, causing decreased seminiferous tubule fluid formation and a failure to support germ cells so that they die by apoptosis. Although much work has contributed to the development and testing of this hypothesis, many issues remain to be addressed, including (1) structural studies to determine the biochemical basis for altered microtubule assembly, (2) a molecular explanation for the selective sensitivity of the nervous system and testis to 2,5-hexanedione, (3) the details of how Sertoli cell microtubule-dependent transport supports seminiferous tubule fluid formation, and (4) a better molecular understanding of the machinery that regulates the germ cell apoptotic response to Sertoli cell injury.

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3-nitroacetanilide, are identified in in vitro cocultures of Sertoli and germ cells exposed to 1,3-dinitrobenzene [7, 241]. Based on previous studies of nitrobenzene reduction [242, 243], a general scheme for the metabolism of 1,3-dinitrobenzene is proposed [244] that features a variety of reduced intermediates including the nitroxyl anion radical, 3-nitrosonitrobenzene, and 3-nitrophenylhydroxylamine (Fig. 20.10). Using germ cell detachment from rat Sertoli–germ cell cocultures as a marker of toxicity, 3-nitrosonitrobenzene is a more potent toxicant than 1,3-dinitrobenzene [244], while neither 3-nitroaniline nor 3-nitroacetanilide are toxic [63, 241]. Further, the toxicity of both 1,3dinitrobenzene and 3-nitrosonitrobenzene is enhanced by decreasing intracellular thiol levels and ameliorated by scavenging reactive intermediates [244]. This evidence implicates reduction to 3-nitrosonitrobenzene as a required step for testicular toxicity and suggests that further reductive steps to highly reactive intermediates are involved in the injury [244].

C. Nitroaromatics Nitroaromatics are produced in large amounts and represent a diverse chemical class of compounds. The most common and obvious toxicity associated with exposure to nitroaromatics is methemoglobinemia and anemia. In addition, various nitroaromatics produce testicular injury, including nitrobenzene [231], 1,3-dinitrobenzene [7], 2,4-dinitrotoluene [232, 233], 2,4-diaminotoluene [234], the pesticide dinoseb [235], and 2,4,6-trinitrotoluene [236]. This review will focus on 1,3-dinitrobenzene, a commonly used intermediate in the chemical industry for the synthesis of pesticides, dyes, and explosives and the nitroaromatic studied in greatest depth with regard to testicular injury. 1. Chemistry and Metabolism Past in vivo studies of 1,3-dinitrobenzene [237, 238] and nitrobenzene [239, 240] identify nitro reductions, ring hydroxylations, and conjugation reactions as major steps in metabolism that vary between species and strains. Two reduced metabolites, 3-nitroaniline and

FIGURE 20.10 The nitro group of 1,3-dinitrobenzene is metabolized through a series of reduced intermediates to an amine which is then acetylated. Both 1,3-dinitrobenzene and 3-nitrosonitrobenzene are Sertoli cell toxicants in vitro, whereas neither 3-nitroaniline nor 3-nitroacetanilide is toxic. (Modified from Cave and Foster [244].) (Reproduced with permission of Cache River Press.)

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Blood levels and cumulative exposures of 1,3-dinitrobenzene that produce testicular injury in rats produce no testicular toxicity in hamsters, an observation that is explained by the much lower levels of 3-nitroalaniline in hamsters compared to rats [245, 246]. Even though 3-nitroaniline is not itself a testicular toxicant [247], precursors of this metabolite that accumulate in the rat, but not in the hamster, are candidate testicular toxicants [245]. However, recent investigations provide a more compelling hypothesis to explain species differences in response to 1,3-dinitrobenzene. These investigations indicate that 1,3-dinitrobenzene metabolism occurs in the mitochondria and that the specific level of mitochondrial glutathione and its participation in the metabolism/detoxification of 1,3-dinitrobenzene are the most important determinants of testicular toxicity [248]. These studies show that mitochondrial preparations of in vitro seminiferous tubule cultures prepared from hamsters have a greater capacity for reductively metabolizing 1,3-dinitrobenzene to the active metabolite compared to rats [248, 249]. However, levels of cellular glutathione and ATP are not depleted as quickly in hamster preparations as those from rats [248]. Recent studies indicate that the 1,3-dinitrobenzene metabolites, nitrosonitrobenzene and nitrophenylhydroxylamine, react with glutathione nonenzymatically and result in decreased levels of mitochondrial glutathione [250, 251]. Taken together, these findings indicate the critical role of metabolism of 1,3-dinitrobenzene for its toxicity and that the capacity of the glutathione-mediated detoxification reaction underlies the mechanistic basis for the differential species sensitivity to testicular toxicity. 2. Testicular Toxicity Subchronic exposure of rats to 1,3-dinitrobenzene in the drinking water [252] or by gavage [253] results in testicular atrophy and infertility at doses that produce few, if any, systemic symptoms. Another model appropriate for histopathological and mechanistic studies is acute high-dose exposure. A single exposure to 25 mg/kg 1,3-dinitrobenzene results in Sertoli cell vacuolation in stages VIII and XI first observed 12–24 hr after exposure, a time when germ cells appear normal [7, 64]. This initial Sertoli cell injury is followed by apoptosis of late pachytene spermatocytes, primarily of stages VI–VIII and IX–XIII, 24–48 hr after exposure [254, 255]. Microscopic examination of the caput epididymis [64] and flow cytometric analysis of cells in caput epididymal fluid [65] show the presence of immature germ cells presumably shed from the seminiferous epithelium. Hormonal analyses confirm the direct

toxicity of 1,3-dinitrobenzene for the testis without primary involvement of the hypothalamic–pituitary– testicular axis [256]. Studies of fertility, sperm parameters and testicular and epididymal histopathology performed after a single 48 mg/kg dose of 1,3-dinitrobenzene demonstrate an early and profound degeneration of spermatocytes in stages VII through XIII and effects on stage I through V spermatids [79, 257, 258]. In vitro fertilization using epididymal sperm of rats indicates that germ cells of stages III, IV, XII, and XIV are the most sensitive to disruption by acute 1,3-dinitrobenzene exposure [80]. Species differences are noted with rats more sensitive than mice after a single high-dose exposure to 1,3-dinitrobenzene [65]. In addition, young mice and rats are more resistant than older animals to testicular injury. This sensitivity of older rodents to dinitrobenzene is explained by differences in metabolism or pharmacokinetic differences [245, 259–261]. However, the metabolic/pharmacodynamic differences observed do not explain fully the increased sensitivity of adult rodents to dinitrobenzene [248]. Further support for the Sertoli cell as the initial target of 1,3-dinitrobenzene testicular toxicity derives from in vitro studies. Germ cell detachment, a presumptive in vitro marker of Sertoli cell toxicity, occurs with noncytotoxic doses of 1,3-dinitrobenzene and its reduced metabolite 3-nitrosonitrobenzene [7, 241, 244]. In addition, 1,3-dinitrobenzene elevates the level of lactate and pyruvate secreted by Sertoli cells in culture by an FSH-independent mechanism [26, 262]. This increase in lactate and pyruvate is not observed with 1,2- or 1,4-dinitrobenzene, congeners of 1,3-dinitrobenzene that lack testicular toxicity [26]. Recent findings also indicate that agents such as 1,3-dinitrobenzene act directly on germ cells. The Fas signaling system plays a crucial role in germ cell apoptosis resulting from phthalate-induced Sertoli cell injury [263]. It is hypothesized that activation of the Fas system is a common response for inducing germ cell apoptosis after Sertoli cell injury [263]. However, Fasor FasL-deficient mice (lprcg or gld mice, respectively) display an increased sensitivity to 1,3-dinitro-benzene– induced testicular germ cell apoptosis [264]. These findings are contrary to the prediction that the Fas system is required for the initiation of germ cell death after toxicant-induced Sertoli cell injury. It is possible that the explanation for the ability of 1,3-dinitrobenzene to induce spermatocyte apoptosis in these mice is rooted in the potential direct interaction of 1,3-dinitrobenzene on germ cells. However, the fact that the Fas-deficient mice are more sensitive than the wild-type mice indicates that the loss of function of Fas or FasL sensitizes the mice to 1,3-dinitrobenzene–induced germ cell apoptosis.

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These observations indicate that inhibition of Fas or FasL function leads to cellular changes in the testis, making spermatocytes more sensitive to the effects of 1,3-dinitrobenzene. These are the same germ cell subtypes that undergo apoptosis in the wild-type mice. Therefore, the profiles of germ cell subtypes undergoing 1,3-dinitrobenzene-induced apoptosis are not changing. Thus, 1,3-dinitrobenzene–induced testicular injury may be a compilation of direct actions on both Sertoli and germ cells. 3. Future Directions The work performed to date on 1,3-dinitrobenzene testicular toxicity provides the necessary basic knowledge of dose and route of exposure for future mechanistic studies. Both in vivo and in vitro results support the Sertoli cell as the initial target of injury within the testis, whereas studies of 1,3-dinitrobenzene metabolism identify 3-nitrosonitrobenzene as a proximate toxic metabolite. However, germ cells also are influenced directly by 1,3-dinitrobenzene exposure [264], and the participation of direct germ cell injury by this agent requires further investigation. Redox cycling of reactive metabolites of 1,3-dinitrobenzene is proposed as the underlying chemical basis to testicular injury [265]. The ability of diethyl maleate to exacerbate and of ascorbate and cysteamine to ameliorate in vitro toxicity supports a free-radical–mediated mechanism of injury. These agents could be used as potential probes into the fundamental biochemical processes that modulate 1,3-dinitrobenzene–induced Sertoli cell dysfunction and germ cell loss.

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testicular effects is unclear, and this review focuses on postnatal phthalate injury, because evidence points to the Sertoli cell as the initial target in that model. Several comprehensive reviews provide greater detail concerning the chemistry and diverse biological effects of these compounds [268–270]. 1. Chemistry and Sources Phthalates, also known as phthalic acid esters, are commonly used as plasticizers to impart flexibility to plastic containers and wrappings. The toxic phthalates of concern are usually diesters of phthalic acid, the ortho form of benzenedicarboxylic acid. The best studied and most abundantly produced phthalic acid ester is di-(2-ethylhexyl)phthalate (Fig. 20.11). Di-(2-ethylhexyl)phthalate is a colorless liquid with a high boiling point, a low vapor pressure, and an extremely low water solubility. Because the phthalate plasticizers are not covalently bound within the final product, these compounds are removed gradually from a plastic matrix by lipophilic agents. Thus, environmental distribution of phthalates is widespread [271]. Exposure to di-(2-ethylhexyl)phthalate within the general human population is estimated at 3–30 μg/kg/day, and exposure is ubiquitous [272, 273]. Patients receiving frequent transfusions are exposed to relatively high concentrations of phthalates leaching out of polyvinylchloride medical devices [274]; exposures are particularly high for neonates requiring multiple medical interventions, reaching levels greater or equal to 3.3 mg/kg/day [275, 276].

D. Phthalates The phthalates are the most completely characterized chemical class of Sertoli cell toxicants. In the first study of phthalate-induced testicular injury, dietary exposure of rats to 0.9 or 1.9 g/kg daily of di-(2-ethylhexyl) phthalate for 90 days produced histopathological changes limited to the testis and described as “tubular atrophy and degeneration resembling senile changes” [266]. Since then, various aspects of phthalate toxicity have been investigated including testicular injury, peroxisome proliferation in the liver, hepatocarcinogenesis, and hypolipidemia. This section will examine a selection of those studies, with an emphasis on findings pertinent to understanding the potential mechanisms of phthalate-induced postnatal testicular toxicity in rats. Currently, much attention is devoted to phthalate effects on male reproductive tract development [267]. At this time, the cellular target producing fetal phthalate

FIGURE 20.11 Di-(2-ethylhexyl)phthalate is activated by hydrolysis to the ultimate testicular toxicant mono-(2-ethylhexyl)phthalate. (Reproduced with permission of Cache River Press.)

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2. Metabolism and Pharmacokinetics Phthalate exposure occurs predominantly by the ingestion of diesters. A combination of in vivo and in vitro studies provides strong evidence that the active toxic metabolite of di-(2-ethylhexyl)phthalate is the monoester derivative, mono-(2-ethylhexyl)phthalate (Fig. 20.11). In vivo, mono-(2-ethylhexyl)phthalate is a more potent testicular toxicant than the corresponding diester [277]; several other metabolites, including 2-ethylhexanol, lack testicular toxicity [278]. Studies using testicular cells in culture show specific alterations in structure and function following exposure to low concentrations of mono-(2-ethylhexyl)phthalate that are not produced by di-(2-ethylhexyl)phthalate, 2-ethylhexanol, or other selected polar metabolites [26, 102, 278]. These findings, added to the demonstration that enriched cultures of Sertoli cells and Leydig cells are essentially incapable of further metabolism of mono-(2-ethylhexyl)phthalate [100], implicate phthalic acid monoesters as the ultimate toxic metabolites. The phthalate diesters undergo hydrolytic cleavage to the corresponding monoesters and alcohols in the small intestine and liver [279–281]. The relative toxicity of diester phthalate congeners depends on the in vivo generation of monoester phthalate metabolites and their inherent toxicity. Therefore, di-(n-pentyl)phthalate is more potent in vivo than di-(2-ethylhexyl)phthalate [282], probably because of differential rates of intestinal hydrolysis to the corresponding active monoesters [280]. Mono-(2-ethylhexyl)phthalate is the most potent monoester congener in an in vitro toxicity assay [102]. After diester administration, mono-(2-ethylhexyl)phthalate appears in the rat testis and persists there with a biological half-life of more than 2 days [283]. Most species excrete glucuronides of the parent compounds; however, rats metabolize the phthalate diester side chains to water-soluble derivatives [284, 285]. 3. Testicular Toxicity: General Features Several important variables are identified that modify testicular toxicity in phthalate-exposed animals, including species, age of the animal, and dosing regimen. Oral administration of di-(n-butyl)phthalate produces severe testicular atrophy in rats and guinea pigs, focal or no atrophy in mice, and no testicular injury in hamsters [286, 287]. Hamsters are insensitive to di-(2-ethylhexyl)phthalate but show testicular damage when administered mono-(2-ethylhexyl)phthalate, a difference in susceptibility explained by the relatively poor hydrolysis of diester to monoester by hamsters compared to rats [286]. Microscopic evidence of testicular injury also is reported following long-term exposure of the ferret to di-(2-ethylhexyl)phthalate [288].

Young rats are more susceptible to phthalateinduced testicular injury than adults. Four-week-old rats exposed by gavage to 2.8 g/kg of di-(2-ethylhexyl)phthalate for 10 days show a decrease in testis weight and uniform atrophy of the seminiferous tubules, whereas 10-week-old rats show normal testis weight and 5–50% affected tubules; the testes of 15-week-old rats are unaffected by this exposure [289]. Lesions are produced in mature rats by phthalate exposure; however, the extent of the injury is limited [9]. Neonatal rats exposed to di-(2-ethylhexyl)phthalate show a transient decrease in Sertoli cell proliferation and a decrease in spermatid maturation 4 weeks after exposure but no consistent adverse effects on fertility or other reproductive parameters [86, 290]. One possible explanation for the greater susceptibility of young rats to oral exposure by phthalate diesters is enhanced absorption, resulting in greater total exposure to monoester [291]. Perhaps the most attractive dosing regimens for mechanistic studies are acute high-dose exposures of the most active phthalate congeners in young animals of a sensitive species. A single 2.2 g/kg dose of di-(n-pentyl) phthalate or 2.2 g/kg mono-(2-ethylhexyl)phthalate administered by gavage to young rats produces histopathological and functional alterations in Sertoli cells within hours [8, 18, 103]. On the other hand, longer exposures to lower doses of phthalates are most useful in assessing general reproductive effects [292]. 4. Testicular Toxicity: Pathology The Sertoli cell is identified as the primary cellular target for postnatal phthalate-induced testicular injury by (1) the early histopathological changes occurring in this cell type following both in vivo and in vitro exposure, (2) the characteristic and early alterations in Sertoli cell function and biochemistry, and (3) the rapid disruption of the Sertoli–germ cell physical interaction. Three hours after oral administration of di-(n-pentyl)phthalate to sexually immature rats, electron microscopy reveals extensive vacuolation in the basal endoplasmic reticulum of Sertoli cells in a small portion of seminiferous tubules [8]. By 6 hr, Sertoli cell vacuolation is seen in nearly all seminiferous tubules, whereas spermatocytes and spermatids show early degenerative changes. In addition, an acute interstitial inflammatory infiltrate is present. By 24 hr, the seminiferous epithelium is disorganized with shedding and degeneration of spermatocytes and spermatids and a resolving acute interstitial inflammatory response. Repeated dosing over several days produces seminiferous

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tubules populated by Sertoli cells and occasional spermatogonia, but otherwise depleted of germ cells. The interstitium contains normal-appearing Leydig cells and a negligible inflammatory infiltrate [8]. Treatment of the sexually mature rat results in similar histopathological alterations, which are limited to stages XI−XIV, I, and II [9]. In addition, ectoplasmic specializations between Sertoli cells and between Sertoli and germ cells are disrupted, with fragmentation or retraction of Sertoli cell processes and convolution of the plasma membranes. After in vitro exposure of Sertoli–germ cell cocultures to mono-(2-ethylhexyl)phthalate and mono-(n-pentyl)phthalate [293], Sertoli cells develop finger-like processes that invaginate into adjacent cells, actin filaments increase in density and are altered in distribution, and increased densities of ribosomes, smooth endoplasmic reticulum, and Golgi complex are seen. In sum, similar ultrastructural effects are observed in vivo in immature and mature rats and in vitro in cocultures of rat Sertoli and germ cells after phthalate exposure [293]. Additional support for the Sertoli cell as the phthalate target comes from studies examining Sertoli cell proliferation and vimentin intermediate filaments [18, 86, 290]. One-week-old animals exposed to 1 g/kg of di-(2-ethylhexyl)phthalate by gavage for 5 days show a 35% decrease in Sertoli cell number per seminiferous tubule relative to age-matched controls. The number of Sertoli cells present is similar both before (at age 6 days) and after (at age 11 days) treatment, indicating that phthalate exposure produces a specific arrest in the normal Sertoli cell proliferation that occurs at this developmental age [86]. Using 3-day-old rats exposed to di-(2-ethylhexyl)phthalate, a transient, dose-dependent decrease in Sertoli cell proliferation is observed at 24 hr of exposure with coincident formation of multinucleated germ cells [290]. Sertoli–germ cell cocultures from 2-day-old animals exhibit a similar inhibition of Sertoli cell proliferation following mono-(2-ethylhexyl)phthalate exposure [294]. When pubertal rats are given an oral gavage of 1 g/kg mono(2-ethylhexyl)phthalate, Sertoli cell vimentin intermediate filaments that normally extend for the Sertoli cell nucleus to sites of germ cell attachment collapse around the basal nucleus [17], and this collapse occurs prior to an increase in germ cell death [18]. One remarkable feature of phthalate-induced testicular injury is the rapid shutdown of seminiferous tubule fluid formation that occurs after exposure. Complete inhibition is observed when this Sertoli cell– dependent secretory process is evaluated by efferent duct ligation beginning 1 hr after exposure of immature rats to 2.2 g/kg of di-(n-pentyl)phthalate [10]. The cessation in Sertoli cell–dependent fluid production is

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accompanied by a block in androgen binding protein secretion. Other rapid alterations in Sertoli cell biochemistry are observed, including a decrement in mitochondrial succinic dehydrogenase activity 3 hr after oral administration of di-(n-pentyl)phthalate to sexually immature rats [295]. By 6 hr, a complete loss of mitochondrial succinic dehydrogenase activity is seen in Sertoli cells, whereas this enzymatic activity in germ cells is unaffected. Studies using Sertoli cells in culture confirm the occurrence of phthalate-induced alterations in biochemistry and secretion, including a striking increase in lactate production and a variable decrease in pyruvate production [26, 296, 297]. These initial observations are extended with a more detailed analysis of phthalate-induced abnormalities in the glycolytic pathway [262, 298]. Other findings include a mild decrease in cellular ATP levels [296], a decrease in mitochondrial succinic dehydrogenase activity that is dependent on the presence of phthalate in the assay [296], an inhibition in transferrin secretion that can be reversed by added FSH [299], and an inhibition of FSH-stimulated cAMP accumulation [35]. Oral administration of 2 g/kg mono-(2-ethylhexyl)phthalate to pubertal rats indicates that germ cells die by apoptosis, with significant apoptotic induction at 6 hr postexposure [18]. Curiously, induction of apoptosis is preceded by a decrease in germ cell apoptotic rate at 3 hr. Phthalate-induced apoptosis appears to involve Fas ligand–dependent signaling between Sertoli and germ cells. Fas receptor is expressed by germ cells and mono-(2-ethylhexyl)phthalate increases Sertoli cell expression of Fas ligand [112, 202]. In mice with a mutant form of Fas ligand unable to bind Fas receptor, phthalate-induced germ cell apoptosis is inhibited significantly [263]. However, germ cell apoptosis still occurs in these mutant mice, demonstrating the importance of other apoptotic mechanisms. In vitro toxicity studies with Sertoli–germ cell cocultures pioneered by Gray and Beamand and continued by others [102, 294] show that mono-(2-ethylhexyl)phthalate induces a concentration-dependent increase in the rate of germ cell detachment, a unique effect not produced by di-(2-ethylhexyl)phthalate or 2-ethylhexanol [102] or other selected polar metabolites [278]. Importantly, germ cell detachment from cocultures occurs at doses much lower than those that cause general cytotoxicity, suggesting that release is a specific consequence of exposure. 5. Testicular Toxicity: Mechanisms The rapid onset of phthalate-induced testicular injury suggests a relatively specific and simple mechanism of action. The six distinct mechanistic hypotheses

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proposed to explain postnatal phthalate-induced testicular injury implicate widely divergent aspects of testicular function: (1) zinc-dependent enzymatic activities, (2) hormonal status, (3) metabolic function, (4) FSH receptor-dependent pathways, (5) peroxisome proliferator-activated receptor–dependent pathways, and (6) Sertoli–germ cell adhesion. These hypotheses will be discussed in their historical order of development to provide a sense of their conceptual evolution with respect to general knowledge in the field. The observation that nutritional zinc deficiency results in testicular atrophy is basic to the hypothesis that the testicular toxicity of phthalates is mediated by zinc depletion [83]. Young rats administered di-(n-butyl)phthalate by gavage show testicular atrophy associated with increased urinary excretion of zinc and decreased testicular zinc [84]. Coadministration of zinc with phthalates either protects against [84] or has no effect on [300] the occurrence of testicular atrophy. Decreased zinc content in testis following phthalate exposure is species and congener specific and correlates with the extent of testicular injury [282, 286, 301–303]. X-ray microanalysis shows zinc deposits, which are limited to round spermatids, decreasing in abundance 6 hr after a single phthalate dose [85]. Three studies demonstrate the occurrence of phthalate-induced testicular injury in young rats without alterations in zinc content or with alterations in zinc occurring subsequent to germ cell apoptosis [86, 277, 304]. These data suggest that previous observations of loss of testicular zinc are not mechanistically important and may reflect shedding and apoptosis of spermatids. The mechanism by which dietary zinc deprivation produces testicular atrophy may be via a direct effect on testicular steroidogenesis [305, 306]. For this reason, numerous investigators have studied hormonal status after postnatal phthalate exposure. Serum concentrations of testosterone are reported as either decreased [307] or unchanged [308] after phthalate exposure, while testicular concentrations of testosterone are either elevated [302, 303] or decreased [287, 309]. Phthalates produce marked decreases in the testicular cytochrome P-450–dependent microsomal steroidogenic enzyme activities 17 α-hydroxylase and 17-20 lyase [310]. On the other hand, a single toxic dose of phthalate has no effect on plasma FSH levels [277], and simultaneous administration of testosterone or FSH to phthalate-treated animals does not ameliorate the testicular injury [289]. Further, the phthalate-induced testicular injury in adult rats is limited to stages XI–XIV, I, and II [9], whereas the stage most sensitive to testosterone deficiency is stage VII [311]. Unlike postnatal phthalate exposure, fetal exposure to di-(2-ethylhexyl)phthalate or di(n-butyl)phthalate via maternal oral

gavage produces a marked reduction in testosterone synthesis with subsequent effects on testosteronedependent tissues [312, 313]. Although the fetal phthalate exposure model produces multiple effects on the developing testis including changes in Leydig cell steroidogenesis, morphology, and proliferation as well as gonocyte morphology, the cellular target or mechanism of action is unclear [270]. The observation that hepatic mitochondria exposed to phthalates are functionally abnormal [314] provided the impetus to study Sertoli cell metabolism after phthalate treatment. Subtle alterations in mitochondrial function are observed, though no changes in Sertoli cell ATP levels occur in the first 8 hr after in vitro phthalate exposure [296]. Clear alterations in metabolism are observed in vitro, most notably a large increase in lactate production [26, 296, 297] due to stimulation of glycolysis or the pentose phosphate pathway [262, 298]. These findings demonstrate that phthalates cause metabolic abnormalities; however, the nature of the changes suggests that metabolic dysfunction is not the primary mechanism of Sertoli cell damage leading to germ cell loss. The observation that phthalate-induced testicular injury of adult rats is limited to stages XI–XIV, I, and II [295], those stages with the highest FSH responsiveness [315], prompted investigation of the FSH receptor and its associated signaling system as a molecular target. A further attraction of the FSH receptor system as a mechanistic target is its unique expression by Sertoli cells. Using cultured Sertoli cells, mono-(2-ethylhexyl)phthalate specifically inhibits FSH-stimulated accumulation of the second messenger cAMP [35, 316]. The FSH-stimulated inhibition of cAMP accumulation is partial, time dependent, dose dependent, independent of cAMP breakdown or adenylate cyclase inhibitory pathways, and FSH receptor signal transduction system dependent. FSH binding to Sertoli cell membranes is inhibited only after preincubation with mono(2-ethylhexyl)phthalate and is a consequence of altered affinity rather than a reduction in receptor number [317]. These results suggest that mono-(2-ethylhexyl)phthalate acts as a noncompetitive inhibitor of FSH signal transduction. The noncritical role played by FSH itself in adult testicular function and the delayed effect of phthalate on FSH binding to its receptor argues against the FSH receptor as an important target of toxicity [317, 318]. Because G-protein–coupled receptors exhibit extensive cross-regulation [319], the delayed and indirect nature of the phthalate effect of FSH receptor function suggests that another Sertoli cell G-protein–coupled receptor signaling pathway is an important phthalate target. Because phthalates are liver peroxisome proliferators, a mechanistic role for peroxisome proliferator-activated

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receptors (PPARs) in phthalate testicular toxicity has been studied. The three PPARs (α, β, and γ) are members of the nuclear hormone receptor family and regulate transcription of genes involved in lipid and carbohydrate metabolism [320]. Rodent testis produces all PPAR isoforms with expression of PPARα reported in Leydig cells, Sertoli cells, and spermatocytes; PPARβ in Leydig and Sertoli cells; and PPARγ in Sertoli cells [321, 322]. Using PPARα knockout mice, the liver effects of phthalate exposure are attributed to activation of this isoform [323, 324]. In contrast, PPARα-deficient mice develop testicular atrophy following chronic, high-dose (12,000 ppm in the diet) di(2-ethylhexyl)phthalate exposure [324]. Based on a qualitative analysis of seminiferous tubule lesion development, PPARα knockout mice show a delay in lesion onset [324], but this result may be due to unexplored variations in these mice such as changes in phthalate pharmacokinetics. These data indicate that PPARα activation is not mechanistically crucial for phthalate testicular toxicity; however, a vital role for PPARβ or γ remains possible. A hallmark of phthalate testicular injury is the separation of germ cells from Sertoli cells. In the pubertal testis, this separation is an early event during injury and manifest as an adluminal shedding of spermatocytes and round spermatids, suggesting that adhesion between Sertoli and germ cells is disrupted [8]. Similarly, phthalate exposure of Sertoli–germ cell cocultures produces extensive germ cell detachment [10, 294]. Although it is not known whether Sertoli–germ cell adhesion junctions are targeted directly, the collapse of junction-associated Sertoli cell vimentin intermediate filaments prior to an increase in germ cell apoptotic endpoints is indicative of early alterations in Sertoli– germ cell adhesion junctions [18]. Germ cells adhere selectively to Sertoli cells, and in model systems, intercellular adhesion is a prosurvival mechanism [4, 11]. These data suggest that phthalates target the unique aspects of Sertoli–germ cell adhesion, leading to germ cell detachment and apoptosis. Important unanswered questions in such a mechanism are the factors involved in Sertoli–germ cell adhesion and whether detachment precedes apoptotic induction. 6. Future Directions A satisfactory molecular explanation of the mechanism of phthalate-induced testicular injury must account for the known unique aspects of this injury: (1) the greater susceptibility of young rather than old animals, (2) the greater susceptibility of certain stages of the cycle of the seminiferous epithelium, (3) the fact that neonatal Sertoli cells are injured,

(4) the histopathological evidence of rapid injury to Sertoli cells including vacuolation and retraction of processes, (5) the occurrence of an acute inflammatory infiltrate soon after exposure, (6) the rapid shutdown of seminiferous tubule fluid formation, (7) the modulation of G-protein–mediated Sertoli cell signaling, (8) the alterations in vimentin intermediate filaments with detachment of germ cells, and (9) the downregulation of testosterone synthesis during gestational exposure. These unique characteristics suggest that phthalate exposure quickly alters a Sertoli cell–specific process already expressed by early postnatal Sertoli cells. Although unknown, the Sertoli cell may be a common target in both gestational and postnatal injuries. Examining the available data as a whole, an attractive hypothesis is that the phthalate target is associated with the Sertoli cell plasma membrane and expressed coincident with the developmental specification of Sertoli cells.

E. Other Toxicants 1. Cisplatin Cis-diammine dichloroplatinum (cisplatin) is a chemotherapeutic drug used for the treatment of solid tumors and is highly efficacious in the treatment of most germ cell-related malignancies. Chemotherapyassociated azoospermia in men [325] may result from a loss of stem spermatogonia. However, azoospermia and frequently permanent infertility occur despite the presence of spermatogonia [14, 326, 327]. There is considerable debate about the cellular target of cisplatin in the testis. While exposure to cisplatin, even at low doses, causes germ cell death, the Sertoli cell is widely believed to be the testicular target for cisplatin. Disruption of the blood–testis barrier in response to cisplatin exposure [326] has suggested Sertoli cell injury. Cisplatin-induced reduction in the production of inhibin B and transferrin from the Sertoli cell have also been frequently reported [14, 328–335]. In addition, the increased sodium and decreased potassium concentrations in seminiferous tubular fluid noted in treated animals indicate abnormal Sertoli cell secretory function [14]. Upregulation of Fas ligand on Sertoli cells is a hallmark of injury-associated reduction in Sertoli cell supportive capacity [112]. Recently, an upregulation in the level of FasL in response to subchronic exposure to cisplatin was observed; increased expression is observed even after recovery for two spermatogenic cycles (J. H. Richburg, unpublished observations). These observations suggest that the cisplatin primary cellular target is the Sertoli cell. However, cisplatin-induced disruption of the inter-Sertoli cell tight junctional

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complexes does not result in the expected systemic leakage of androgen binding protein [332], which is maintained at normal serum and intratesticular levels [14]. Further, cisplatin transiently reduces both serum and testicular testosterone levels [14, 336], apparently by inhibition of testicular cytochrome P-450– dependent steroid biosynthesis, and adversely affects the viability and proliferation of spermatogonia [337]. The apparent multiplicity of cell types involved indicates both a complex mode of action and the requirement of additional studies to dissect the contribution of Sertoli cell dysfunction to cisplatin-induced testicular injury. 2. Colchicine Colchicine specifically binds to tubulin and inhibits its polymerization into microtubules [338]. Medically, colchicine is used to treat chronic inflammatory disorders and produces a variety of systemic toxic effects [339, 340]. Therapeutic doses alter sperm production in humans, including induction of reversible azoospermia [341, 342]. Ultrastructural studies of the effects of colchicine on the testis demonstrate that, although axonemal and manchette microtubules are quite stable, both germ cell spindle and Sertoli cell microtubules are depolymerized readily by this agent [343, 344]. Because of its action on germ cell spindles, colchicine is used to study the effects of chemically induced aneuploidy [345, 346]. A detailed investigation of the effects of intratesticularly injected colchicine on Sertoli cell ultrastructure shows that microtubules in the Sertoli cell cytoplasm rapidly depolymerize [16]. Following colchicine injection, groups of germ cells with attached Sertoli cell cytoplasm appear in the lumen, a process called apical sloughing, presumably due to cleavage of the seminiferous epithelium between cohorts of germ cells. A more detailed evaluation of colchicine-induced apical sloughing identifies rat stages IX–XIV as most sensitive to sloughing, whereas stages VII–VIII exhibit the most dramatic alterations in tubulin immunostaining [347], suggesting that other factors, such as vimentin, are likely contributing to the structural integrity of the seminiferous epithelium. Given that sloughing is a major consequence of colchicine exposure, microtubules may be necessary for the support of the delicate Sertoli cell processes in the adluminal compartment [16]. Using the ground squirrel as a model, colchicine markedly decreases the number of Sertoli cell microtubules after both acute and subacute subcutaneous exposure [30]. In the absence of Sertoli cell microtubules, elongate spermatids develop abnormal acrosomes and fail to undergo apical migration, while the Sertoli cell smooth

endoplasmic reticulum accumulates basally [30]. These reports of the testicular effects of colchicine are consistent with the original proposal by Christensen [348] that “microtubules support the tenuous cytoplasmic extensions of the Sertoli cells, and may also produce the various movements that spermatids undergo in the epithelium over the course of spermiogenesis.” 3. Indazole-3-Carboxylic Acids Following the observation that 1-(4-chlorobenzyl)1H-indazole-3-carboxylic acid possesses significant antispermatogenic activity [349–352], a large series of halogenated 1-benzyl-1H-indazole-3-carboxylic acids were synthesized and tested for testis effects [353]. The halogenated 1-benzyl-1H-indazole-3-carboxylic acids with significant antispermatogenic activity that have been studied most intensely are (1) the original compound, 1-4(4-chlorobenzyl)-1H-indazole-3-carboxylic acid (AF 1312/TS); (2) 1-(2,4-dichlorobenzyl)-1H-indazole-3-carboxylic acid (AF 1890 or lonidamine); and (3) 1-(4-chloro-2-methylbenzyl-1H-indazole-3-carboxylic acid (AF 1923 or tolnidamine). In addition, more recent investigations focus on 1-(2,4-dichlorobenzyl)indazole-3-carbohydrazide (AF-2364) and 1-(2,4dichlorobenzyl)-indazole-3-acrylic acid (AF-2785) (reviewed in [354]). Within hours of exposure of rats, rabbits, dogs, or monkeys to a single dose of either AF 1312/TS or lonidamine, similar early alterations in Sertoli cells are observed, characterized by vacuolation of the cytoplasm and retraction of the apical cytoplasm with germ cell shedding [352, 355]. Because morphological changes are observed first in Sertoli cells, this cell is proposed as the target for antispermatogenic activity [352, 356]. The mechanisms by which halogenated 1-benzyl1H-indazole-3-carboxylic acids produce their testicular effects remain unknown. These compounds may act by perturbing Sertoli–germ cell adherens junctions, thus causing germ cell loss from the seminiferous epithelium (reviewed in [354]). Studies examining 1-(2,4-dichlorobenzyl)-indazole-3-carbohydrazide (AF-2364) and 1-(2,4-dichlorobenzyl)-indazole-3acrylic acid (AF-2785) show detachment of round and elongated spermatids into the seminiferous tubule lumen [357]. The disruption of the Sertoli–germ cell adherens junctions is indicated by the increased expression of testin, a marker whose expression correlates with the integrity of Sertoli–germ cell junctions [357, 358]. As determined by autoradiography, lonidamine gains access to the seminiferous epithelium [359]. Beginning 2 days after exposure to lonidamine, serum FSH becomes elevated and remains elevated while LH and testosterone do not change [360].

Chapter 20 Sertoli Cell Toxicants

Interestingly, serum androgen binding protein is elevated within hours of lonidamine exposure accompanied by a decrease in testicular and epididymal androgen binding protein [360]. This systemic release of androgen binding protein may be explained by alterations of the blood–testis barrier [15]. Studies of Sertoli cells in culture exposed to 1-benzyl1H-indazole-3-carboxylic acids demonstrate changes in morphology and secretion [361–363], and AF 1312/TS is very active in enhancing germ cell release from Sertoli–germ cell cocultures [102]. Finally, Sertoli cell mitochondria apparently function normally in the presence of these toxicants [364], although 1-benzyl-1H-indazole-3-carboxylic acids adversely affect mitochondrial function in germ cells and tumor cells [365, 366]. 4. Indenopyridine Developed initially as potential antihistamines some 25 years ago, indenopyridine derivatives elicit a toxic response that is confined to the testis [367]. Acute oral indenopyridine exposure produces a similar sequence of histopathological changes in dogs, mice, and rats. By 24 hr of dosing, seminiferous epithelium histopathology includes vacuolation of the basal epithelium (interpreted as within Sertoli cells), extensive apoptosis of spermatocytes and spermatogonia, and generation of multinucleated germ cells [60, 368, 369]. Evidence for the Sertoli cell as the initial target is suggested by the following data. Androgen status is not altered, and only mild elevations in serum FSH and LH are observed [60, 370]. On the other hand, indenopyridine adversely affects several Sertoli cell structural/functional parameters early after exposure, including seminiferous tubule fluid production, serum inhibin B levels, and vacuolation [60]. Depending on the dosing regimen and indenopyridine derivative, acute effects produce infertility with only Sertoli cells and occasional spermatogonia present in the seminiferous epithelium, and exposure consequences may be irreversible [368, 369]. 5. 2-Methoxyacetic Acid The commercial solvent 2-methoxyethanol (also known as ethylene glycol monomethyl ether) produces testicular degeneration in numerous species resulting from its metabolism by alcohol/aldehyde dehydrogenases to 2-methoxyacetic acid (MAA), the ultimate toxic metabolite [371–373]. Following an in vivo exposure, germ cells undergo apoptosis within 24 hr, with pachytene spermatocytes in stages XII to XIV being the most sensitive germ cell population [295, 374, 375]. This specific germ cell depletion, associated with mild

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morphological alterations in Sertoli cells, suggests that MAA directly targets germ cells. Such a conclusion is questioned by results from in vitro experiments. Ku and Chapin [376] show that MAA-induced germ cell degeneration seen in vivo is recapitulated with exposure of intact seminiferous tubules but not with exposure to either Sertoli–germ cell cocultures or isolated germ cells. These data suggest that Sertoli cells are affected directly by MAA, and this reasoning is supported by additional data as follows. Although seminiferous tubule fluid production is unaffected [374], toxicity is associated with increased plasma inhibin levels and decreased lactate production in vitro [94, 377]. Although androgen binding protein levels in rete testis fluid are not changed, plasma androgen binding protein levels increased [374, 378]. In addition, quantitative changes to Sertoli cell–expressed mRNAs including androgen binding protein [379], a phosphodiesterase and stress protein [93], and others [92] are observed early after MAA exposure. Finally, the stage-dependent distribution of androgen receptor protein and mRNA in Sertoli cells is altered after MAA exposure [379]. Collectively, these data show that Sertoli cells are altered by MAA exposure with kinetics approximating alterations to pachytene spermatocytes. Because an intact seminiferous tubule environment is important for toxicity, the MAA toxicity mechanism may involve Sertoli cells as a direct target. At the very least, the Sertoli cell is an important component in the toxicity mechanism. 6. Tri-o-Cresyl Phosphate Tri-o-cresyl phosphate is an organophosphorus ester compound used commercially as a plasticizer and lubricant. It is known to produce a delayed polyneuropathy following exposure [380]. Investigations in both rats and mice describe reduced fertility following tri-o-cresyl phosphate exposure with altered sperm motility and histopathological abnormalities in the testes [381, 382]. Tri-o-cresyl phosphate-induced testicular abnormalities also are observed in roosters [383] and chickens [384] and explored in detail in rats [385–387]. The initial testicular abnormality observed in rats is vacuolization of the Sertoli cell cytoplasm, identifying the Sertoli cell as a possible initial target of exposure. Sertoli cell vacuoles occur contemporaneously with an abnormal basal location of elongate spermatids, apparently due to Sertoli cell phagocytosis that precedes extensive degeneration of round spermatids and spermatocytes. No abnormalities are observed in serum, interstitial fluid testosterone, serum FSH, serum LH, or seminiferous tubule fluid production as assessed by efferent duct ligation [385].

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By electron microscopy, widespread dilation of the Sertoli cell smooth endoplasmic reticulum is observed [387]. The active metabolite of tri-o-cresyl phosphate, saligenin cyclic-o-tolyl phosphate, is present in high concentrations in the testis, probably due to selective Leydig cell metabolism [101]. It is proposed that saligenin cyclic-o-tolyl phosphate exerts its damaging effects by acting as an alkylating agent [388]. Sertoli cells in culture exposed to saligenin cyclic-o-tolyl phosphate display minor morphological alterations and an increased lactate production without alterations in pyruvate production, synthesis, or secretion of proteins, or the cAMP response to FSH stimulation [389]. Following in vivo exposure to tri-o-cresyl phosphate, striking decreases are detected in testicular nonspecific and neurotoxic esterase activities, enzyme activities that are also targeted in the nervous system [385, 386]. In both Leydig cell and Sertoli cell cultures, nonspecific esterase activity is inhibited by saligenin cyclic-o-tolyl phosphate [101, 389]. The high concentration of saligenin cyclic-o-tolyl phosphate within the testis likely explains the occurrence of organ-specific toxicity; however, the underlying mechanism of Sertoli cell dysfunction and potential targets, other than nonspecific esterase, remain unknown.

postspermatogonial germ cells is observed at all time points with a relatively constant population of 3−4 spermatogonia per 100 Sertoli cells in atrophic seminiferous tubules (Fig. 20.12). Because stem germ cells normally are present in approximately this number [390, 391], these results suggest that 2,5-hexanedione exposure induces an “irreversibly” injured testis containing residual stem germ cells [78]. Indeed, actively proliferating stem cells are found in the atrophic testes of 2,5-hexanedione-exposed rats [204]. Modeling of the frustrated spermatogenesis in these atrophic testes suggests that spermatogonia are destroyed by apoptosis, creating a block to further maturation, which explains the persistence of the atrophic state [205]. Interestingly, pharmacological interventions that suppress testosterone levels allow reestablishment of spermatogenesis in the 2,5-hexanedione-exposed atrophic testis [207]. The underlying mechanisms responsible for toxicantinduced induction of this block to spermatogonial

VI. REVERSIBILITY In this section, the issue of the reversibility of germ cell loss after exposure to Sertoli cell toxicants is reviewed briefly. The reversibility of a toxicantinduced testicular injury is an important issue from both scientific and societal viewpoints. Socially, it is important that fertility return after a toxic exposure, such as chemotherapy in cancer patients who want to start or enlarge their families. Scientifically, models of depopulation and repopulation provide one avenue to a better understanding of the underlying processes that control germ cell proliferation and differentiation. Sertoli cell toxicants can cause an apparently “irreversible” testicular atrophy, studied in greatest detail with 2,5-hexanedione. Even relatively short exposures, as little as 2 weeks of 1% 2,5-hexanedione in the drinking water, produce long-lasting testicular atrophy with many seminiferous tubules lacking differentiated germ cells 22 weeks later [59]. Interestingly, when these atrophic seminiferous tubules are examined closely, approximately 2 spermatogonia per 100 Sertoli cells are observed [59]. A more detailed evaluation of this phenomenon was undertaken in young adult rats exposed to 1% 2,5-hexanedione in the drinking water for 5 weeks with morphological assessment of the testes 27, 60, and 75 weeks after exposure [78]. A generalized loss of

FIGURE 20.12 Atrophic seminiferous tubules (T) contain spermatogonia 75 weeks after a 5-week exposure to 1% 2,5-hexanedione in the drinking water. Upper panel: A solitary spermatogonium is identified by its basal location within the seminiferous tubule, its limited and prominent cytoplasm, and its nuclear features (×990). Lower panel: In this seminiferous tubule, a pair of spermatogonia is connected by an intercellular bridge (arrow) (×870). Interstitial space (I). (Reproduced with permission of Cache River Press.)

Chapter 20 Sertoli Cell Toxicants

maturation and its release by hormonal manipulation are not understood. This topic of recovery from “irreversible” testicular injury is explored in greater detail elsewhere (see Chapter 23).

6.

VII. CONCLUSIONS The focus in the first edition of this chapter, written more than a decade ago, was on morphological and biochemical manifestations of Sertoli cell injury. Although these remain important approaches for identifying Sertoli cell toxicants, the power of molecular tools is now obvious. We are immersed in the “omics” revolution, and the application of genomics and proteomics methods to previously intractable problems has opened up new avenues of research. Despite these advances, the identification of the Sertoli cell as the cellular target of a testicular toxicant remains a process of combining various manifestations of an injury (morphological, biochemical, and molecular) that point to the Sertoli cell in the aggregate. Although we still do not know the molecular mechanisms of action for most of the toxicants discussed in this chapter, the rapid advances in the analytical techniques of molecular toxicology are promising. The excitement and energy of this new research era indicates that mechanistic investigations of Sertoli cell toxicants have passed from infancy to adolescence, and that the pace of progress and deeper understanding is accelerating.

7.

8.

9.

10. 11.

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14.

15.

Acknowledgment The authors would like to recognize the contributions of Dr. Pragati Sawhney for her critical review of this work and Jeanne Galbo for her excellent technical support. This work was supported by grants from the NIEHS/NIH (RO1 ES05033 awarded to K.B.; RO1 ES11632 awarded to K.J.J.; RO1 ES09145 awarded to J.H.R.), NIH Center Grant P30 ES07784 (J.H.R.), and NIH Training Grant ES07247 (J.H.R.).

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C H A P T E R

21 Conditions Affecting Sertoli Cells WAEL A. SALAMEH AND RONALD S. SWERDLOFF Division of Endocrinology, David Geffen School of Medicine, University of California at Los Angeles, Torrance, California

After birth in humans, two waves of Sertoli cell proliferation occur, the first at 18 months of life and then during early puberty. The final number of Sertoli cells determines the efficiency of spermatogenesis. During adulthood the Sertoli cell provides the nurturing milieu necessary for spermatogonial stem cells to commit to proliferation and differentiation in continuous waves of maturing male gametes. It is thus useful to classify the entities that affect Sertoli cell function depending on when they occur. Some genetic diseases affecting Sertoli cells in utero will predominantly interfere with the process of sexual differentiation. Other entities occurring early postnatally will affect Sertoli cell number and hence the efficiency of spermatogenesis. Finally, testicular exposure to noxious environmental stimuli in adulthood may lead to the inability of the Sertoli cells to nurse the male gametes in their maturational process and to various degrees of impairment in spermatogenesis and male factor infertility. In this chapter we comprehensively review the clinical disorders associated with perturbations in Sertoli cell function in each of these developmental stages.

I. INTRODUCTION II. SERTOLI CELL MARKERS AS PREDICTORS OF SERTOLI CELL HEALTH AND THERAPEUTIC OUTCOME III. GENETIC AND ENVIRONMENTAL DISORDERS AFFECTING DEVELOPMENT OF SERTOLI CELL FUNCTION IV. SERTOLI CELL ONLY SYNDROME V. SERTOLI CELL NEOPLASIA VI. GENETIC SYNDROMES WITH A SERTOLI CELL PHENOTYPE VII. NEONATAL, PUBERTAL, OR ACQUIRED ADULT CONDITIONS AFFECTING SERTOLI CELL FUNCTION VIII. SUMMARY References

I. INTRODUCTION Previous chapters have paved the way for us to understand the critical functions the Sertoli cell plays in testicular determination and subsequently in the initiation and maintenance of spermatogenesis. Our understanding of these functions is not complete, but there is consensus that this cell has different roles during different stages of male gonadal maturation. A timely recent review explores these functions in detail [1]. During embryonic life and after sex determination through expression of Sry occurs, the undifferentiated gonad becomes committed to a male fate. The Sertoli cells secrete Müllerian inhibiting substance (MIS) and the Müllerian ducts undergo regression. SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

II. SERTOLI CELL MARKERS AS PREDICTORS OF SERTOLI CELL HEALTH AND THERAPEUTIC OUTCOME Several Sertoli cell products have been touted as being clinically useful to indicate damage to the Sertoli cell and the probability of success for different therapeutic interventions aimed at improving infertility in men. Cardinal among those is inhibin B, a dimer

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whose subunits are secreted and assembled by the Sertoli cells. Inhibin B is predominantly responsible for inhibition of follicle-stimulating hormone (FSH) secretion by the pituitary. The availability of radioimmunoassay (RIA) for measuring this biologically active form and not its individual subunits has made possible its use as a marker for Sertoli cell function [2, 3]. The regulation of inhibin B α and β subunit secretion from the Sertoli cells is influenced by circulating FSH originating in the pituitary. In addition, it is regulated by other inhibins and activins secreted by the Leydig cells and as-yet unidentified factors from the meiotic and haploid germ cell (Fig. 21.1). The levels of circulating inhibin B and FSH change in a dynamic fashion from birth to adulthood, reflecting a changing relationship between these two hormones. In infant males during early puberty, an original positive correlation exists between these two hormones, possibly reflecting the role of FSH in stimulation of Sertoli cell proliferation in these two stages of development (Fig. 21.2). At more advanced stages of puberty and adulthood, this correlation becomes a negative one, reflecting the traditional role of inhibin as a suppressor of FSH secretion [2]. Low semen counts in the adult positively correlate with low inhibin B levels [3] (Fig. 21.3). Other markers of Sertoli cell function or maturity have been either used clinically [MIS, androgen receptor (AR), cytokeratin 18 (CK-18)] or used in animal models as a means of determining the impact of various interventions on Sertoli cell function [androgen-binding protein (ABP)]. Throughout the chapter, we will refer to the specific studies detailing the utility of these markers in the various conditions affecting Sertoli cell function.

III. GENETIC AND ENVIRONMENTAL DISORDERS AFFECTING DEVELOPMENT OF SERTOLI CELL FUNCTION A. Disorders of Sexual Differentiation 1. True Hermaphroditism and Mixed Gonadal Dysgenesis Disorders of sexual differentiation make up a complex set of entities that generate much controversy in terms of their classification and management. In this section we discuss these entities only as they relate to Sertoli cell function. Mixed gonadal dysgenesis (MGD) is characterized by various degrees of testicular differentiation ranging from immature testes to streak gonads. On the other hand, true hermaphroditism (TH) is characterized by the presence in either or both gonads of well-differentiated ovarian and testicular

tissues [4]. Unilateral or bilateral cryptorchidism always accompanies testicular dysgenesis [5]. In both entities the Sertoli cells that are present in the male portion of the gonad or the whole gonad are immature, displaying round to ovoid nuclei and inconspicuous nucleoli [4]. Gonadoblastomas and more differentiated malignant germ cell tumors or Sertoli cell tumors develop in all cases of MGD, leading to the recommendation of bilateral gonadectomy of all Y-chromosome–containing material. In TH removal the opposite gonad to the assigned gender and a biopsy of the remaining gonadal tissue is sufficient for neoplasia [4]. These findings set the stage for a redundant theme in this chapter: that immature Sertoli cells may be a predisposing factor for testicular neoplasia. MIS continues to be secreted by the immature Sertoli cells; however, its low levels and persistence of Müllerian derivatives observed in these patients reflect either a low number of immature but functional Sertoli cells or, rarely, mutations in the IMS gene [6]. 2. Persistence of Müllerian Derivatives Syndrome in Males As described in Chapters 8 and 14 and reviewed elsewhere [7–15], MIS, also called anti-Müllerian hormone (AMH), is an important Sertoli cell product belonging to the transforming growth factor β (TGFβ) family and induced in the primordial Sertoli cells after commitment of the undifferentiated gonad to a testicular fate. Its main role during this period of mammalian development is complete regression of Müllerian duct derivatives such as the uterus and fallopian tubes. Lack of MIS activity in utero leads to persistence of Müllerian duct derivatives syndrome (PMDS) such as development of a uterus and fallopian tubes in a genetic male. A dog model for this syndrome has been described [16]. In this model bioactive MIS levels were detected in affected pups, leading the authors to speculate that the defect is caused by the inability of the MIS receptor to respond to the normal ligand. In humans, PMDS is a rare autosomal recessive disorder due to mutations in both alleles encoding MIS or in its type II receptor [17, 18]. Clinically, this syndrome is characterized by individuals with male phenotype with findings of a uterus and fallopian tubes that are not regressed and that are usually discovered at surgery [18]. Patients present with either bilateral cryptorchidism (rarely) or, more commonly, with unilateral descent of the testes in the scrotal sac, herniation of the ipsilateral uterus and fallopian tube, and contralateral testicular ectopia [18]. In this syndrome, the testes are not well anchored by the gubernaculum

FIGURE 21.1 Local regulation and partitioning of inhibin B secretion from the Sertoli cell. Production of

inhibin subunits by the Sertoli cell with possible production of βΒ subunit by spermatocytes. Stimulatory influence of FSH is shown, together with the modulatory influence of the germ cell component. The assembled, dimeric form of inhibin B is then secreted bidirectionally, into both seminiferous tubule and into the interstitium. Modulation of Sertoli and Leydig cell function by inhibins and activins is indicated. (Adapted from Anderson, R. A., and Sharpe, R. M. [2].)

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FIGURE 21.2

Changes in circulating inhibin B concentrations and FSH levels in blood from birth to adulthood. The neonatal peak in inhibin B is more prolonged than is that of FSH, and may reflect Sertoli cell proliferation. Following the quiescent period of childhood, puberty is associated with a progressive increase in inhibin B. Initially, this increase is associated positively with FSH levels but at a later (underlined) stage in puberty there is a switch to an inverse relationship between FSH and inhibin B that is characteristic of the adult human male. (Adapted from Anderson, R. A., and Sharpe, R. M. [2].)

and are more mobile and predisposed to testicular torsion. Abnormalities in spermatogenesis depend on the duration of cryptorchidism and the aplasia of the epididymis and the upper portion of the vas deferens, which adjoin the lateral uterine wall and mesosalpinx. Consequently, fertility is rare in these individuals [18]. In one study of 69 families with this syndrome, mutations in the MIS gene have been described in 45% of the affected patients. In 39% of affected subjects, individuals had normal MIS levels, and mutation in the MIS type II receptor was detected. The most common mutation is a 27-base-pair deletion in exon 10 of the receptor. In 16% of individuals, no specific mutation in either of these two genes could be detected, indicating a role for other genes important

FIGURE 21.3 Correlation between sperm count and inhibin levels (r = 0.39). (Adapted from Bohring, C., and Krause, W. [3].)

in the processing or signaling through MIS [18]. Wnt7a mutant mice also fail to have regression of the Müllerian ducts [19]. Furthermore, cooperation between Wnt signaling through β catenin and steroidogenic factor 1 (SF-1) are required for MIS type II receptor transcriptional regulation [20]. There are no described Wnt7a mutations in males with pseudohermaphroditism due to PMDS. Finally, PMDS has been associated with an increased incidence of testicular (predominantly seminomatous) cancers [7, 18, 21–23].

B. Testicular Dysgenesis Syndrome Several authors have hypothesized that environmental conditions [24, 25] may affect Sertoli and Leydig cells functions in fetal life and eventually lead to either (1) immediate manifestations such as incomplete or absent testicular descent (cryptorchidism) or to some of the above-described disorders of sexual differentiation or (2) delayed manifestations, including testicular cancer or spermatogenic abnormalities such as a low semen count. This group of interrelated conditions, hypothetically causally linked by a failure of the Sertoli cells to mature, has been termed testicular dysgenesis syndrome (TDS) [1, 26–28]. The full manifestations of this syndrome—hypospadias, cryptorchidism, testicular cancer, and oligo/azoospermia—are rarely seen in one individual, the most common presentation being altered spermatogenesis [28]. The impetus for the description of this syndrome as a separate clinical entity comes from epidemiological

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data linking hypospadias, cryptorchidism, and varying degrees of impaired spermatogenesis to testicular cancer [29–32]. Support for the authors’ contentions that a link between a Sertoli cell immaturity and testicular cancer exists includes the demonstration that biopsies of the contralateral (tumor-free) testes of individuals with testicular cancer show evidence of focal Sertoli cell only tubules, immature Sertoli cells, microliths, carcinoma in situ (CIS), and abnormal seminiferous tubule architecture [28]. Evidence suggests that in utero exposure to phthalate esters may lead to a syndrome similar to TDS [25], as reviewed in depth by Sharpe [24]. Recently, it has been shown that glial cell line–derived neurotrophic factor (GDNF) a Sertoli cell product that binds to the Ret proto-oncogene and in testes to its cognate receptors GDNF receptor α-1 and α-2 (GFR-α1 and α2) [33] regulates spermatogonial cell fate in the testes. GDNF levels are regulated by FSH [34]. Mice transgenic for GDNF initially accumulate undifferentiated spermatogonia, are infertile, and within a year frequently develop bilateral testicular tumors [35]. It would be tempting to speculate that there is a link between conditions leading to Sertoli cell immaturity and dysregulation of the GDNF/GFR-α1 signaling pathway at least in some cases of TDS, but no such clear linkage has been demonstrated.

C. Y-Chromosome Deletions Y-chromosome microdeletions in interval Yq11.23 have been associated with male infertility. These were attributed to the deletion of an azoospermia factor (AZF). In actuality, these deletions map to three different AZF loci termed: AZFa, b, and c (Fig. 21.4). AZFa deletions are almost always associated with Sertoli cell only syndrome (SCOS). AZFb deletions usually display a meiotic arrest, whereas deletions in the AZFc locus may display phenotypes varying from oligospermia to SCOS with azoospermia [36]. Sertoli cell function in affected individuals has only been partially studied. Staining for CK-18, a cytoskeletal marker that stains only prepubertal and, thus, immature Sertoli cells, was not found in subjects with AZF deletions indifferent of degree of impairment of

Pseudoautosomal region 1

spermatogenesis, but it was present in subjects with other causes of SCOS or mixed testicular atrophy [37, 38]. On the other hand, Foresta et al. [39] compared inhibin B and FSH levels among 27 subjects with AZF deletions and 75 subjects with severe testiculopathies that were either idiopathic or of known etiologies (cryptorchidism, testicular trauma, etc.). All subjects had either SCOS or severe oligospermia associated with severe hypospermatogenesis. Although FSH elevations and lower inhibin levels were present in both groups, they were significantly less pronounced in the group with AZF deletions [39]. These results are at odds with another study comparing inhibin B, FSH, and other hormonal and clinical variables between four groups: a subjects predominantly with AZFb and AZFc deletions, b subjects with idiopathic azoo/severe oligospermia, c subjects with Klinefelter’s syndrome (see Section VI.D), and d normal subjects [40]. The authors found that the elevations in FSH were indistinguishable between subjects with Y-chromosome deletions and subjects with idiopathic azoo/severe oligospermia compared to controls. In this study, substantial overlap existed in inhibin B levels between control and the Y-chromosome-deleted and idiopathic azoo/severe oligospermia subjects, making this comparison inconclusive. These findings cumulatively indicate that Sertoli cell function in subjects with Yq deletions is only partially and minimally affected and that there may be no difference between the impact of Y-chromosome deletions and other idiopathic causes of azoo/severe oligospermia on Sertoli cells. Of note is that higher inhibin B levels were observed in the previously mentioned Foresta study [39] in subjects with AZFc or AZFb+c deletions in comparison with the few (two subjects) with AZFa deletions. The authors speculated that AZFc or AZFb+c encodes for testes-specific genes, whereas AZFa encodes, in addition, for genes with ubiquitous expression in the somatic lineage, which may have a more direct role in Sertoli cell function and are, thus, are more likely to affect Sertoli cell function. This hypothesis is of interest but needs further corroboration with a higher representation of the more difficult to find subjects with AZFa deletions [39].

Heterochromatin Pseudoautosomal region 2

Centromere

AZFb

Yq

Yp AZFa

FIGURE 21.4

AZFc

Y-chromosome deletion map. (Adapted from Tomasi, P. A., et al. [40].)

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D. FSH Gene and Receptor Mutations FSH is a major regulator of Sertoli cell number and function (Chapters 10 and 11). The phenotype of mice lacking either the FSHβ subunit or the FSH G-protein coupled receptor and humans with various mutations has been extensively reviewed and is the subject of a vigorous debate on the difference in phenotypic expression between mice and man [41–52]. Male knockout mice for the FSHβ subunit have decreased FSH levels, normal T levels, a 50% reduction in testes size, and oligoasthenospermia, but they are fertile [53]. This phenotype is rescued by transgenesis of the human FSHβ gene on this background [54]. FSH receptor knockouts display a similar phenotype with differences from mice with FSHβ gene deletion, as described in Table 21.1. The most salient ones are elevated FSH levels and low T levels [55–57]. The elongated spermatids appear to have inadequate chromatin condensation [56]. The reduction in T levels is unexpected and has been attributed to loss of ligand-independent constitutive activity of the FSH receptor that is required for adequate proliferation of Leydig cells [58]. In the human, the clinical presentation of isolated FSH deficiency that is either (1) idiopathic (no detectable mutation in FSHβ) [59] (Fig. 21.5), (2) due to FSHβ [60–62] , or (3) due to FSHr inactivating mutations [48, 63] is intriguing in that there a high degree of variation in the clinical presentation with males being either azoospermic, as in FSH deficiency, or oligospermic, as in FSH receptor inactivating mutation (Table 21.2). In mice, an FSH deficiency or FSH receptor inactivation can lead to oligospermia, but fertility is maintained. This indicates possible species differences

TABLE 21.1 Characteristics of Homozygous FSHβ and FSHR Male Knockout Mice FSHβ Knockout

FSHR Knockout

Phenotype

Fertile (reduced?)

Fertile (reduced)

Testes size

Decreased 50%

Decreased 20–30%

Sperm concentration

Decreased 75%

Decreased 35%

Sperm motility

Decreased 40%

Decreased 25%

Normal sperm

Not provided

Decreased 30%

FSH

Low

3-fold increase

LH

Normal

Normal

Testosterone

Normal

Decreased by 67%

Heterozygote males are normal. (Adapted from Layman L.C. (2000). Mutations in the follicle-stimulating hormone beta (FSHβ) and FSH receptor genes in mice and humans. Semin. Reprod. Med., 18(1), 5–10.)

FIGURE 21.5 Results of repeated GnRH injections (100 μg every 2 hours) in a patient with selective FSH deficiency. (A), Variations of LH and FSH circulating levels. (B), Variations of inhibin B and activin A circulating levels. (Adapted from Mantovani, G., et al. [59].)

in how critical the FSH function is for the initiation and maintenance of spermatogenesis. Another phenotypic variation among the males within the various reported cases of FSH deficiency is that some have absent puberty and others do not. In addition, the T levels can either be normal or low. Only one activating mutation in the FSH receptor has been reported in humans [64]. The patient had a hypophysectomy for a pituitary tumor and subsequently no detectable gonadotropins and was treated with T alone; nevertheless he was able to sustain spermatogenesis and father three children.

IV. SERTOLI CELL ONLY SYNDROME SCOS was first described by del Castillo in 1947 and is a histological finding on testicular biopsies done for either obstructive or other causes of azoospermia. In one retrospective analysis of clinical features associated with bilateral SCOS, in 14% of cases, it was associated with severe oligospermia [65]. Its other clinical correlates are variable, ranging from normal (36%) to reduced testes size along with a similarly variable increased FSH (43% normal, 21% moderately raised,

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Chapter 21 Conditions Affecting Sertoli Cells TABLE 21.2 Summary of Published Human FSHβ and FSHR Gene Mutations in Males Males

Mutation

FSH

LH

Testosterone

Testes

Phenotype

FSH deficiency

None

Very low

Normal

Normal

Small

Infertility, azoospermia

FSHβ No. 1

Cys82Arg

Low

High

Normal

Small

Normal puberty, azoospermia

FSHβ No. 2

Val61X

Low

High

Low

Small

No puberty, azoospermia

FSHβ No. 3

Tyr76X

Unmeasurable

High

Normal

Small

Normal puberty, azoospermia

FSHR No. 1

Ala189Val (inactivating)

High

High

Normal

Small

Normal puberty, oligospermia, reduced fertility?

FSHR No. 2

Asp567Gly (activating)

Low

Low

On therapy

Normal

Normal sperm despite hypophysectomy

(Updated from Layman L. C. (2000). Mutations in the follicle-stimulating hormone beta (FSHβ) and FSH receptor genes in mice and humans. Semin. Reprod. Med., 18(1), 5–10.)

36% very high) [65]. It is most likely that this histological diagnosis is the end manifestation of several clinical entities. In this series, 18% of the cases were due to cryptorchidism and attributed to heat damage. Other entities leading to this histological phenotype are discussed in later sections. A nagging question is whether some forms of SCOS are due to alterations in the level of a critical Sertoli cell factor(s) crucial for support of germ cell proliferation or survival. We have recently shown that overexpression of human BCL-2 in Sertoli cells in mice under the control of the α-inhibin promoter leads to patchy and variable defects of spermatogenesis in seminiferous tubules [66]. Electron microscopy showed increased intercellular spaces and large vacuoles in Sertoli cells [66]. This form of patchy SCOS tubules has been associated with immaturity of Sertoli cell function as manifested by focal reappearance in the involved tubules of proteins that are usually downregulated in adult Sertoli cells such as MIS and CK-18 [14, 67–69] and loss of maturity markers such as the AR [70]. An argument is often made that absence of the germ cells from the seminiferous tubule will also lead to dedifferentiation of Sertoli cells into an immature functional state [1, 71–73]. This does not always appear to be the case since deletions in the Y-chromosome AZF intervals, although associated with SCOS, maturation arrest, or oligospermia, affect little if any of the maturity of Sertoli cells (Section III.C) [37–39, 67, 74]. We can conclude that many conditions, but not all, primarily leading to germ cell depletion and an SCOS phenotype may be associated with Sertoli cell immaturity. Much needs to be learned about the reverse scenario, whereby genetic or environmental conditions

first affect the Sertoli cell, then lead to this immaturity, causing subsequent germ cell loss and SCOS. It is not clear if the immature phenotype is an epiphenomenon or a necessary step in the pathogenesis of SCOS.

V. SERTOLI CELL NEOPLASIA In prepubertal boys, Sertoli cell tumors constitute 3% of all primary testicular tumors [75]. In the adult, estimates range from 0.4 to 1.5% of all testicular neoplasms [76]. They are histologically classified into three different categories: classic, large cell calcifying Sertoli cell tumors (LCCCST), and a sclerosing Sertoli cell tumor subtype (SSCT) [76]. In a recent compilation of all reported cases in the literature, the vast majority fall into the classic subtype followed by LCCCST and finally the much rarer SSCT, and different clinical prognostic characteristics are associated with each. Some of these tumors are associated with rare genetic diseases leading to multiple organ neoplasias. One such disease is Carney’s complex (CNC), an autosomal dominant multiple neoplasia syndrome, associated with spotty skin pigmentation, cardiac myxomas, and endocrine tumors. These include primary pigmented nodular adrenocortical hyperplasia leading to Cushing’s disease, growth hormone and prolactin secreting pituitary adenomas, and finally Sertoli cell tumors of the LCCCST subtype occurring in at least 33% and probably more of affected males [77]. Thyroid neoplasia and ovarian cysts have also been described [77]. Although this is a rare entity, the recent identification of the gene responsible for this experiment of

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nature may shed some light on one of the somatic “hits” or mutations required for the development of the spontaneously occurring nonfamilial variety of the LCCCST subtype of Sertoli cell tumors. Two groups have separately identified the mutation gene in kindreds afflicted with this disease [78–80]. This PRKAR1A gene encodes for the type 1α regulatory subunit of protein kinase A (PKA). PRKAR1A is then a tumor suppressor controlling PKA activity, which is known to be critical in mediating signaling in many endocrine tissues. As a matter of fact, there is evidence of a twofold increase in cAMP production in CNC tumors in comparison with non-CNC tumors [78]. Genetic heterogeneity is seen, however, because mutations in PRKAR1A can be found in all 40% of affected families [79]. Some clinical overlap is also seen between CNC and another hereditary multiple neoplasia syndrome, Peutz-Jeghers syndrome (PJS), which is characterized by hamartomatous polyposis of the gastrointestinal tract, mucocutaneous pigmentations [81], and association with sex cord tumors of the ovary and Sertoli cell neoplasms of the calcifying variety LCCCST [82–85] as in CNC. In some PJS kindreds, mutations have been found in another tumor suppressor gene, the serine threonine kinase STK11/LKB1 [81, 86–88]. Somatic mutations of these genes in nonfamilial forms of both diseases have been identified. In a subject with a Sertoli cell tumor, the clinician should look carefully for other manifestations of these diseases and, if identified, recommend genetic screening to the families of probands. Asymptomatic subjects with the mutation need intensified surveillance for other types of tumors associated with these diseases. Another gene found to be of importance in the development of Sertoli cell tumors is the α subunit of inhibin. Knockout mice for the α-inhibin gene invariably develop Sertoli cell tumors [89, 90]. FSH is necessary but not sufficient for development of these tumors [91]. Follistatin, estrogen receptors α and β, and several cell cycle regulators such as p27 Kip and cyclin D2 modify the Sertoli cell tumorigenicity of α-inhibin null mice [92–94]. For example, it has been suggested that increases in the cyclin D2/CDK4 activity necessary for the G1 to S phase transition and decreases in the tumor suppressor/CDK inhibitor p27 Kip levels are contributory to the tumorigenic process. Indeed, α-inhibin/ cyclin D2 double knockout mice live longer and have delayed development of Sertoli cell tumors [94], whereas α-inhibin/p27 Kip double knockouts develop early tumors and die sooner when compared to α-inhibin knockout mice [92]. Inhibin-α/estrogen receptors α and β triple knockouts are less prone to early Sertoli cell tumor formation, indicating the need for redundant signaling through both estrogen

receptors for a more aggressive Sertoli cell tumor phenotype to occur [95].

VI. GENETIC SYNDROMES WITH A SERTOLI CELL PHENOTYPE A. Fragile X Syndrome Fragile X syndrome presents with a cluster of symptoms including hyperactivity, cognitive impairment, autistic features, and—most interestingly from our perspective—macroorchidism or ovarian failure [96–98]. It is the most common form of inherited mental retardation in humans. Expansion of CGC repeats in the 5’ untranslated region of the FMR gene leads to methylation of its promoter and lack of its expression in several target tissues, leading to the observed phenotype [99]. Although larger testes volumes can be observed as early as between the ages of 2 and 7 in Fragile X prepubertal boys, frank macroorchidism is the rule after the age of 8 [100]. The macroorchidism was first studied histologically in two men with the syndrome by testicular biopsies [101]. These showed interstitial edema, increased lysosomal inclusions in Sertoli cells, and impaired spermiogenetic differentiation [101]. These changes were attributed to increased pressure from the interstitial edema. Autopsy findings in two affected individuals who died with sudden death revealed “interstitial cell hyperplasia” in the macroorchid testes [102]. Unilateral macroorchidism has also been reported [103]. Not all cases of mental retardation and macroorchidism have been associated with the CGC expansion of the FMR gene [104], indicating some genetic heterogeneity. The most well-characterized testicular phenotype comes from the work of the Themmen group, which performed a detailed characterization of the histological phenotype of testes of FMR1 knockout mice. Their findings in FMR–/– mice point toward a significant increase in Sertoli cell proliferation (by BRDU labeling) from day 12 postcoitus to day 15 postnatally and an increase in testes weight that becomes significant on day 15 postnatally exceeding wild type by 30% by 6 months [98, 99, 105]. The distribution of FMRP in mice at days 3 to 7 postnatally is predominantly in the cytoplasm of spermatogonia and to a lesser extent in Sertoli cells. Subsequently, Sertoli cell staining disappears and is limited to spermatogonia and, in addition, in adults to midpachytene spermatocytes [106]. Unfortunately, this study did not determine FMRP expression in the critical period when the first Sertoli cell proliferation wave occurs.

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FIGURE 21.6 Alignment of FMRP and related proteins: FXR1P and FXR2P. (Adapted from Hoogeveen, A. T., et al. [99].)

The increase in Sertoli cell proliferation in utero was not attributable to changes in FSH levels by RIA and FSH receptor mRNA by RNA protection assay as determined in both the wild-type and FMR knockout mice [104]. The FSH receptor mRNA was equally downregulated by FSH injection in both types of mice [104], leading the authors to conclude that FSH signal transduction is unchanged in the knockouts mice and is not the cause of the increase in Sertoli cell proliferation in these mutant mice. FMR1 belongs to a family of RNA binding proteins FXR1 and FXR2 that share significant homology and conserved domains (Fig. 21.6). Knockouts for FXR1 die shortly after birth, whereas FXR2 knockouts display a somewhat similar phenotype as FMR1 but do not seem to have the macroorchidism [98, 107]. It is thought that all three proteins play a role in the transport of specific proteins from the nucleolus/ nucleus into the ribosome [99, 108]. FMRP associates with the translating polyribosomes in an RNAdependent manner via messenger ribonucleoproteins (mRNP) [109, 110]. A mutation in FMRP preventing this polyribosome association leads to full manifestation of the disease [111]. It is likely then that FMRP plays the role of a transcriptional regulator and in the brain plays a role in modulating neuronal plasticity [112–114]. A number of mRNAs that associate with the FMRP–mRNP complex and that play a role in these processes have been isolated (Table 21.3). A similar approach can be utilized to identify mRNAs of testicular origin that play a role in the FSH-independent phase of control of Sertoli cell proliferation in utero. In summary, in spite of an enlargement of testes in both species, the description of the histology of macroorchidism in human Fragile X–affected individuals does not parallel what has been described in mice. In the mouse, the phenotype is attributed to an increase in Sertoli cell proliferation in utero whose mechanism is still poorly understood.

B. Adrenoleukodystrophy Adrenoleukodystrophy (ALD) is an X-linked disorder characterized by neuron demyelination, adrenal insufficiency, and testicular dysfunction. The disorder is attributed to accumulation of very long chain fatty acids (VLCFAs) in various tissues due to inactivating mutation of ALDP. This gene is similar to the ATP-binding-cassette (ABC) transporter proteins and is responsible for the microsomal β-peroxidation of saturated VLCFA. ALD is a major cause of adrenal insufficiency in the Western world. According to one report [115], it accounts for 35% of idiopathic Addison’s disease. The testicular phenotype of subjects with ALD is less pronounced and consists of trilamellar inclusions in Leydig cells on electron microscopy associated with VLCFA accumulation. Less frequently there is a decrease in the number and vacuolization of Sertoli cells [116–121]. These changes are occasionally progressive with germ cell depletion varying from abnormal

TABLE 21.3 Synaptic plasticity

Axon guidance

Fragile X mental retardation protein 1

MAP1B

Munc13

NAP-22

NAP-22

Semaphorin 3F

Rab6-interacting protein 1

Inhibitor protein ID3

SAPAP4 Sec7-related guanine-nucleotide exchange factor Fragile X mental retardation protein (FMRP) ligands related to synaptic plasticity and axon guidance. It is not clear if some of these ligands are relevant to FMRP action in the testes. (Jin, P., and Warren S.T. (2003). New insights into fragile X syndrome: from molecules to neurobehaviors. Trends Biochem. Sci. 28, 152–158.)

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FIGURE 21.7

LH (a), FSH (b), testosterone T (c), and free testosterone (d) concentrations in adrenoleukodystrophy (ALD) patients (also known as adrenolmyeloneuropathy [AMN]) with (X) and without (W) adrenocortical insufficiency in comparison with healthy controls (O). The continuous lines represent the mean values and the dashed lines the upper and lower normal range. (Brennemann, W., et al. [123].)

semen characteristics to maturation arrest and azoospermia [120, 122]. In 49 patients with ALD, endocrine investigations revealed that 31 subjects had elevated luteinizing hormone (LH), whereas 28 patients had an increase in FSH levels (Fig. 21.7) without any significant change in inhibin B levels. In spite of the LH elevation, T levels were maintained, probably at the expense of the high LH, which indicates a state of compensated hypogonadism [123]. In spite of the vacuolization of some of the Sertoli cells and the elevated FSH, the normal inhibin B indicates a less severe Sertoli cell dysfunction. A mouse deletion mutant of ALD shows some of the morphological changes observed in human testes (Fig. 21.8). The mice have delayed infertility, and testicular atrophy was seen in at least one animal. Unfortunately, the testicular and reproductive hormone profiles of these animals were not studied in detail.

C. Congenital Adrenal Hyperplasia The major manifestations of the nonclassical virilizing form of congenital adrenal hyperplasia (CAH) are due to diminished activity of the enzyme 21 hydroxylase in the adrenal gland. This leads in this mild form to enough corticosteroids being produced but at the expense of a high ACTH and shunting of the 21-hydroxylase precursors to the androgen synthesis pathway. In the female, the phenotype is straightforward, presenting with virilization and polycystic ovary syndrome (PCOS). However, various degrees of male infertility do occur in some of the afflicted males and some present with azoospermia [125–129]. It seems that the etiology of the infertility in these males is heterogenous. Theoretically, the increase in androgen production from the adrenals could block the gonadotropins and lead to a decrease in intratesticular T and impaired spermatogenesis.

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product of these tumors suppressing inhibin B secretion, which would explain the loss of the negative feedback loop on FSH. One publication [125] actually measured inhibin B levels in pubertal boys with nonclassic CAH and some with adrenal rest tumors. These levels were normal, however, because all of the subjects in this study had normal FSH.

D. Klinefelter’s Syndrome

FIGURE 21.8

Testis of a 6-month-old mutant (ALD -/Y) mouse. Note the atrophy of seminiferous tubules (pale), an increased number of interstitial cells (darker staining), and the lack of sperm cells. (Adapted from Forss-Petter, S., et al. [124].)

Some of the subjects develop adrenal rest tumors, which are derived from ACTH-responsive cells that have erratically migrated to the testes. This type of infertility has been blamed either on the tumor obstructing the hilus of the testes leading to azoospermia [127] or to tumor expansion and destruction of testicular parenchyma [127, 129]. In two separate series, subjects with the adrenal rest tumors had high FSH levels, suggesting either direct obstructive damage or impaired Sertoli cell function due to a paracrine

This syndrome, characterized by at least one extra X chromosome (XXY), is best known for its association with infertility, small testes, and late-onset hypogonadism. However, the clinical presentation is more elaborate with multiple organs and systems affected [130]. The number of extragonadal abnormalities correlates with the number of supernumerary X chromosomes [131, 132]. XXY patients usually present with a learning disability and cognitive executive and neurobehavioral and psychological dysfunction all recently described by our group [130, 132]. This makes it conceivable that at least some of the manifestations of this syndrome are due to overdosage of genes that escape methylation on the X chromosome [133]. Abnormalities in skeletal features, dentition, and other features have also been described [130, 134–136]. Because the germ cell compartment accounts for 85% of the volume of the testis, it is no surprise that the loss of germ cells and hyalinization of the seminiferous tubules leads by puberty to small firm testes and azoospermia [137]. In Klinefelter’s syndrome, spermatogenesis may be occasionally observed in isolated tubules [138, 139]. It is unclear if this prepubertal depletion of germ cells is due to dosage excess of a gene(s) on the X chromosome expressed in the germ line or in the somatic lineage. Several lines of evidence support a role for the Sertoli cell in the pathogenesis of the testicular phenotype. In a review of 36 biopsies in XXY subjects aged 15 to 66, Skakkebaek described, in addition to the hyalinized seminiferous tubules, two types of nonhyalinized tubules [140]: type A, with classical Sertoli cells, and type B, with cells with ovoid nuclei probably representing atypical Sertoli cells (Fig. 21.9). Other authors have found abnormalities of Sertoli cell morphology and ultrastructure such as an altered chromatin pattern, absence of annulate lamella, lipid droplets, and glycogen-filled vacuoles in the cytoplasm [137, 141–143] . These findings were not universal among all studies. A more significant indication of late Sertoli cell dysfunction in this syndrome comes from a recent report characterizing longitudinally the changes in inhibin B levels in affected XXY boys [144].

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FIGURE 21.9

Types of Sertoli cells in patients with Klinefelter’s syndrome: Left panel: Type A tubules with classical Sertoli cells. Right panel: Type B tubules with cells with ovoid nuclei probably representing atypical Sertoli cells. (Adapted from Skakkebaek, N. E. [140].)

The levels were normal before puberty but then started rising with early puberty and then fell to the low/undetectable levels in XXY adults (Fig. 21.10) corresponding to the high FSH levels found in this agegroup. The authors concluded that Sertoli cell function based on inhibin B levels is normal in prepubertal boys, but starts declining during adulthood, when its presence is significantly reduced due to lack of positive paracrine influences from the dying germ cells. In XXY mice, Hunt et al. [145] reported early germ cell loss in utero but interestingly found that the XXY fetal germ cell proliferative potential was unaffected in vitro when these cells were cultured on euploid somatic feeder layer cells. This suggests a perturbation in the

XXY Sertoli cell function as the potential etiology for the cessation in proliferation of fetal germ cells in utero in this model. Our group has recently characterized the progressive germ loss in a similar XXY mouse model and found that the germ cell loss is complete by day 10 postnatally [146], paralleling the time course of germ cell loss observed in humans. Interestingly, the ultrastructure of Sertoli cells in this model exhibited a scanty amount of cytoplasm and complete absence of lateral process characteristic of active Sertoli cells. In addition, nests of apparently degenerating Sertoli cells were noted in some tubules. These changes are indications of the cellular inactivity of the Sertoli cells [146] (Fig. 21.11).

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FIGURE 21.10 Left, Longitudinal changes in inhibin B levels in 11 control boys and 7 boys with Klinefelter’s syndrome undergoing serial blood sampling as they went through puberty. Samples were classified into time periods before and after the onset of puberty, defined as a testicular volume larger than 3 ml (prepuberty, 12–24, 6–12, and 0–6 months; and postpuberty, 0–6, 6–12, 12–24, and 24–36 months postpuberty). Right, Longitudinal inhibin B levels in 15 young adults with Klinefelter’s syndrome are shown in relation to a 95% prediction interval of healthy normal men. Data are expressed as medians, with error bars representing the range. (Adapted from Christiansen, P., et al. [144].)

A comparison of inhibin B levels and FSH levels in a study described earlier found that FSH elevations and inhibin B suppression were most pronounced in subjects with Klinefelter’s syndrome when compared with subjects with idiopathic or Y-chromosome deletions related to azoo/severe oligospermia. The authors speculate that there is a lower Sertoli cell mass in XXY males [40]. In a recent attempt to test the hypothesis that transplanted normal bull germ cells may partially rescue the azoospermia of a nonmosaic XXY bull, the donor germ cells were rejected and could not incorporate in the testes. This germ cell rejection invalidated the experiment and left the contribution of XXY Sertoli cells to the germ cell depletion in Klinefelter’s as an important question that our group along with other collaborators is trying to solve.

E. McCune Albright Syndrome Macroorchidism has been reported in a prepubertal boy with McCune Albright syndrome and a Gsα gene activating mutation in Sertoli cells, leading to Sertoli cell autonomous hyperfunction and hyperplasia, increased inhibin and MIS production, and blunting of FSH response to GnRH [147].

VII. NEONATAL, PUBERTAL, OR ACQUIRED ADULT CONDITIONS AFFECTING SERTOLI CELL FUNCTION A. Hypothyroidism Thyroid hormone (TH) deficiencies or excesses have varying effects on the male gonad, depending on

the timing of the hormonal perturbation during development [148]. It appears that hypothyroidism does not significantly affect the hypothalamic pituitary testicular access during adulthood. Goitrogen-induced neonatal hypothyroidism extending to puberty appears to retard Sertoli cell proliferation and final testes size. On the other hand, a transient state of hypothyroidism occurring neonatally but with recovery prior to puberty leads to a net increase in the final Sertoli cell number with an associated increase in testes size and sperm number. In this section, we focus on the effects of transient and permanent hypothyroidism on Sertoli cell number and function. Most boys with prepubertal onset hypothyroidism present with macroorchidism and an elevated FSH. After treatment, FSH elevation and the macroorchidism may or may not normalize [148, 149]. The only histological information available is through the rare biopsy or testicular specimen available from affected individuals. Prepubertal testes show an increase in the seminiferous tubule compartment, early onset of spermatogenesis, and no change in the number of Leydig cells. If left untreated, juvenile hypothyroidism leads by puberty to hyalinization of the seminiferous tubules and peritubular and interstitial fibrosis with a decreased number of Leydig cells [148, 149]. An ontogeny of the expression of thyroid hormone receptors (TRs) from fetal to adult human testes shows that TRβ was not expressed in any of the tissues. However, TRα2 and TRα1 were both coexpressed exclusively in immature Sertoli cells predominantly during the fetal and prepubertal periods. The TRα1 expression waned to barely detectable levels by adulthood [150]. Similar data show that both TRα1 and TRα2 are also expressed in the same narrow developmental window in the rat

FIGURE 21.11

Ultrastructural characteristics of testes from XXY mice compared to wild type controls: A, XY male mouse showing normal ultrastructure of Sertoli and germ cells. B, Sertoli cell–only tubule from an XXY male; Sertoli cells have spheroidal nuclei (N) and a scanty cytoplasm (asterisk) with absence of lateral processes characteristic of active Sertoli cell. C, XXY tubule showing formation of a nest of degenerating Sertoli cells. D, XXY interstitial tissue with hypertrophied Leydig cell (L) and a macrophage (M). Note the abundant endoplasmic reticulum the large systems of concentric cisternae (asterisk). (Adapted from Lue, Y., et al. [146].)

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function of the cyclin inhibitor p27Kip1 in the suppression of the Sertoli cell cycle.

B. Cryptorchidism

FIGURE 21.12

Comparison of the levels of THRalpha1 mRNA and TH nuclear binding during the rat testis development. The nuclear TH binding capacity in whole testis during fetal and postnatal development was also plotted (broken line). The correlation between THRalpha1 mRNA levels and TH binding in testis of different ages is shown in the inset. (Adapted from Jannini, E. A. et al. [148].)

testes in the nuclei of both Sertoli cells and spermatogonia [148, 151–154] (Fig. 21.12). Although most actions of TH are generally mediated through TRβ, it appears that the actions of TH in the testes are exerted through TRα1, predominantly through action on the immature Sertoli cell. A large body of literature [155–165] implicates thyroid hormone in the maturation of Sertoli cells. Transient treatment with goitrogens of neonatal rats leads to a delay in the maturational process and continued proliferation of the Sertoli cells. Neonatal hypothyroidism leads to retention of expression of markers of Sertoli cell immaturity such as MIS and suppression of Sertoli cell lactate production and ABP expression [164] as well as delayed expression of inhibin and clusterin [155, 158, 159]. This is accompanied by an increase in the Sertoli cell mitotic index [166, 167]. It, thus, seems likely that thyroid hormone enhances Sertoli cell maturity at the expense of a suppression of proliferation. Neonatal and prepubertal transient hypothyroidism delays the terminal differentiation of Sertoli cells and allows for an increase in the length of period in which Sertoli cells proliferate and consequently for an increase in germ cell output leading to the observed macroorchidism. If this process extends beyond puberty it becomes deleterious and lead to fibrosis and hyalinization of the seminiferous tubules. In other tissues, TH inhibits the cell cycle by upregulating the cyclin kinase inhibitor p27Kip1. Mice lacking p27Kip1 have large testes. It appears that TH exerts the same regulatory effects on the Sertoli cell cycle, providing a rationale for the increased proliferation of Sertoli cells observed during prepubertal hypothyroidism [168]. So far we have described at least two hormones (TH and inhibin ) that regulate the

Cryptorchidism, defined as failure of the testes to reach the scrotal sac, is an extremely common entity with many potential clinical ramifications in adulthood such as impacts on fertility and incidences of testicular cancer. The details of the journey of the testicle from its original abdominal location to the scrotum reviewed by Ivell and Hartung [169] are shown in Figure 21.13. Ongoing debate surrounds data that suggest an increase in the incidence of cryptorchidism in the United States and Western Europe and the linkage of the surge to environmental endocrine disruptors [24–28, 32, 170–172]. Central to this hypothesis, as discussed in Section III.B, is that prenatal exposure to estrogens (DES), xenoestrogens (organochlorine pesticides), and antiandrogens (flutamide, phthalates) leads to alterations in Sertoli cell function that predispose perinatally to cryptorchidism and in adult life to TDS with a high propensity for testicular cancer and lower semen counts. The germ cell death in testes subjected to longstanding cryptorchidism is related to the relatively higher (2–8°C) temperatures encountered in the abdomen or the inguinal canal as opposed to the 32 degrees prevalent in the scrotum [173, 174]. The cells most sensitive to induction of apoptosis by the heat effect are the meiotic lineage and round early spermatids in boys. However, there is also evidence for a reduction in number of spermatogonia and a failure of morphological maturation of at least some of the Sertoli cells starting at puberty [175, 176]. In cases of unilateral cryptorchidism, abnormalities in spermatogenesis in multiple species are encountered in the noncryptorchid testes [177–179]. These changes, ranging from a reduction in spermatogenesis up to SCOS in the contralateral testes, argue for either an independent primary defect that affects both spermatogenesis and testicular descent independent from heat, or a secondary effect induced on the contralateral testes by the degenerating cryptorchid testis. So for the purposes of our discussion, one can frame the question as to whether primary genetic or environmentally induced Sertoli cell changes are responsible for (1) failure of descent and spermatogenesis descent in the affected testes, (2) concomitant spermatogenetic and Sertoli cell defects in the opposite descended testis, or (3) if all manifestations are a direct consequence of heat exposure on one or both testis. In favor of the first scenario, we review data implicating either genetic or environmental factors that are the primary trigger

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FIGURE 21.13 Schematic illustration of the events involved in the descent of the differentiating testis from a perirenal to an inguinal position. (A) The undetermined gonad, at a time equivalent to mouse embryonic day 12, with Müllerian (pink) and Wolffian (blue) derivatives both present, and the undifferentiated gonad in a perirenal position. (B) Shortly before birth androgens cause an involution of the cranial suspensory ligament (CSL), allowing the thickening gubernaculum, under the influence of insulin-like factor-3/ relaxin-like factor (INSL3/RLF), to retain the testis in the inguinal region while the embryo grows, effectively moving the kidney to a relatively more dorsal location. Müllerian inhibiting substance (MIS) from developing Sertoli cells causes the involution of the Müllerian duct, while androgens continue to stimulate the development of Wolffian derivatives and secondary sexual characteristics. Finally, in (C), the gubernaculum facilitates the passage of the testis into the scrotum with the development of an inguinal canal. (D) In the female, the CSL is retained holding the ovary close to the kidney, as the remaining abdomen grows ventrally. The Müllerian duct is retained later to form the oviduct and uterus, while the Wolffian derivatives and gubernaculum involute. (Adapted from Ivell, R., and Hartung, S. [169].)

leading to both abnormalities. Foresta et al. (Table 21.4) reported that 27.5% of males with severe impairment of spermatogenesis and bilateral cryptorchidism and 25% of subjects with idiopathic severe male factor infertility were affected by Y-chromosome deletions [180]. Cryptorchidism due to etiologies leading to various forms of hypothalamic hypogonadism or incomplete androgen insensitivity have also been reported [181–191]; however, in one study, no relationship was found between AR mutations and cryptorchidism [192], and not all subjects with cryptorchidism respond to hCG or GnRH treatment [193–198]. Thus, androgen insufficiency alone is insufficient to explain all cases of cryptorchidism. As previously discussed, some mutations in the MIS gene may well present with cryptorchidism [17, 199, 200]. Recently, a gene expressed in the Leydig cell that encodes insulin-like factor 3 (INSL3, also called relaxin-like factor) was recently characterized. Mice lacking this gene present with a surgically correctable form of cryptorchidism as their only phenotype [169, 201]. No human mutations in either INSL3 or its receptor have been found so far in cryptorchidism in men [169]. Mice lacking Hoxa-10 or mice with severed

genitofemoral nerves and, hence, a decrease in its neurotransmitter CGRP (calcitonin related peptide) also have cryptorchidism [202–205]. Based on the multiple but rare genetic entities may manifest as cryptorchidism and may be useful tools in deciphering the mechanism of testicular descent. It is unlikely, however, that these entities account for a high percentage of the rather frequent incidence of cryptorchidism. Much work remains to be done to fully understand how environmental factors affect testicular descent. However, a considerable body of literature supports the notion that environmental endocrine disruptors in the form of exoestrogens or antiandrogens lead to Sertoli cell immaturity or functional dedifferentiation of the Sertoli cells (as evidenced by a panoply of Sertoli cell maturity or immaturity markers discussed in Section II) and play a role in the vast majority of cases with acquired cryptorchidism and its sequelae such as TDS [1, 25, 28, 70, 171, 206–220].

C. Varicocele Several mechanisms have been implicated in the pathophysiology of varicocele-induced negative

399

Chapter 21 Conditions Affecting Sertoli Cells TABLE 21.4 Clinical Characteristics of Patients with Severe Bilateral Testiculopathies, with and without Y-Chromosome Deletions, Compared with Patients with Testiculopathies of Known Etiology or with Less Severe Testiculpathy or Fertile Controls Patients

n

Seminal pattern (cells x 106 mL)

11

0.7 ± 0.6

29

28

Testicular volume (mL)

FSH (IU/L)

LH (IU/L)

T (nmol/L)

9.5 ±3.0a

18.2 ±5.5a

3.4 ±1.2

14.5±4.2

0.9 ±0.9

10.1 ±3.7a

17.4 ±6.7a

3.1 ±0.9

15.6±3.9

0.8 ± 0.6

10.3±2.1a

17.1 ±5.6a

3.1 ±1.2

15.8±4.3

82

1.0 ± 1.3

9.8 ±2.3a

16.3 ±8.2a

2.9 ±1.0

14.9±3.6

20

14.7 ±2.9

UT:9.7 ±2.8a

3.6 ±1.5

3.3 ±1.7

15.4±4.2

Unilateral cryptorchidism with severe bilateral testiculopathy (Group 1) With deletion Without deletion Idiopathic bilateral severe testiculopathy (Group 3) With deletion Without deletion Unilateral cryptorchidism with

DT:15.8 ±4.3

normal descended testis (Group 2) Idiopathic bilateral moderate

20

13.8±3.3

14.1 ±3.6

3.7 ±1.9

3.7 ±1.5

16.2±3.8

50

1.1±0.8

10.2±3.1

15.8 ±4.8a

3.8 ±1.3

15.7±3.2

100

53.6±37.1

15.6 ±3.4

3.2 ±1.2

3.6 ±0.8

16.1±3.1

testiculopathy (Group 4) Severe testiculopathy of known etiology (Group 5) Fertile controls

UT, undescended testis; DT, descended testis. 0.05 vs. controls. Foresta, C., Moro, E., Garolla, A., Onisto, M., and Ferlin, A. (1999). Y chromosome microdeletions in cryptorchidism and idiopathic infertility. J. Clin. Endocrinol. Metab. 84, 3660–3665. aP<

change in semen parameters. These include an increase in testicular temperature and the possible tissue hypoxia and oxidative damage associated with venostasis. The impact of varicocele on Sertoli cell function has been studied both in animal models and in humans before and after varicocelectomy using markers of Sertoli cell function. Surgically induced varicocele by partial ligation of the left renal vein in rat leads to a significant decrease in ABP and transferrin expression in the testis by immunohistochemistry [221]. In men with varicocele, apical Sertoli cell cytoplasmic vacuolization or truncation is occasionally found on testicular biopsy [222, 223]. Granular transformation of Sertoli cells with accumulation of secondary lysosomes and scant organelles has also been found [224]; however, this finding is nonspecific and can be found in biopsies of testes affected by other etiologies. Significantly, changes in Sertoli cells functions have also been found and were frequently corrected by surgery. Many of these trials were not appropriately controlled. Most compared Sertoli cell markers to either a population of fertile men and/or to subjects with varicocele before and after varicocelectomy. Nevertheless, Kosar et al. found that semen transferrin levels in humans were significantly reduced in subjects with

varicocele and had only a minor nonsignificant improvement in semen transferrin levels after surgery [225]. A similar significant decrease in both transferrin and ABP immunoreactivity has been found in Sertoli cells after surgically induced varicocele in rodents. Another observational study found a significant increase in mean inhibin B levels along with an improvement in semen parameters after varicocele surgery [226] (Fig. 21.14). No other hormonal parameters changed. A universal improvement in inhibin B levels after surgery is not observed in all patients [227]. Similarly the decrease in inhibin levels in varicocele patients is not consistently observed in varicocele patients [228–232]. From these data, it is unclear if the changes, occasionally observed in varicocele patients in the morphology or function of Sertoli cells, represent a significant step in the sequence of pathophysiological events leading to impairment in semen parameters or just a nonspecific marker of alteration in Sertoli cell health.

D. Orchitis One frequent reason for orchitis in adolescent boys is caused by Paramyxoviruses or mumps, which predominantly involves the parotid glands but may also

FIGURE 21.14

Individual levels of inhibin B, FSH, free androgen index, sperm count and sperm motility before and after varicocelectomy. Dots above and below the line y = x represent patients with an increase and decrease in the variable after surgery, respectively. (Adapted from Pierik, F. H., et al. [226].)

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involve the testes. Consequences are testicular atrophy and potential sterility [233, 234]. It is observed to occur alone or in a few case reports in association with idiopathic thrombocytopenic purpura (ITP) in adults vaccinated with mumps vaccine [233, 235]. One speculative mechanism as to how Sertoli cell function might be affected in this context is that signal transducer and activation of transcription (STAT1 and 3), usually produced by Sertoli cells [236–238], is directed for proteosome degradation by the virus, depriving the cell from transcription factors necessary for the immediate early gene response and for induction of several interferons’ production, which is necessary for mounting an antiviral response [239–243]. Evidence for involvement of the Sertoli cell is present in at least one model of autoimmune orchitis. In this model, autoimmune orchitis was produced through immunization of rats with testes homogenates and adjuvant, and it led to the appearance of mononuclear cellular infiltrates and germ cell sloughing. Associated with these changes was a significant decrease in inhibin B levels, which was observed in the experimental group as opposed to controls, correlating with a threefold increase in FSH levels [244]. The Sertoli cell contributes, through its junctional complexes, providing an immune privileged environment to germ cells. Violation of that environment through trauma, surgery (inguinal hernia repair, vasectomy), or other types of infection, may lead to exposure of germ cell antigens to the host immune system and generation of autoimmune orchitis [234, 245–259].

E. Testicular Torsion There is substantial evidence that nitric oxide (NO) plays a role in the induction of apoptosis in the ischemia/reperfusion injury induced by testicular torsion/detorsion [260–267]. It is unclear whether the associated increased apoptotic changes, which include demise of some Sertoli cells and an increase in the phagocytic activity of others [268], all observed in the contralateral testes of these subjects [269, 270], are due to (1) disruption of the testicular blood–testes barrier, (2) increased production of circulating cytokines from the contralateral affected testes [270, 271], (3) or a manifestation of a primary genetic defect that leads to both the testicular torsion and the apoptotic changes observed in the contralateral testes. This last hypothesis seems unlikely. One publication claims that the release of NO from the ischemic testes may have an antiapoptotic effect in the contralateral testes [272]. In delayed repair of testicular torsion or in experimental models of unilateral testicular torsion, inhibin B levels decrease, indicating damage to the Sertoli cell in the “unaffected” testes [273, 274].

F. Vasectomy Vasectomy is one of the two most widely used methods of contraception in men [275]. Although fertility can be achieved after vasectomy reversal, there is evidence that a large number of men develop antisperm antibodies, limiting the success of fertility reversal after this procedure [255, 275]. There is conflicting evidence on the effect of vasectomy on Sertoli cell function. Measurement of some markers of Sertoli cell function (inhibin B, transferrin, LDH-X prostaglandin D synthase) in seminal plasma may in some cases be reduced [276–279], but these are not useful in predicting the presence or reversibility of Sertoli cell damage. Measurement of serum inhibin B levels in subjects with all causes of obstructive azoospermia showed no difference with normal controls [280]. Histological examinations of testicular biopsies in men postvasectomy show an increase in the thickness of the seminiferous tubules, interstitial fibrosis, sperm granulomas, and a reduction in Sertoli cell number [281, 282]. Others have reported only a minor insult to spermatogenesis after this procedure [283]. In Langur monkeys, 540 days after vas occlusion, there was vacuolization of Sertoli cells and other focal degenerative changes in germ cells. This is in contrast with an earlier study, which failed to show any alteration in spermatogenesis at 18 months after vasectomy of Macaca fascicularis monkeys [284]. In rats, 3 and 8 wk after vasectomy, there was no difference in ABP accumulation after ligation of the efferent duct in either vasectomized or sham operated animals, leading the authors to conclude that in rats, this procedure had no impact on Sertoli cell function [285]. Studies in mice are not in agreement, with some showing a modest but statistically significant increase in Sertoli cell nuclei per tubule [286]. Dramatic changes in all testicular cells were recently observed in a different strain of mice [287]. In addition, in guinea pig and in rabbit, evidence for an autoimmune orchitis with various degrees of Sertoli cell damage was also found [288, 289]. Species and strain differences, length of duration of ligation of the vas, and technique may account for the variable effects on Sertoli cell function. The results in men support the notion that long-standing obstruction after vasectomy may have a subtle negative impact on Sertoli cell count and function.

G. Diabetes Most of the discussion surrounding the impact of diabetes mellitus (DM) on male reproduction focuses on diabetes’ contributions to male sexual dysfunction. There is, however, some data to suggest, both in animal models and men, that in a few instances the same

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genetic or environmental defects affecting Sertoli cell function are linked to the development of DM or abnormalities in the availability or storage of nutrients. For example, hemochromatosis is a cause of diabetes due to both infiltration and toxic effects on β and liver cells and it leads to insulinopenia and insulin resistance [290]. However, hemochromatosis is also a cause of pituitary failure and of testicular dysfunction due to deposits of hemosiderin and lipofuscin in the pituitary and in the testes including in Leydig and Sertoli cells [290, 291]. Prader-Willi syndrome, which is attributed to loss of paternal allelic expression of several genes on chromosome 15q11-q13, is associated with hypothalamic hypogonadism, central obesity, insulin resistance, and mental retardation, but may also lead to cryptorchidism and primary testicular failure and SCOS [292, 293]. It is also clear that some alterations in the reproductive axis are directly induced by diabetes, but it is unclear as to what their contribution is to idiopathic male factor infertility or if they worsen the phenotype of other causes of Sertoli cell dysfunction. Multiple clinical studies reviewed elsewhere [294] show mild to moderate changes in semen parameters. The severity of these changes is dependent on duration of disease, level of diabetic control, and concomitant presence of neuropathy. This review also points out that in welldesigned studies, and if patients are metabolically well controlled, there is little significant change in pituitary gonadotroph secretion. The Sertoli cell ultrastructural phenotype of testicular biopsies obtained from men with diabetes is not consistent. In one study, testicular biopsies from men with either oligospermia or impotence showed vacuolated Sertoli cells and a high degree of apical cell membrane degeneration and redundancy along with altered Sertoli–to-spermatid junctional specializations [295]. However, an earlier study by Faerman et al. reports hypospermatogenesis, but few changes in Sertoli cell morphology [296]. Measurement of inhibin levels in 33 adolescent males and young adult patients affected by Type 1 diabetes, with a mean disease duration of 12.7 ± 5.8 yr and suboptimal, but not severe, lack of metabolic control, and in an age-matched group of 36 healthy control subjects showed comparable levels of FSH and inhibin B levels [297]. These studies in men contrast with studies in animals or in vitro models where an association between diabetes and infertility- or hyperglycemia-induced alteration in Sertoli cell function is more readily demonstrable. For example, in two models of diabetes—streptozotocin-induced diabetes in male Sprague-Dawley or Wistar rats and diabetes naturally occurring in BB rats—a marked suppression of LH and

steroidogenesis [298–300] is seen. In one of these studies, FSH levels were also suppressed [299], and epididymal ABP levels were elevated. In the BB rat, there is some ultrastructural evidence of disruption of Sertoli–Sertoli junctional complexes [301], and more severe alterations in Sertoli cell morphology is apparent after 250 days of age in these rats [300]. Incubation of Sertoli cells with a cocktail of substances simulating diabetic conditions, including an increase in glucose concentrations, leads to increased lactate production and suppression of FSH-stimulated ABP production [302]. Sertoli cells from nonobese diabetic mice have a reduced capacity to survive high doses of glucose concentrations compared with normal mice [303]. Insulin signaling is mediated through the insulin regulatory substrate (IRS) family of proteins. IRS-2 knockout males are adequate breeders until they develop diabetes and then become totally infertile [304]; however, the male phenotype was not studied in enough detail to allow comment on the mechanistic involvement of Sertoli cells. Similarly CDK4-deficient mice develop diabetes due to lack of proliferation of islet cells and some peripheral insulin resistance associated with infertility; however, the male factor infertility does not develop until after 4 months of age [305]. Again, it is unclear if the development of infertility in the male is due to a primary essential role for CDK4 in testes where it is expression includes Sertoli cells [306] or if it is due to the development of diabetes or both. These data may indicate that diabetes in good control may not be sufficient to cause a significant negative impact on Sertoli cell function, but poorly controlled diabetes with or without a second “hit” may indeed lead to male infertility, at least in rodents. This probably occurs through multiple mechanisms, some of which may involve Sertoli cell function. It is difficult to adequately study Sertoli cell function in men with poor diabetic control due to the scarcity and difficulty of obtaining testicular tissue. Animals models, however, do suggest that in spontaneously occurring or genetically engineered diabetes models, diabetes may affect Sertoli cell function, the mechanism of which remains to be determined. Another new exiting “link” between diabetes and Sertoli cells aims at the therapeutic utilization of the immune privileged properties of Sertoli cells to cure or improve Type 1 diabetic control [307–314]. This would be achieved either through xenotransplantation in insulinopenic diabetics of either genetically engineered Sertoli cells expressing insulin, or newborn pig islet and Sertoli cells into a subcutaneously implanted device. In both instances, the use of Sertoli cells would obviate the need to rely on immunosuppressive

Chapter 21 Conditions Affecting Sertoli Cells

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therapy for ensuring survival of the islet cells xenotransplant. One preliminary trial of the latter approach performed in humans shows promising results, although controversy still surrounds the interpretation of the data [308], and no independent replication of these results in higher mammals or humans has been done.

H. Radiation and Chemotherapy A significant number of young males suffering from testicular cancer, Hodgkin, non-Hodgkin lymphomas, and leukemia are subjected to radiation or chemotherapy or both prior to exerting their fertility potential. Annually in the United States, 1400 young adults receiving such therapy have residual azoospermia [315]. This number does not include pediatric cases, which are treated in the same fashion for similar malignancies. As previously discussed in the case of testicular cancer (Section III.B), and as in lymphoma, significant alterations in spermatogenesis may predate the institution of cytotoxic therapy. These alterations may be etiologically related to the development of testicular cancer or may be a by-product of the altered cytokine milieu associated with lymphoproliferative disease [315]. Both acute and chronic toxicity have been documented especially with cisplatin-based chemotherapeutic regimens. Chronic toxicity resulting in 50% of treated subjects becoming azoospermic is dose dependent (Table 21.5). It is beyond the scope of

TABLE 21.5 Doses of Cytotoxic Agents Causing Prolonged Azoospermia in 50% of Patientsa Group

Definite Gonadotoxicity

Dose

Alkylating agents

Cyclophosphamide

7.5 g/m2

Chlorambucil

1.4 g/m2

Mustine Melphalan Busulfan Carmustineb

0.5 g/m2

Lomustineb

1.0 g/m2

Antimetabolites

Cytarabine

Vinca alkaloids

Vinblastine

Others

Procarbazine

4.0 g/m2

Cisplatin

0.6 g/m2

aBased on the data of Meistrich, M.L. (1998). Hormonal stimulation of the recovery of spermatogenesis following chemo- or radiotherapy. Review article. Apmis. 106, 37–45. bBased on the treatment of prepubertal males; doses of other agents are based on the treatment of adult men.

FIGURE 21.15

Individual serum FSH levels in 232 patients with germ cell tumors followed up to 153 months after completion of chemotherapy. Dotted line indicates lower limit of normal range. (Adapted from Brennemann, W., et al. [316].)

this chapter to provide a detailed review of the impact of these agents on spermatogenesis or mechanistic explanations on how Sertoli cell damage occurs. We focus instead on evidence for an impact on Sertoli cell function in men. Azoospermia secondary to chemotherapy or radiation is probably multifactorial, but damage to the Sertoli cell may be implied by the elevated levels of FSH encountered in most subjects [315]. The elevation in FSH is observed in most patients at 6 months after chemotherapy. Recovery of spermatogenesis has been observed on occasion up to 8 years after chemotherapy, but a significant proportion will have a residual elevation of FSH (Fig. 21.15). The persistent elevation of FSH after therapy associated with a Sertoli cell only histology on testes biopsy is indicative of probable permanent damage and little chance for recovery. Suppressed inhibin B levels have been shown in some patients following chemotherapy [316, 317]. Earlier studies utilizing a less specific inhibin assay failed to show a significant drop of inhibin after chemotherapy [318, 319]. The impact of testicular irradiation on inhibin secretion depends on the dose of irradiation delivered to the testes. In men treated with testicular irradiation (14–20 Gy) for possible residual tumors after orchiectomy for unilateral CIS, inhibin levels were undetectable after 2–12 months of therapy [320]. Men treated with I131 for thyroid cancer showed a transient decrease in inhibin B levels at 3 and 6 months after therapy [321]. Although proliferating spermatogonia may be very sensitive to even small doses of radiation (<0.1 Gy), Sertoli cells require a higher dose (>0.2 Gy) for a transient or permanent rise in FSH to occur [322]. Children may be more prone to permanent Sertoli cell damage induced by radiation or chemotherapy since

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this treatment occurs during the period of Sertoli cell proliferation and might leave these patients with a reduced complement of Sertoli cells, leading to lower inhibin B levels and higher FSH levels during adulthood [323, 324]. Examination of Sertoli cell function after chemotherapy in prepubertal boys (age median of 4.5 yr), showed little effect on either inhibin B or FSH levels [325].

5.

6.

VIII. SUMMARY

7.

In this chapter, we have described the many clinical conditions affecting Sertoli cell function. Unfortunately, the quality of the clinical studies is quite variable and conclusions might seem at times contradictory. Nevertheless, all studies have as a common denominator and as an endpoint either a descriptive morphological change in Sertoli cell structure or an alteration in markers of Sertoli cell function. This level of knowledge is unfortunately insufficient to delineate where the pathways for Sertoli cell injury are distinct and where they merge. Although there may be a threshold beyond which Sertoli cell damage is irreversible, there is sufficient information to suggest that subtle insults may be associated with partial or complete recovery of Sertoli cell function. It is, thus, incumbent on the scientific community of basic and clinical andrologists to go beyond the observational. In this era of genomics, proteomics, and metabolomics, we should be able to start delineating the early as well late pathophysiological changes leading to alteration in Sertoli cell function in each of the discussed disease entities. Such knowledge will facilitate design of novel therapies and protective interventions to prevent irreversible damage from occurring. This is important, not only for the individuals afflicted with these conditions, but for our understanding of the global impact of environmental changes on male reproductive function and, thus, on the reproductive prospects and health of the species as a whole.

8.

9. 10. 11.

12. 13.

14.

15. 16.

17.

18. 19.

20.

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273. Kubini, K., Zachmann, M., Albers, N., Hiort, O., Bettendorf, M., Wolfle, J., Bidlingmaier, F., and Klingmuller, D. (2000). Basal inhibin B and the testosterone response to human chorionic gonadotropin correlate in prepubertal boys. J. Clin. Endocrinol. Metab. 85, 134–138. 274. Ozkan, K. U., Kucukaydin, M., Muhtaroglu, S., and Kontas, O. (2001). Evaluation of contralateral testicular damage after unilateral testicular torsion by serum inhibin B levels. J. Pediatr. Surg. 36, 1050–1053. 275. Anderson, R. A., and Baird, D. T. (2002). Male contraception. Endocr. Rev. 23, 735–762. 276. Orlando, C., Casano, R., Caldini, A. L., Forti, G., Barni, T., Bonfanti, L., and Serio, M. (1988). Measurement of seminal LDH-X and transferrin in normal and infertile men. J. Androl. 9, 220–223. 277. Fenichel, P., Rey, R., Poggioli, S., Donzeau, M., Chevallier, D., and Pointis, G. (1999). Anti-Müllerian hormone as a seminal marker for spermatogenesis in non-obstructive azoospermia. Hum. Reprod. 14, 2020–2024. 278. Liu, D. Y., Cooper, E. J., and Baker, H. W. (1986). Seminal transferrin, an index of Sertoli cell function: Is it of clinical value? Clin. Reprod. Fertil. 4, 191–197. 279. Garem, Y. F., Arini, A. F., Beheiry, A. H., Zeid, S. A., and Comhaire, F. H. (2002). Possible relationship between seminal plasma inhibin B and spermatogenesis in patients with azoospermia. J. Androl. 23, 825–829. 280. Foresta, C., Bettella, A., Petraglia, F., Pistorello, M., Luisi, S., and Rossato, M. (1999). Inhibin B levels in azoospermic subjects with cytologically characterized testicular pathology. Clin. Endocrinol. (Oxf). 50, 695–701. 281. Jarow, J. P., Budin, R. E., Dym, M., Zirkin, B. R., Noren, S., and Marshall, F. F. (1985). Quantitative pathologic changes in the human testis after vasectomy. A controlled study. N. Engl. J. Med. 313, 1252–1256. 282. Loughlin, K. R. (1988). Complications of vasovasostomy. Urol. Clin. North Am. 15, 243–248. 283. Hirsch, I. H., Sedor, J., Kulp, D., McCue, P. J., and Staas, W. E., Jr. (1994). Objective assessment of spermatogenesis in men with functional and anatomic obstruction of the genital tract. Int. J. Androl. 17, 29–34. 284. Hadley, M. A., and Dym, M. (1983). Spermatogenesis in the vasectomized monkey: Quantitative analysis. Anat. Rec. 205, 381–386. 285. Harris, J. D., and Lipshultz, L. I. (1981). The effect of vasectomy on Sertoli cell function in rats. Invest. Urol. 18, 305–307. 286. Croft, B. T., and Bartke, A. (1976). Quantitative study of spermatogenesis in vasectomized mice. Int. J. Fertil. 21, 61–64. 287. Singh, S. K., and Chakravarty, S. (2000). Histologic changes in the mouse testis after bilateral vasectomy. Asian J. Androl. 2, 115–120. 288. Bigazzi, P. E., Kosuda, L. L., Hsu, K. C., and Andres, G. A. (1976). Immune complex orchitis in vasectomized rabbits. J. Exp. Med. 143, 382–404. 289. Aitken, H., Kumarakuru, S., Orr, R., Reid, O., Bennett, N. K., and McDonald, S. W. (1999). Effect of long-term vasectomy on seminiferous tubules in the guinea pig. Clin. Anat. 12, 250–263. 290. Stremmel, W., Niederau, C., Berger, M., Kley, H. K., Kruskemper, H. L., and Strohmeyer, G. (1988). Abnormalities in estrogen, androgen, and insulin metabolism in idiopathic hemochromatosis. Ann. NY Acad. Sci. 526, 209–223. 291. Vogt, H. J., Weidenbach, T., Marquart, K. H., and Vogel, G. E. (1987). Idiopathic hemochromatosis in a 45-year-old infertile man. Andrologia 19, 532–538. 292. Nagai, T., and Mori, M. (1999). Prader-Willi syndrome, diabetes mellitus and hypogonadism. Biomed. Pharmacother. 53, 452–454.

293. Goldstone, A. P. (2004). Prader-Willi syndrome: Advances in genetics, pathophysiology and treatment. Trends Endocrinol. Metab. 15, 12–20. 294. Sexton, W. J., and Jarow, J. P. (1997). Effect of diabetes mellitus upon male reproductive function. Urology 49, 508–513. 295. Cameron, D. F., Murray, F. T., and Drylie, D. D. (1985). Interstitial compartment pathology and spermatogenic disruption in testes from impotent diabetic men. Anat. Rec. 213, 53–62. 296. Faerman, I., Vilar, O., Rivarola, M. A., Rosner, J. M., Jadzinsky, M. N., Fox, D., Lloret, A. P., Bernstein-Hahn, L., and Saraceni, D. (1972). Impotence and diabetes. Studies of androgenic function in diabetic impotent males. Diabetes 21, 23–30. 297. Salardi, S., Zucchini, S., Cicognani, A., Gualandi, S., Barbieri, E., and Cacciari, E. (2002). Inhibin B levels in adolescents and young adults with type 1 diabetes. Horm. Res. 57, 205–208. 298. Perez Diaz, J., Benitez, A., and Fernandez Galaz, C. (1982). Effect of streptozotocin diabetes on the pituitary-testicular axis in the rat. Horm. Metab. Res. 14, 479–482. 299. Hutson, J. C., Stocco, D. M., Campbell, G. T., and Wagoner, J. (1983). Sertoli cell function in diabetic, insulin-treated diabetic, and semi-starved rats. Diabetes 32, 112–116. 300. Murray, F. T., Cameron, D. F., Orth, J. M., and Katovich, M. J. (1985). Gonadal dysfunction in the spontaneously diabetic BB rat: Alterations of testes morphology, serum testosterone and LH. Horm. Metab. Res. 17, 495–501. 301. Cameron, D. F., Rountree, J., Schultz, R. E., Repetta, D., and Murray, F. T. (1990). Sustained hyperglycemia results in testicular dysfunction and reduced fertility potential in BBWOR diabetic rats. Am. J. Physiol. 259, E881–889. 302. Hutson, J. C. (1984). Altered biochemical responses by rat Sertoli cells and peritubular cells cultured under simulated diabetic conditions. Diabetologia 26, 155–158. 303. Gondos, B., Rivkind, Y., and Jovanovic, L. (1998). Effect of increasing glucose concentrations on Sertoli cell viability in the nonobese diabetic mouse. Ann. Clin. Lab. Sci. 28, 236–241. 304. Burks, D. J., de Mora, J. F., Schubert, M., Withers, D. J., Myers, M. G., Towery, H. H., Altamuro, S. L., Flint, C. L., and White, M. F. (2000). IRS-2 pathways integrate female reproduction and energy homeostasis. Nature 407, 377–382. 305. Mettus, R. V., and Rane, S. G. (2003). Characterization of the abnormal pancreatic development, reduced growth and infertility in Cdk4 mutant mice. Oncogene 22, 8413–8421. 306. Rhee, K., and Wolgemuth, D. J. (1995). Cdk family genes are expressed not only in dividing but also in terminally differentiated mouse germ cells, suggesting their possible function during both cell division and differentiation. Dev. Dyn. 204, 406–420. 307. Check, E. (2002). Diabetes trial stirs debate on safety of xenotransplants. Nature 419, 5. 308. Birmingham, K. (2002). Skepticism surrounds diabetes xenograft experiment. Nat. Med. 8, 1047. 309. Luca, G., Calvitti, M., Neri, L. M., Becchetti, E., Capitani, S., Basta, G., Angeletti, G., Fanelli, C., Brunetti, P., and Calafiore, R. (2000). Sertoli cell-induced reversal of adult rat pancreatic islet beta-cells into fetal-like status: Potential implications for islet transplantation in type I diabetes mellitus. J. Investig. Med. 48, 441–448. 310. Dufour, J. M., Rajotte, R. V., Korbutt, G. S., and Emerich, D. F. (2003). Harnessing the immunomodulatory properties of Sertoli cells to enable xenotransplantation in type I diabetes. Immunol. Invest. 32, 275–297. 311. Korbutt, G. S., Elliott, J. F., and Rajotte, R. V. (1997). Cotransplantation of allogeneic islets with allogeneic testicular cell aggregates allows long-term graft survival without systemic immunosuppression. Diabetes 46, 317–322.

Chapter 21 Conditions Affecting Sertoli Cells 312. Dufour, J. M., Rajotte, R. V., Kin, T., and Korbutt, G. S. (2003). Immunoprotection of rat islet xenografts by cotransplantation with Sertoli cells and a single injection of antilymphocyte serum. Transplantation 75, 1594–1596. 313. Emerich, D. F., Hemendinger, R., and Halberstadt, C. R. (2003). The testicular-derived Sertoli cell: Cellular immunoscience to enable transplantation. Cell Transplant. 12, 335–49. 314. Dufour, J. M., Hemendinger, R., Halberstadt, C. R., Gores, P., Emerich, D. F., Korbutt, G. S., and Rajotte, R. V. (2004). Genetically engineered Sertoli cells are able to survive allogeneic transplantation. Gene Ther. 11, 694–700. 315. Schrader, M., Muller, M., Straub, B., and Miller, K. (2001). The impact of chemotherapy on male fertility: A survey of the biologic basis and clinical aspects. Reprod. Toxicol. (City) 15, 611–617. 316. Brennemann, W., Stoffel-Wagner, B., Bidlingmaier, F., and Klingmuller, D. (1992). Immunoreactive plasma inhibin levels in men after polyvalent chemotherapy of germinal cell cancer. Acta Endocrinol. (Copenh.). 126, 224–228. 317. Wallace, E. M., Groome, N. P., Riley, S. C., Parker, A. C., and Wu, F. C. (1997). Effects of chemotherapy-induced testicular damage on inhibin, gonadotropin, and testosterone secretion: A prospective longitudinal study. J. Clin. Endocrinol. Metab. 82, 3111–3115. 318. Tsatsoulis, A., Shalet, S. M., Robertson, W. R., Morris, I. D., Burger, H. G., and De Kretser, D. M. (1988). Plasma inhibin levels in men with chemotherapy-induced severe damage to the seminiferous epithelium. Clin. Endocrinol. (Oxf). 29, 659–665.

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319. Tsatsoulis, A., Shalet, S. M., Morris, I. D., and de Kretser, D. M. (1990). Immunoactive inhibin as a marker of Sertoli cell function following cytotoxic damage to the human testis. Horm. Res. 34, 254–259. 320. Petersen, P. M., Andersson, A. M., Rorth, M., Daugaard, G., and Skakkebaek, N. E. (1999). Undetectable inhibin B serum levels in men after testicular irradiation. J. Clin. Endocrinol. Metab. 84, 213–215. 321. Wichers, M., Benz, E., Palmedo, H., Biersack, H. J., Grunwald, F., and Klingmuller, D. (2000). Testicular function after radioiodine therapy for thyroid carcinoma. Eur. J. Nucl. Med. 27, 503–507. 322. Howell, S., and Shalet, S. (1998). Gonadal damage from chemotherapy and radiotherapy. Endocrinol. Metab. Clin. North Am. I (City) 27, 927–943. 323. Schmiegelow, M., Lassen, S., Poulsen, H. S., Schmiegelow, K., Hertz, H., Andersson, A. M., Skakkebaek, N. E., and Muller, J. (2001). Gonadal status in male survivors following childhood brain tumors. J. Clin. Endocrinol. Metab. 86, 2446–2452. 324. Cicognani, A., Cacciari, E., Pasini, A., Burnelli, R., De Iasio, R., Pirazzoli, P., and Paolucci, G. (2000). Low serum inhibin B levels as a marker of testicular damage after treatment for a childhood malignancy. Eur. J. Pediatr. 159, 103–107. 325. Crofton, P. M., Thomson, A. B., Evans, A. E., Groome, N. P., Bath, L. E., Kelnar, C. J., and Wallace, W. H. (2003). Is inhibin B a potential marker of gonadotoxicity in prepubertal children treated for cancer? Clin. Endocrinol. (Oxf). 58, 296–301.

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VIII SPERMATOGONIAL STEM CELLS

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22 Gonocyte Development and Differentiation PETER J. DONOVAN AND MARIA P. DE MIGUEL Stem Cell Program, Institute for Cell Engineering, Johns Hopkins University, School of Medicine, Baltimore, Maryland

germline progenitors, so-called primordial germ cells (PGCs), is achieved by movement of these cells into the gonads. Once in the gonads, the PGCs proliferate for a short period of time before entering a mitotic arrest in males or entering meiotic prophase and then meiotic arrest in females (Fig. 22.1). The germ cells, now called gonocytes, remain in these arrest states until after birth. In the testis, the gonocytes will resume mitosis in the first few days after birth depending on the species. At that first cell division the gonocyte gives rise to spermatogonia, the stem cells of the adult testis. Although gonocytes have over the years received little attention, they are increasingly seen as representing an important stage in germline development. Rather than being viewed as a quiescent period in which germ cells lie in wait for the onset of spermatogenesis, this stage of germ cell development can be viewed as an active and important period of differentiation. As the progenitors of the stem cells of the adult, gonocytes provide the necessary foundation for adult fertility. Failure of gonocytes to survive correctly in the developing gonad can result in complete sterility. Alternatively, other studies suggest that failure of gonocytes to differentiate correctly in the fetal and postnatal gonad can result in the development of testicular cancer, the most common cause of cancer in young men. Improper descent of the testes (cryptorchidism) can affect the differentiation of gonocytes and lead to the development of testicular cancer. The vulnerability of this population of germ cells in late fetal and early postnatal life is being realized and the impact of toxicants

I. II. III. IV. V. VI.

INTRODUCTION A BRIEF HISTORY OF THE GERMLINE FORMATION OF PGCS FROM PGCS TO GONOCYTES FROM GONOCYTES TO SPERMATOGONIA PGCS, GONOCYTES, AND THE ORIGIN OF TESTICULAR CANCER VII. ANALYZING AND MANIPULATING GENE EXPRESSION IN THE MAMMALIAN GERMLINE VIII. CONCLUSIONS References

I. INTRODUCTION The survival of animal species is dependent on their ability to pass their genome on to the next generation. In mammals, as in all vertebrate and many invertebrate species, this task is accomplished by a small population of highly specialized cells termed germ cells. The fusion of two germ cells, the gametes, at fertilization creates a cell capable of recapitulating the entire program of development, thereby creating a new organism. The formation of the gametes in the adult could be viewed as the end stage of a long process of germline development that begins in the embryo and continues through fetal and postnatal life. The mammalian germ cell population is formed early in postimplantation development and arises outside of the gonad anlagen. Colonization of the gonad by the SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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Copyright 2005, Elsevier Science (USA). All rights reserved.

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FIGURE 22.1 Schematic representation of germ cell development from PGC to spermatogonia in the mouse. PGCs are first identified at 7.2 dpc by tissue-nonspecific alkaline phosphatase and Oct-4 expression, as they migrate toward the developing gonad. At 10.5 dpc they also express c-kit. c-Kit expression continues until the gonocyte stage in the gonad becomes mitotically arrested at 14.5 dpc. They then lose alkaline phosphatase and Oct-4 expression, and begin to express vasa (Mvh) and germ cell nuclear antigen (GCNA). These two markers will continue to be expressed in all germ cell types until completion of meiosis. Gonocytes become migratory again at 1 dpp, in order to migrate from the center of the seminiferous cords to the basement membrane. Note the large increase in size of gonocytes at this time. c-Kit is reexpressed when the gonocytes reenter mitosis and form type A spermatogonia.

and teratogens on these cells is being scrutinized in earnest. In the last decade genetic studies in mice utilizing knockout models have begun to elucidate the genetic pathways affecting PGC and gonocyte development. Similarly, molecular cloning of mutant loci associated with infertility in human populations has provided a unique insight into germline development in humans and an approach that is complementary to the genetic studies in mice. The application of genome-wide expression analysis to all stages of germ cell development also promises to provide unique new insights into gonocyte development and differentiation. At the same time techniques for manipulating gene expression in germ cells and their surrounding somatic cells have improved sufficiently such that it is now feasible to envisage functional studies on genes identified in genome-wide expression screens as potential regulators of germ cell development. This chapter focuses on outlining our current knowledge of gonocyte development and differentiation as well as looking to future developments in this field.

II. A BRIEF HISTORY OF THE GERMLINE Mammalian PGCs can be traced in the embryo by staining embryo sections for tissue-nonspecific alkaline phosphatase (TNAP), by antigens recognized by rabbit polyclonal antisera and mouse and rat monoclonal antibodies [1–4], and by the expression of the POU domain transcription factor Oct4. That these TNAP-positive, Oct4-positive cells are indeed PGCs is confirmed by the fact that these cells are deficient or absent in mouse mutants that are sterile [5–7]. PGCs have been traced in sections of mouse embryos by means of antibodies to carbohydrate differentiation antigens, so-called stage-specific embryonic antigens (SSEAs), and by histochemical staining for TNAP. PGCs arise outside of the gonad anlagen, and colonization of the gonad is brought about partly through the morphogenetic movements of the embryo and partly through active directed migration [1]. During this period of migration and gonad colonization, germ cell numbers increase. In the mouse embryo the earliest

Chapter 22 Gonocyte Development and Differentiation

identifiable population of PGCs is small (8 to 10 cells) [3, 4, 8, 9]. By the time the embryonic gonad is fully colonized by PGCs [approximately 12.5 days postcoitus (dpc) in the mouse] the population of PGCs is greatly increased and is estimated to be about 25,000 to 30,000 cells [1, 10]. In male embryos, once the PGCs have reached the embryonic gonad they cease proliferation and enter a period of cell cycle arrest, at which time they are referred to as gonocytes [1]. This phase of differentiation is also marked by loss of expression of some of the SSEAs including SSEA-1. In the next phase of normal differentiation, the gonocytes move between adjacent Sertoli cells and become located in the center of the seminiferous cords. At the same time, gonocytes lose markers expressed by PGCs, a process that begins at around 13.5 dpc in the mouse [2, 11]. In rodents, the gonocytes will remain in cell cycle arrest until a few days after birth depending on the species, at which time they will resume mitosis (the so-called onset of spermatogenesis). At this time, gonocytes extend processes toward the basal lamina and relocate themselves peripherally between the cords in a position apposed to the basal lamina [12]. Relocation to the basement membrane is necessary for their survival, because those remaining in the center of the testis cords eventually degenerate [13]. Gonocytes located at the basement membrane will divide mitotically to form the stem cells of the testis, the spermatogonia (for reviews, see [14, 15]). The transition from gonocytes to spermatogonia in the first wave of spermatogenesis is still not well characterized. It has been hypothesized that, at the start of spermatogenesis, gonocytes give rise to either A stem (As) spermatogonia or directly to A2 spermatogonia [16]. In the adult, the As cell will divide to give rise to A paired (Apr) and A aligned (Aal) undifferentiated spermatogonia, which remain connected by intercellular bridges. Then Aal spermatogonia will differentiate into A1 spermatogonia, which are the first generation of differentiating type A spermatogonia. These differentiating spermatogonia go through a series of six divisions, via A2, A3, A4, In, and B spermatogonia, that give rise to spermatocytes (for reviews, see [14, 17]). The preceding description provides some information on how the various stages of germ cell development in a male mammal are linked, but provides little information about the mechanisms regulating growth and differentiation. Information regarding the mechanisms regulating these processes has come from genetic studies defining loci associated with sterility, which by definition identify genes involved in one or other aspect of gametogenesis. A second set of data has come from in vitro studies in which some of the factors regulating germ cell survival, proliferation,

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and differentiation have been identified. Defining the timing and extent of germ cell proliferation are important because this information is vital to any meaningful understanding of how and when germ cell development is controlled in mammals.

III. FORMATION OF PGCS The formation of PGCs occurs early in postimplantation development in mammals. Because the PGC population gives rise to the gonocyte population, factors affecting PGC numbers can affect the formation of the gonocyte pool. Therefore, in considering gonocytes, it is worth noting how the PGC pool is formed and what factors affect that cell population. In mice, fate mapping studies have identified the PGCs progenitors as a small population of approximately 8 to 10 cells as early as 6.5 dpc [3, 4, 9]. Some of the molecular mechanisms regulating the formation of the germline have been defined. Importantly, bone morphogenetic protein 4 (BMP4), produced in the extraembryonic mesoderm, plays a critical role in induction of PGCs in the epiblast. Mice lacking BMP4 are severely deficient in PGCs. Interestingly, the effects of BMP4 can be recapitulated in culture and BMP4 can induce germ cell formation from epiblast cells in culture. Another member of the BMP family, BMP8b, also plays a critical role at this time and, again, mice lacking BMP8b are severely deficient in PGC numbers. Because the PGC pool is reduced, the numbers of germ cells that are available to form gonocytes and then spermatogonia is compromised. Consequently, male mice lacking BMP8b are sterile. These data point to an important role for BMPs during many stages of germ cell development, but raise the interesting question of how the signals from a variety of BMPs are integrated to regulate germ cell development. During the next phase of development, PGCs will move toward the developing embryonic gonad. Migration to the gonad in the first phase is thought to be brought about by morphogenetic movements of the embryo such that germ cells are brought into the region of the hindgut through invagination of that structure [1]. In the next phase of development, germ cells actively migrate from the hindgut epithelium toward the genital ridges or gonad anlagen (Fig. 22.2). The factors controlling germ cell migration are complex. Some studies suggest that germ cell interaction with the extracellular matrix is important for gonad colonization. Consistent with this notion, extracellular matrix molecules such as fibronectin, laminin, and type IV collagen are expressed by the coelomic epithelial

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FIGURE 22.2 Bright-field microphotograph of 8.5-dpc mouse PGCs isolated and cultured and labeled with BrdU (brown) and alkaline phosphatase (pink) to differentiate between proliferating (arrowheads) and quiescent (arrows) PGCs in culture.

cells on which germ cells migrate. Moreover, germ cells change adhesiveness to laminin and fibronectin during this period. Importantly, germ cells express a β1 integrin on their cell surface. Integrins act as cell surface receptors and bind to extracellular ligands, typically components of the extracellular matrix. This class of receptor occurs in two forms, α and β each with a large number of isoforms. Heterodimers of various α and β isoforms act as receptors for different ligands including fibronectin and laminin. For example, α3β1, α6β1, and α6β1 act as laminin receptors. The importance of β1 integrin for germ cell migration is indicated by the reduced number of germ cells in mice lacking β1 integrin [18]. However, germ cells are not completely absent, suggesting that other mechanisms also play a role in germ cell migration. Mice lacking α3 and α6 showed no defects in germ cell migration, suggesting that these components are not part of a putative germ cell receptor for laminin [18]. Therefore, at present it is unclear what heterodimeric partners are associated with β1 integrin and what the putative ligand is. Until recently, evidence for chemotactic guidance of germ cells to the gonad was scarce. Recent studies, carried out in both zebra fish and mice, have provided strong evidence for the role of a chemokine in chemotaxis in germ cell migration. Stromal derived factor 1 (SDF-1) is a chemokine that binds to the AIDS coreceptor CXCR4, a G-protein coupled receptor. Genetic screens in zebra fish identified SDF-1 as an important germ cell chemoattractant. In mice, germ cells express CXCR4 on their cell surface, and mice lacking SDF-1 show defects in germ cell colonization of the gonad [19, 20]. During this period of gonad colonization PGC numbers increase. By 10.5 dpc approximately 1000 PGCs are identified in the mouse embryo [5]. In the fully colonized gonad at 12.5 dpc 25,000 germ cells are estimated to be present (for a review, see [1]). The population doubling time of PGCs has been estimated to be around 16 hr [10]. When PGCs are isolated into

FIGURE 22.3 Hematoxylin-stained histological section of the aorta–gonad–mesonephros region of an 11.5-dpc mouse showing the aorta (arrowheads), the developing gonad (arrow), and the rudimentary mesonephros.

culture during the time period in which they are migrating toward, the mouse gonad (8.5–11.5 dpc), they continue to proliferate and incorporate 3H-thymidine or BrdU (Fig. 22.3) [2, 21, 22]. Gonocytes isolated from 12.5-dpc or older embryonic mouse gonads show a decreased ability to proliferate [2]. Thus, their in vitro proliferation seems to mirror their in vivo proliferation, perhaps suggesting that PGC development is regulated by a clock mechanism that times developmental events as has been suggested in other cell lineages. Taken together these data suggest that the transition from a PGC to a gonocyte involves not only changes in the cell cycle but an ongoing program of differentiation [2, 23, 24]. Some of the factors controlling PGC survival and proliferation during the period of gonad colonization have been identified in genetic studies and in cell culture experiments. Among the factors that act on PGCs, the best studied are the c-kit receptor tyrosine kinase, encoded by the murine Dominant White Spotting locus on chromosome 5, and the ligand for the c-kit receptor, kit ligand (KL), encoded by the murine Steel (Sl) locus on chromosome 10 ([25–27], for a review, see [28]). Mice homozygous for many mutations at the Sl and W loci are sterile, demonstrating the critical role for KLmediated activation of c-kit signaling in PGC survival (for a review, see [29]). In this context, note that PGCs express c-kit during their migration toward the embryonic gonad, and that kit ligand is expressed by the cells along the migratory path, indicating that KL is required for PGC survival during this period [30]. Recent studies have shown that KL restores telomerase activity in mouse mitotic PGCs in culture, suggesting that one of the features of pluripotent cells is maintained in PGCs during this period of development [31]. The effect of the W and Sl mutations can be detected as early as 9.5 dpc. That PGC numbers are not affected at earlier stages suggests that PGC development prior to this stage is c-kit independent and points

Chapter 22 Gonocyte Development and Differentiation

to a dramatic change in PGC growth factor dependence at 9.5 days of development. In vitro culture systems have been used extensively to both identify and to study the factors that regulate PGC growth. Isolation and culture of PGCs has been described extensively elsewhere [2, 32]. Briefly, the most successful and most commonly used method involves culturing PGCs on a confluent fibroblast feeder layer. In this system, the PGCs can be identified using the classical germ cell markers such as TNAP and SSEA-1 [2]. Many of the factors that have been identified using this system have been subsequently shown to be relevant to PGC growth in vivo. Moreover, culture systems have also been useful for studying other aspects of PGC development in addition to growth regulation, especially PGC differentiation and migration [2, 33, 34]. PGCs are only able to survive in vitro in coculture with feeder cells that produce KL [2, 21, 22, 34, 35] consistent with the in vivo situation. Importantly the in vitro studies revealed a distinction between different isoforms of KL. The KL factor is a transmembrane growth factor produced in two basic forms. One form has a cleavage site in the ectodomain that can be cleaved to produce a soluble factor. Another form lacks the proteolytic cleavage site and remains as a transmembrane growth factor. Importantly, long-term survival of PGCs in culture is dependent on the full-length transmembrane growth factor. When PGCs are cultured on cells that only produce soluble forms of KL, they cannot survive for more than 24 hr [22]. These data nicely explain the observation that in Sldickie (Sld) mice, which can only produce a soluble form of KL, germ cells die and the animals are sterile [36, 37]. The realization that KL was a survival factor led to the identification of other factors that act as PGC mitogens, including members of the interleukin 6 (IL-6) family of growth factors. Most likely members of the IL-6 family of cytokines play an important role in PGC survival and, together with KL, promote PGC proliferation. A member of the IL-6 cytokine family, leukemia inhibitory factor (LIF), has been shown to increase PGC survival in vitro and, in combination with KL, induces proliferation [2, 21, 22, 34, 35, 38, 39]. Other members of this cytokine family, ciliary neurotrophic factor (CNTF) and oncostatin M (OSM), have also been shown to increase mouse PGC survival [40, 41], whereas IL-6 itself does not affect PGC growth in vitro [38]. These cytokines have distinct ligand-binding subunits, but notably activate intracellular signaling via a shared signaling component, gp130. Importantly, mice lacking the gp130 signaling component are reported to show reduced PGC numbers (Yoshida and Taga, unpublished data, cited in [42]). In addition, incubation of

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cultured PGCs with an anti-gp130 antibody leads to PGC apoptosis [42]. Recently, further examination of PGC numbers in gp130 knockout mice revealed a less drastic, but significant, reduction in PGC numbers in male but not in female embryos at 13.5 dpc [43]. These data demonstrate that signaling via gp130 is required for some aspect of male PGC growth but not necessarily for PGC survival. Some studies have suggested that gp130 signaling may inhibit entry of male germ cells into meiosis in the developing gonad. Male PGCs isolated from embryos begin to express meiosis-specific genes in culture, indicating that when germ cells are removed from the male gonadal environment they enter meiosis autonomously. In culture, appearance of meiosis-specific markers is suppressed by LIF, suggesting that gp130-mediated signaling blocks entry into meiosis. Indeed expression of LIF in the developing gonads between 11.5 and 13.5 dpc seems to be stronger in male than female gonads assayed by reverse transcriptase–polymerase chain reaction (RT-PCR). Although factors such as KL might be seen as simple survival factors or mitogens, the situation might be more complex. A number of studies point to a role for KL in regulating PGC migration. One study analyzed PGC migration in compound heterozygous embryos carrying one copy of the original Sl mutation (a null allele) and a copy of the Sld allele, which can only produce a soluble form of KL. In these animals 60% of PGCs were found migrating off the normal migration pathway, suggesting that KL might be a component of the migratory pathway that guides PGCs to the embryonic gonad [5]. Similar studies analyzed PGC migration in compound heterozygous mice carrying one allele of the original W mutation and the other the Wextreme (We) mutation. In these embryos PGCs were found to be clumped together at 9.5 dpc, suggesting that expression of the c-kit receptor might be required for adhesion of PGCs to somatic cells and commencement of PGC migration [44]. The failure of PGCs to develop in W/We mice results in adult sterility. The interpretation of the actions of these factors is also complicated by the fact that agents that appear to act as chemoattractants, such as SDF-1, also appear to affect PGC numbers, suggesting that they also act as mitogens. Growth of germ cells at any stage of development may not be controlled simply by growth factors. Adhesion molecules such as E-cadherin [45] are also implicated in PGC in vitro proliferation, but their role in regulating PGC numbers in vivo remains unclear. As described earlier, PGCs express b1 integrin on their cell surface during migration into the developing gonad, and in mice lacking b1 integrin, PGC numbers are reduced. It seems likely that germ cell growth and survival are controlled very strictly temporally and

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spatially by a variety of factors including cell adhesion molecules, extracellular matrix components, growth factors, and cytokines. These overlapping mechanisms for regulating growth and migration could provide redundant pathways to regulate PGC migration, survival, and proliferation to ensure that PGCs only survive and proliferate at the right time and in the right place. In this way the animal also ensures that the correct number of PGCs will reach the developing gonad, while at the same time ensuring that PGCs do not survive and proliferate elsewhere. Other growth factors such as tumor necrosis factor α (TNFα) [46], IL-4 [47], epidermal growth factor (EGF) (Cooke, unpublished data, cited in [48]), basic fibroblast growth factor (bFGF) [35, 49], retinoic acid [50], and IL-11 [42] have all been shown to stimulate PGC growth in vitro. In addition, forskolin and cholera toxin, potent cAMP agonists, have also been shown to increase survival and proliferation of PGCs in vitro (Donovan, unpublished observations [51]). The significance of these data are unclear, however, because of the difficulty involved with interpreting effects of growth factors in a feeder-dependent culture system and because of the paucity of in vivo evidence supporting the role of these factors in regulating germline development. In contrast, the Sky receptor tyrosine kinase is expressed on PGCs after 11.5 dpc and Gas 6 (a soluble ligand for Sky) stimulates PGC proliferation in culture [52]. Similarly, members of the transforming growth factor β (TGFβ) superfamily, specifically BMP2, BMP4, and BMP8b [53, 54] have all been demonstrated to increase PGC growth in vitro. Some of these factors have been deleted in mouse knockout models, providing interesting insights into their function. As described earlier, embryos homozygous for a null mutation in the BMP4 gene were found to lack germ cells [8, 9]. Moreover, BMP4 has been shown to induce a PGC phenotype in a subset of epiblast cells [54, 55]. Similar to BMP4 null mutants, BMP8b null embryos have severe defects in PGC development [53, 56]. Additionally, BMP2-homozygous-null and heterozygous embryos have reduced numbers of PGCs, suggesting a role for BMP2 in PGC generation [57]. Taken together these data suggest that BMPs play significant roles at different stages of germ cell development. Consistent with this notion, targeted deletion of some of the downstream effectors of the BMP signal transduction pathway, the SMADs, leads to germ cell loss in mice [58–60]. An important question is how the signals emanating from the BMP receptors act differently at different stages of germ cell development to control germ cell growth and development.

Recently, a peptidyl-propyl isomerase, Pin1, has been demonstrated to regulate PGC proliferation as early as 8.5 dpc, because Pin null mice show a decrease in PGC numbers and a prolonged cell cycle [61]. These are important observations because we know so little about how the cell cycle is controlled in germ cells and how the transition from a mitotic PGC to a quiescent gonocyte is controlled by cell cycle regulators. Using gene knockout strategies in mice, a number of other genes have also been demonstrated to be important for PGC survival or proliferation such as TIAR [62], the Fanconi anemia complementation group c gene (Fancc) (Fanconi anemia, [63]) and POG [64, 65]. On the other hand, other factors have been found to have no effect on PGC survival in culture. These include insulin-like growth factor 1 (IGF-1), IL-1, IL- 3, plateletderived growth factor (PDGF) [38], and β nerve growth factor (β-NGF) [49]. At first glance this in vitro data seem somewhat meaningless except that this information provides an important contrast to data obtained from studies of cultured gonocytes (see later discussion). Finally, other factors have been described to inhibit PGC proliferation in vitro and include TNFβ, TGFβ1 [66], and activin [23]. The fact that TGFβ1 itself inhibits PGC proliferation, but other members of the TGF superfamily, such as BMP4, promote PGC proliferation has led to the hypothesis that this family of growth factors controls PGC numbers through a balance of positive and negative signals [54]. The transition from a PGC to a gonocyte is accompanied by dramatic changes in growth factor dependence and in cell cycle kinetics, indicating that this transition is likely marked by dramatic changes in the balance of such positive and negative regulators. Defining the molecules that control gonocyte proliferation and growth will be important because of the fundamental place occupied by gonocytes in gametogenesis.

IV. FROM PGCS TO GONOCYTES The transition from a PGC to a gonocyte is most notably marked by a decrease in germ cell proliferation and correlates with a decrease in the levels of expression of the c-kit receptor tyrosine kinase (see later discussion and [67, 68]; for a review, see [28]) (Fig. 22.1). Downregulation of c-kit levels in gonocytes occurs at about the same time as sexual differentiation is detected in the somatic cells of the gonad as evidenced by the formation of testis cords (Fig. 22.4). At the same time gonocytes begin to change surface antigens and begin to express intracellular antigens that distinguish them from PGCs (Fig. 22.1). For example gonocytes

Chapter 22 Gonocyte Development and Differentiation

FIGURE 22.4 Section of a human fetal testis at 20 weeks of gestation, showing the gonocytes (thin arrows) inside the seminiferous cords, which are surrounded by peritubular cells (thick arrows). Fetal Leydig cells (star) are located between the seminiferous cords inside the seminiferous cords. Stain: hematoxylin.

gradually lose expression of SSEA-1, which is expressed by PGCs. In contrast, they begin to express the germ cell nuclear antigen 1 (GCNA-1), an antigen of unknown function, recognized by a rat monoclonal antibody. The germ cells also lose adhesiveness to fibronectin and laminin [11, 69] and at the same time increase in cell size more than fourfold [70] (Fig. 22.1). The proliferation rate of embryonic mouse gonocytes decreases dramatically after they reach the embryonic testicular cords as described earlier. At 14 dpc, 7.7% of the mouse gonocytes are estimated to be proliferating, whereas this number drops to 0.2% at 16 dpc.1 From 16 dpc until birth, the gonocytes are arrested at the G1 phase of the cell cycle [71] (Fig. 22.1). In the rat, this process is similar, but these events take place 2 days later than in the mouse. For example, 5% of rat gonocytes are proliferating at 16 dpc, 5.5% at 17dpc, and 1% at 18 dpc [72]. In mammals, testis development is initiated in the embryo by expression of the sex-determining gene, Sry, in Sertoli cell precursors. Subsequently, Sertoli cells play a central role in male-specific cell interactions [73]. Expression of Sry in Sertoli cell precursors results in a cascade of signaling pathways that leads to complete differentiation of Sertoli cells and the recruitment and differentiation of other supporting cells including peritubular myoid cells and Leydig cells. Gonocytes become engulfed by Sertoli cells in the first phase of testis differentiation. Sertoli cells are then themselves surrounded by precursors of the peritubular myoid cells that migrate into the gonad from the adjacent mesonephric region, effectively enclosing Sertoli cells and forming seminiferous tubules (Fig. 22.4). 1 For the purposes of this chapter, to make the numbers derived from various studies comparable, we have adjusted the proliferation rates to the percentage of cells labeled after 1 hr of either 3H-thymidine or BrdU exposure.

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Leydig cells are interstitial cells that lie between the seminiferous tubules and can regulate Sertoli cell function. The idea that Sertoli cells control the behavior of gonocytes is supported by the fact that both male and female PGCs show autonomous entry into meiosis if they are set apart from the male gonadal environment [74–77]. These data suggest that specific factor(s) expressed by embryonic Sertoli cells are responsible for inhibition of meiosis in gonocytes in the embryonic testis. Several key genes of spermatogenesis are expressed in a male-specific fashion, such as OSM and desert hedgehog (Dhh), which are strongly expressed by somatic cells in the male embryonic gonad but not in the female (Fig. 22.5) (De Miguel and Donovan, unpublished observations [78]). A number of other genes have been shown to play a key role in the sex differentiation of the gonads, such as Cresp/testatin [79, 80], Sox9, DAX-1, Wilm’s tumor 1 (WT1), and Ad4BP/SF-1 (reviewed in [81]). In the fetal ovary, AMH (anti-Müllerian hormone) reduces the number of oogonia and even provokes sex reversal in early female embryos [82]. However, after 13.5 dpc, female germ cells are committed to meiosis, so even if transplanted into male genital ridges they will enter meiosis [83], suggesting that the molecule that prevents male germ cell entry into meiosis is not AMH, but another molecule downstream of Sry. Because correct formation of the testis cord is required for gonocyte survival, defects in many aspects of cord formation can result in loss of germ cells and complete sterility. For example, the WT1 and steroidogenic factor 1 (SF-1) genes are expressed very early in testis development and WT1 and SF-1 null mice completely fail to form gonads [84]. Genital ridges start to form, but then degenerate, provoking germ cell death at the time of their arrival into the degenerating gonad [84, 85]. Similarly, retinoic acid receptor (RAR) mutants also show gonad agenesis. Nonetheless, due to the presence of numerous RAR isoforms, as well as suspected redundancy, the phenotypes of the different isoform knockouts vary from severe reproductive defects to minimal abnormalities. Mice that are deficient for the RAR-γ2 isoform are fertile; however, males deficient for all RAR-γ isoforms are sterile [86]. Similarly, RAR-α1-null mice appear normal, but males deficient for all RAR-α isoforms are sterile and have severe degeneration of the testis [87]. Correct interaction of Sertoli cells with developing germ cells is essential for germ cell survival. The establishment of Sertoli cell–gonocyte interactions involves, in part, the formation of gap junction communication between these two cell types. In the testis the most abundant gap junction protein is connexin 43 (Cx43).

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A

B

FIGURE 22.5 (A) Histological section of a 14.5-dpc mouse testis showing Sertoli cells identified in brown with anti-OSM immunohistochemistry and unlabeled gonocytes in the seminiferous cords (arrows). Stain: hematoxylin and anti-OSM/DAB. (B) Histological section of a 14.5-dpc mouse ovary showing somatic cells slightly positive for OSM and unlabeled gonocytes (arrows). Stain: hematoxylin and anti-OSM/DAB.

Cx43 protein is expressed in Sertoli cell projections that surround gonocytes. In mice, lack of expression of Cx43 results in male infertility, demonstrating the important role for communication between Sertoli cells and germ cells in germ cell survival [88]. Defects in gonocyte development can also be brought about by defective migration of peritubular myoid cells into the developing gonad. This process is regulated in part by the growth factor neurotrophin 3 (NT-3) produced by Sertoli cells. The migrating cells express the highaffinity trkC receptor for NT-3, suggesting that NT-3 produced by Sertoli cells acts as a chemoattractant for migrating peritubular myoid cells [89]. Importantly, mice lacking trkC receptor show defective cord formation [89]. At 17.5 and 19.5 dpc, gonocyte numbers in trkC null mice are reduced by comparison with wildtype animals, demonstrating that proper migration of peritubular myoid cells into the gonad is required in part for correct cord formation and in turn for germ cell survival [89, 90]. Similarly, defective development of the interstitial Leydig cells can also disrupt cord formation, leading to infertility. In the developing testis, Dhh is expressed in Sertoli cells, whereas its receptor, Patched (Ptch-1), is expressed in fetal Leydig cells [78]. Dhh/Ptch-1 signaling is thought to trigger Leydig cell differentiation. Interestingly, male mice lacking the Dhh gene are infertile [78]. In the absence of Dhh, Leydig cells fail to differentiate, the basement membrane surrounding seminiferous cords may be absent, and many gonocytes are found outside of testis cords [91–93]. These studies demonstrate the profound influence of the somatic Sertoli cells and the other somatic cells of the testis on the germ cells even during embryogenesis. For germ cells, the consequence of being outside of the testis cord is severe. Loss of growth factor signaling likely activates a program of cell death within the

germ cell. Consistent with this idea the pro-apoptotic gene Bax is expressed in germ cells. The action of the Bax protein is thought to lead to apoptosis of germ cells that are ectopic to testis cords. Mice carrying two copies of a hypomorphic allele of the anti-apoptotic gene Bcl-x have reproductive defects and male mice are sterile [94]. PGCs migrate to the genital ridges but germ cells are depleted by 15.5 dpc. Deletion of both copies of the pro-apoptotic Bax gene in Bcl-x hypomorphic mice restored fertility to these animals, suggesting that the balance of pro-apoptotic and anti-apoptotic signals controls germ cell survival [94]. Thus, survival of germ cells in the developing embryo and fetus is controlled in a strict temporal and spatial pattern, presumably to ensure that germ cells do not survive outside of testis cords and develop inappropriately and to ensure that only high-quality germ cells survive to develop into gametes. Because germ cells can give rise to pluripotent stem cells, strict controls of their survival and differentiation are likely to exist. Interestingly, in humans 50% of pediatric germ cell tumors are found outside of the gonads and have been suggested to arise from PGCs that fail to die outside of the gonad. In contrast to germ cells, Sertoli cells seem to be able to differentiate normally in the virtual absence of germ cells, as demonstrated by many sterile mouse mutants (for a review, see [95]), although some abnormalities can be detected in Sertoli cell–only tubules in the adult testis. However, Sertoli cells are not totally independent of germ cell influence. For example, in ovotestes, where oocytes are growing inside testis tubules in hermaphrodites, some Sertoli cells appear to transdifferentiate into follicle cells [96, 97]. Several of the mouse knockout models affecting embryonic germ cell numbers do so at the time when germ cells have already colonized the embryonic gonad, at which time they are considered gonocytes.

Chapter 22 Gonocyte Development and Differentiation

Among the genes implicated in gonocyte development is the evolutionarily conserved vasa gene (Mvh in mammals) encoding a DEAD-box RNA helicase. In Drosophila, the prototypic vasa gene is required for germ cell development [98]. In vasa null mice, the numbers of PGCs in the developing gonad are normal up until 11.5 dpc. At 12.5 dpc a dramatic decrease is seen in gonocyte numbers, suggesting that Mvh may play some role in regulating gonocyte proliferation or survival [99]. Similarly, mutations in the Daz (deleted in azoospermia) RNA-binding protein genes are the most common molecularly defined cause of human infertility [100, 101]. As expected, mice lacking another member of the Daz family, Daz-like (Dazl). Similarly, mice lacking the zinc-finger transcription factor Zfp148 are infertile due to a reduced number of gonocytes starting at 13.5 dpc [102]. Although mutations in these genes results in germ cell loss in the developing gonad, it is not clear that the function of each of these genes per se is to effect germ cell survival. More likely they play a role in ongoing germ cell differentiation. Loss of function of these genes may result in germ cell death, perhaps because exquisite checkpoint mechanisms monitor ongoing germ cell development. When germ cell differentiation is perturbed, a program of cell death or apoptosis is activated. This mechanism would ensure that defective germ cells would not endure and would serve as a quality control mechanism to ensure the survival of only the highest quality germ cells, which would go on to make gametes. Formation of the gonocyte population in the developing gonad prefigures the formation of the spermatogonial stem cell population of the adult and, as such, represents a critical stage of gametogenesis. Although the stem cell population only becomes

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fully functional at puberty, the formation of stem cells from gonocytes takes place early in postnatal life.

V. FROM GONOCYTES TO SPERMATOGONIA As described earlier, mouse gonocytes resume mitosis after birth. At 1 day postpartum (dpp) in the mouse testis, their labeling index is 10.4 and 20.1% at 2 dpp, and 24.1% at 3 dpp [71], at which time the first A spermatogonia are identified (Fig. 22.6A). In the rat, resumption of gonocyte proliferation takes place around the third day of life and is strain dependent (Fig. 22.6B). In the U:WU Wistar rat, gonocyte proliferation rates are 0 during the first 3 days of life, 1.1% at 4 dpp, and 5.5 % at 5 dpp [72]. In a separate study, gonocyte proliferation rates were examined and similar numbers were observed: For the first 3 days of life, no gonocytes incorporated BrdU. At 4 and 5 dpp, 0.5 and 5% of gonocytes incorporated BrdU, respectively [103]. In the rat, A spermatogonia first appear at 6 dpp, and 19% of the spermatogonia incorporated BrdU at that time. At 7 dpp, 15% of the spermatogonia incorporated BrdU [103]. In parallel, in the Chinese hamster the onset of spermatogenesis occurs at 5 dpp [16], whereas in humans the first B spermatogonia are seen at 4 years of age [104]. At least two specific antibodies, Mab4B6.3E10 [105] and the activated leukocyte cell adhesion molecule ALCAM [106], can distinguish gonocytes from spermatogonia. The expression of ckit in spermatogonia and its absence in gonocytes can also be used for differentiating these two morphologically similar cell types (see later discussion). Recently, several genes differentially expressed in rat gonocytes

B

FIGURE 22.6 (A) Paraffin section of a mouse testis at 1 dpp. Sertoli cells are strongly labeled in brown for OSM, and unlabeled gonocytes (arrows) are very large in size and appear to be migrating toward the periphery of the seminiferous cords, where the basal membrane lies. Hematoxylin and anti-OSM/DAB stained. (B) Paraffin section of a rat testis at 3 dpp. Gonocytes (arrowheads) are very large and at this time are migrating towards the basal membrane. Stain: Mayer’s hematoxylin.

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and spermatogonia have been identified and include succinate dehydrogenase, ribosomal protein S15a, and a scaffolding subunit of protein phosphatase A [107], but the function of these genes remains to be elucidated. Nevertheless, such studies represent the first steps toward defining, at the molecular level, the differences between gonocytes and the PGCs from which they are derived and the spermatogonia which they produce. Many of the factors that affect gonocyte proliferation and differentiation remain to be elucidated. Until recently, it had not been possible to culture these cells in the absence of the accompanying Sertoli cells, so most studies have been performed using cocultures of gonocytes and Sertoli cells isolated from neonates (Fig. 22.7). When cultured on a fibroblast monolayer, postmigratory gonocytes do not survive well in vitro [108], reflecting the need for factors expressed by Sertoli cells at this time. Gonocytes are usually cultured together with the accompanying Sertoli cells ([103, 109–111]; for a detailed procedure, see [32]). In this system, gonocytes are identified with anti-vasa [112, 113] and anti-GCNA [114] antibodies (Fig. 22.8), because TNAP levels decrease during embryonic development, and SSEA-1 antigen is no longer expressed by the gonocytes. Recently, new methods have been developed for culturing gonocytes in the absence of Sertoli cells [13, 115]. In these cultures, isolated gonocytes are cultured on Matrigel with 20% FCS and growth factors such as LIF, bFGF, and forskolin. Using these systems, gonocytes have been reported to survive for up to 15 days; however, the possibility that gonocytes differentiate into spermatogonia under these conditions has not yet been ruled out [115]. When newborn rat gonocytes are cultured in serum-free medium over Matrigel, their labeling indexes are 0.2% on day 3, 5% on day 4, and 6.1% on

FIGURE 22.7 Phase-contrast micrograph of a coculture of gonocytes and Sertoli cells isolated from rat testis at 3 dpp. After 6 days in culture in the presence of 10 ng/mL CNTF, Sertoli cells form a compact monolayer and gonocytes (arrowheads) appear refringent, round, and on top of the Sertoli cell monolayer.

FIGURE 22.8 Histological section of a mouse testis at 4 dpp, showing gonocytes and spermatogonia labeled in black with antiGCNA. Note the differences in GCNA intensity between germ cells (arrows). Stain: Mayer’s hematoxylin and anti-GCNA/DAB/Cl2Ni.

day 5 of culture, and they can be cultured for up to 10 days [110, 111]. Surprisingly, the main hormones controlling spermatogenesis in vivo, follicle-stimulating hormone (FSH) and testosterone (T), do not affect newborn rat gonocytes when cocultured with Sertoli cells [108]. Other factors, such as 17-β-estradiol, which is produced locally by Sertoli cells [116] (the so-called “thymuline” from thymus extracts [117]), and activin A [118] increase the growth of cultured gonocytes. Alternatively, AMH, also known as Müllerian inhibitory substance (MIS), has been reported to induce mouse gonocyte differentiation in vitro [119]. In contrast to the relative dearth of information about the endocrine control of gonocyte fate, a wide variety of paracrine growth factors have been reported to affect gonocyte survival, proliferation, or differentiation in vitro. For example, bFGF [120] increases both survival and proliferation of newborn rat gonocytes in coculture with rat Sertoli cells. On the other hand, LIF [111] and OSM [103], both powerful PGC mitogens, stimulate survival of mouse gonocytes and both survival and proliferation of rat gonocytes [41]. CNTF increases gonocyte survival in vitro, but it does not affect their proliferation rate [111]. Because CNTF also increases Sertoli cell survival, the CNTF effect is probably indirect through somatic cells. Note that another member of the IL-6 family of cytokines, IL-6 itself, does not affect gonocyte survival in culture [121]. Most likely this reflects the specific type of ligandbinding receptor expressed by gonocytes because CNTF and IL-6 share a common signal-transducing component of their receptor, gp130. PDGF has also been shown to stimulate proliferation of newborn rat [116] and mouse [122] gonocytes in vitro. The effect of PDGF on gonocytes is in contrast to the effect of this

Chapter 22 Gonocyte Development and Differentiation

factor on PGCs that appear unresponsive. This indicates one important difference in growth factor dependence of these two cell types and also points to the importance of the negative data derived from PGC culture experiments. As stated earlier, another major difference between gonocytes and spermatogonia seems to be in the expression of the c-kit receptor and, therefore, in the dependence on KL. Gonocyte proliferation and differentiation are not regulated by KL, whereas spermatogonial survival and proliferation are. In vitro, KL has no effect on postnatal rat gonocytes [111], and injection of an anti-c-kit antibody into 2-day-old mice does not affect later spermatogenesis [123]. Interestingly, KL seems to play an important role later on in the control of the spermatogonial population and reflects the fact that c-kit levels are upregulated in type A spermatogonia [124]. These data highlight one of the differences between gonocytes and spermatogonia, because gonocytes are unresponsive to KL, whereas KL stimulates type A spermatogonial proliferation. These data indicate that one of the major changes to occur in the transition from a gonocyte to a spermatogonium occurs in growth factor dependence. As can be deduced, the differences in culture systems between migratory and postmigratory PGCs, gonocytes, and spermatogonia likely reflect differences in the growth factor requirements of these different cell types. Defining the factors that control the growth and differentiation of these different germ cell populations will ultimately be one of the key steps to a comprehensive understanding of mammalian gametogenesis. The identification of the factors regulating the growth of germ cells at any stage of development is important because defects in germ cell development at any stage of gametogenesis can result in infertility or in the development of testicular tumors. Because the spermatogonial stem cell population is derived from the gonocytes in the neonatal animal, understanding the growth and differentiation of this population of germ cells is important to our understanding of fertility in mammals. An increasing body of evidence derived from mouse knockout studies indeed points to the critical changes in gene expression that occur at the gonocyte stage of germline development and that are required for adult fertility. Advances in technology for analyzing gene expression in developing germ cells at all stages of development will help us decipher the pathways regulating mammalian gametogenesis. Equally important will be the development of techniques for manipulating germ cells and Sertoli cells to enable functional studies on genes identified as potential regulators of male germ cell development.

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VI. PGCS, GONOCYTES, AND THE ORIGIN OF TESTICULAR CANCER Failure of PGCs to stop proliferating in the fetal gonad is associated with the development of testicular tumors, termed teratomas and teratocarcinomas in mice. In this situation germ cells continue to proliferate and give rise to an embryonal carcinoma (EC) cell. EC cells are pluripotent stem cells and can give rise to derivatives of the three primary germ layers. If the EC cells differentiate into other cells, they form a benign teratoma usually comprised of many tissue types including skin, teeth, hair, muscle, and nerves. If the EC cells fail to differentiate, they form malignant teratocarcinomas comprised entirely of EC cells that can quickly kill the animal. Some of the genes that affect teratoma formation have been identified in mice and humans, but we still know very little about the controls on germ cell proliferation and differentiation in the developing gonad. In mice, the Teratoma (Ter) mutation is the most potent mutation to affect teratoma formation. Introduction of the Ter allele onto the 129/Sv strain background increases tumor incidence to 30% (reviewed in [125]). Several other genes have been shown to affect teratoma formation and include several tumor suppressor genes and the Sl gene. How mutations in a gene that is required for PGC survival would promote the incidence of PGC-derived tumors is difficult to understand. One possibility is that conditions that drive PGC death would select for cells that had acquired or lost oncogenes or tumor suppressor genes that would promote their survival. Although the action of some of the genes implicated in tumor formation are difficult to interpret, the action of the phosphatase and tensin homologue (PTEN) tumor suppressor gene provides some interesting insights into teratoma formation. The PTEN gene is an important molecule in signal transduction pathways in many cell types. Loss of PTEN in PGCs provokes increased proliferation and leads to testicular tumor formation in vivo, as reported by the higher incidence of testicular teratocarcinoma in germ cell-PTEN-null mice [126]. The transition from a mitotic PGC to a mitotically inactive gonocyte is also accompanied by dramatic changes in germ cell developmental potential, assayed by their ability to give rise to pluripotent stem cells. The developmental potential of germ cells has been assayed in vivo by the ability of transplanted germ cells to give rise to teratomas. When gonads are isolated from embryos and fetuses and transplanted to the kidney capsule, they give rise to teratomas that are

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indistinguishable from those that arise spontaneously in the testis [127, 128]. The cell type responsible for teratoma formation is the PGC, which gives rise to a pluripotent EC cell that in turn differentiates to give rise to the teratoma [127]. Notably, germ cells rapidly lose the ability to give rise to teratomas after 12.5 dpc in mice, suggesting that at this time they lose developmental potential. Similarly, in culture, germ cells isolated from 11.5-dpc mouse gonads can give rise to a different but related type of pluripotent stem cells termed an embryonic germ (EG) cell [35, 49]. Like EC cells, EG cells can give rise to cells derived from all three primary germ layers and in chimeric mice can also reenter the germline [49, 129, 130]. The ability of germ cells to form EG cells declines rapidly after 12.5 days of development and coincides nicely with the ability of PGCs to give rise to EC cells in vivo. Thus, the developmental potential of PGCs seems to decline rapidly after 12.5 dpc, indicating that important differentiation events are occurring at this time. Paralleling the loss of developmental potential are observations that differentiating gonocytes have differing abilities to give rise to spermatogonia. When fetal gonocytes from a 14.5-dpc fetus are transplanted into a recipient testis of an adult, they can participate in spermatogenesis [131]. However, when the same experiment is carried out with germ cells from a 12.5-dpc embryo, they are unable to do so. These data suggest that fetal germ cells become committed to differentiation into spermatogonia sometime between 12.5 and 14.5 dpc. At this time of development, germ cells must also undertake erasure and reestablishment of genomic imprints. Imprints mark genes as to their parent of origin and can have profound effects on gene expression from a paternally or maternally derived allele. In the germline this pattern of imprints must be erased at each generation to reimpose imprints appropriate to the sex-specific pattern of the mature gametes. In PGCs this process of imprint erasure is already initiated at 10.5 dpc and is complete by 15.5 dpc [132]. In gonocytes at 15.5 dpc there is evidence of increased methylation, indicating that imprints have begun to be reset by this time [132]. In culture, PGCs will differentiate into gonocytes and appear to execute a cell-intrinsic program of differentiation that might be independent of cell proliferation [24]. When PGCs are cultured in agents that promote DNA methylation or histone acetylation, their differentiation into gonocytes appears to be accelerated. These data suggest that genomic demethylation in PGCs might be part of the mechanism by which germ cell development is coordinated. Taken together these data suggest that during the transition from a PGC to a gonocyte, the germ cell has begun an ongoing program of differentiation at this time that alters

its developmental potential and commits it to the spermatogonial lineage. In humans it is accepted that an inappropriate germ cell number in the testis, brought about by increased temperature, cryptorchidism, or by other physical and genetic conditions, is linked to an increased rate of testicular cancer [133]. For example, cryptorchidism affects the development of both the testis and the associated male reproductive glands. The retention of the testis inside the body cavity after birth increases the testicular temperature, which in turn represses gonocyte proliferation and differentiation [134]. In humans, testicular germ cell tumors have been classified into three categories, based on age of occurrence [135]: (1) pediatric teratomas, teratocarcinomas, and yolk sac tumors, which occur before puberty; (2) seminomas and nonseminomas, which occur between 10 and 34 years of age; and (3) spermatocytic seminomas, which occur in men older than 50 years of age. This classification suggests that the first group derives from PGCs, the second from gonocytes, and the third from spermatocytes. The most common testicular cancer types in men are seminomas and nonseminomas that arise as a result of the occurrence of CIS cells (carcinoma in situ of the testis). CIS cells are thought to be derived from gonocytes and, in fact, they share similar morphological and immunohistochemical characteristics [136, 137]. Genetic studies in human populations have identified several chromosomal translocations associated with testicular tumors. Interestingly, one such frequently observed translocation, isochromosome 12p, contains some genes that could play important roles in germ cell and stem cell development [138]. These include the human homologues of the Stella and Nanog genes. Stella was identified in a screen for genes that regulate early germline development, although its function remains unclear at the present time [139]. Nanog was identified in a screen for genes that regulate the selfrenewal of embryonic stem cells in a LIF-independent manner and is also reported to be expressed in germ cells [140, 141]. The role of these genes in testicular tumor formation is unclear at present, but obviously either of these candidates could play a role in transforming PGCs into pluripotent stem cells. The importance of this research cannot be understated because testicular cancer remains the most frequent type of cancer in young men. As stated earlier, in mice, embryonal carcinoma cells are derived from testicular teratoma and teratocarcinoma that are thought to derive from PGCs [127, 142]. In fact, testicular germ cell tumors in the 129 inbred strain of mice are a model for pediatric testicular germ cell tumors in humans [143]. Mutations in the

Chapter 22 Gonocyte Development and Differentiation

Ter locus are characterized by a high incidence of congenital testicular teratomas only in the 129/Sv background [143]. In addition, spontaneous and engineered mutations in genes such as Trp53 and SlJ on the 129/Sv inbred background have been shown to increase the frequency of spontaneous testicular germ cell tumors [143, 144].

VII. ANALYZING AND MANIPULATING GENE EXPRESSION IN THE MAMMALIAN GERMLINE Advances in the technology for analyzing gene expression in different cell types have greatly improved the ability to comprehensively examine gene expression in cell types that are difficult to isolate in large numbers such as germ cells. A variety of techniques have been developed both for isolating cells in high purity and for analyzing gene expression from a very small number of cells. Expression of green fluorescent protein in specific cell types in transgenic animals allows those cells to be isolated using techniques such as fluorescent-activated cell sorting (FACS). Similarly, microdissection of specific cell types from tissue sections has been made possible by the technique of laser-assisted capture. Once isolated, a variety of techniques can be used to analyze gene expression in cells. Because of the ability to amplify cDNA in a manner that is representative of the original starting cDNA pool using PCR, it is now possible to generate large amounts of cDNA from a small population of cells. The analysis of genes expressed in cell types can be carried out by a variety of techniques such as serial amplification of gene ends (SAGE), suppression subtraction hybridization (SSH), and by means of microarrays. Such techniques are now being applied to germ cells including gonocytes. However, the utility of these techniques depends in large part on the ability to take expression data and move to studies of gene function. To dissect the pathways controlling the growth and differentiation of PGCs, gonocytes, and spermatogonia, researchers need to develop robust systems for manipulating gene expression in the various germ cell types. During the past few years, techniques for manipulating gene expression in various germ cell types and in the supporting somatic cell types have begun to be developed. In most of the examples of cell culture described earlier, several cell types were cultured together. This makes it difficult, and sometimes impossible, to determine if the effects of a specific factor on a specific cell type are direct or indirect.

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Methods for manipulating gene expression in germ cells have been developed, but this technology currently lags behind that available for manipulating somatic cells. Typically germ cells can be manipulated by many of the techniques used to manipulate somatic cells including electroporation, transfection, and viral infection. Electroporation has been used successfully to introduce genes into germ cells in situ in the testis. In this procedure, the DNA is introduced into the seminiferous tubule and the whole testis is then electroporated [145]. A drawback of this system is that DNA is introduced nonspecifically into many cell types. Transfection, using the reagents typically used to transfect somatic cells, can effectively transduce germ cells, but a large majority of cells are killed during the procedure [146]. Nevertheless, genes can successfully be expressed in germ cells following transfection. A variety of viral vector systems have been used to manipulate gene expression in germ cells, including adenoviruses, retroviruses, and lentiviruses. Adenoviruses can infect germ cells in vivo, but only for a short period of time; after 30 days, transgene expression is lost [147]. Whether adenovirus or adeno-associated viruses can infect germ cells in vitro is open to debate. Retroviral vectors have been used for many years and have been demonstrated to lead to long-term in vivo infection of several cell types. Tissue-specific targeting of genes and expression in mouse tissues have been demonstrated previously with the RCAS family of avian leukosis virus (ALV)-based retroviral vectors [148]. The use of this avian retroviral vector system in mammalian cells has several unique advantages. First, this system includes retroviral vectors that are replication competent in avian cells and grow to high titers (>107 IFU/mL). Second, ALVs are naturally replication defective in mammalian cells and, as a consequence, the vectors cannot spread to other cells. The defect(s) results in little or no viral protein expression, but does not affect experimental gene expression. Third, mammalian cells are not susceptible to efficient infection by ALVs because they lack the appropriate receptors. However, the cloning of the receptor for subgroup A ALVs, tvA, [149], made possible the generation of mammalian cell lines and transgenic mouse lines that express the tv-a receptor. PGCs, gonocytes, and spermatogonia isolated from mice expressing tvA ubiquitously from the β-actin promoter can be infected at high efficiency with ALV viruses [150, 151]. This system can therefore be used to address questions about signaling pathways in both PGCs, gonocytes, and spermatogonia. In summary, tissue-specific gene targeting can be achieved with the RCAS family of ALV retroviral vectors by expressing the ALV receptor under the control of a tissue-specific promoter, thereby

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targeting the expression of the experimental gene to the appropriate tissue. It should be possible to further restrict the expression of the exogenous gene by including an internal tissue-specific promoter in the vector. Germ cells can also be successfully infected with more traditional murine moloney leukemia virus– based vectors (MuLVs). Successful infection of germ cells with such viruses depends on several factors, including the viral coat, the method of delivery, and the cell cycle status of the infected cell [151]. For successful integration and expression using retroviral vectors, the infected cell must transit through the cell cycle. For certain cell types that do not divide (e.g., fetal gonocytes), this provides a barrier to using retroviral vectors for manipulating gene expression. Nevertheless, viral infection does not have the same drastic effect on germ cell viability as transfection and can be used to easily manipulate gene expression in vitro. Viruses can also be used to manipulate gene expression in germ cells in situ in the living testis [152, 153]. The use of lentiviral vectors that can successfully transduce noncycling cells can overcome these problems, and these vectors are likely to become more widely used to manipulate gene expression in germ cells. Because gonocytes do not divide for a large part of their existence, lentiviral vectors represent a powerful way in which to manipulate gene expression in these cells. Techniques are now also becoming available for manipulating the somatic cells of the gonad. For example, Shinohara et al. expressed kit ligand in Sl mutant mice using an adenovirus vector [154]. Although expression of kit ligand in Sertoli cells was unable to restore fertility to these animals, the resultant sperm produced could be used to produce normal offspring when injected into oocytes [154]. These data nicely demonstrate both a technique for experimentally manipulating Sertoli cells in vivo as well as a potential technique for the treatment of human infertility. Whereas viral vectors provide a tool for overexpressing genes in Sertoli cells, homologous recombination represents a technique for knocking out gene function. The development of technology for carrying out cell type–specific gene targeting using Cre-Lox mediated recombination provides a technique for examining loss of gene function in specific cell types. The success of this technology depends on the availability of promoter elements capable of driving expression of the Cre recombinase in a cell type–specific manner. To develop this technology for gene knockouts in Sertoli cells, a Sertoli cell expressed gene such as AMH is required. Mice producing the Cre recombinase under the control of the AMH promoter can be used to disrupt gene expression in Sertoli cells in the testis [155]. Using a LacZ reporter, the activity of this

promoter is reported to begin at 15.5 dpc in the developing male gonad, slightly later than expected from the AMH expression pattern [155]. Functional use of this system remains to be demonstrated, but this approach promises to be a powerful one. Thus, these two techniques, viral vector-mediated expression and Cre-mediated, Sertoli cell–specific recombination, can be used to up- and downregulate gene expression in Sertoli cells. Because male germ cells are so dependent on Sertoli cells for their survival, growth, and differentiation, these new technologies represent powerful approaches to dissecting the role of paracrine factors involved in germ cell development.

VIII. CONCLUSIONS The gonocyte represents a seemingly quiescent cell that lies at a developmental boundary between the proliferative and migratory PGCs and the spermatogonial stem cell. In large part it has remained unnoticed and understudied. Because gonocytes are the descendants of the PGCs but the antecedents of spermatogonial stem cells, they lie at a critical stage in germline development. Agents that affect germline development at this critical juncture could clearly have drastic effects on fertility—and, indeed, this is the case. Defective gonad development can result in both infertility and the development of germ cell tumors. For example, one of the most abundant man-made chemicals, di-(2-ethylhexyl)phthalate (DEHP), induces testicular damage in both developing and adult animals (for a review, see [156]). The active metabolite of DHEP, mono-(2-ethylhexyl)phthalate (MEHP), impairs division of neonatal Sertoli cells by acting at a post-cAMP site in the FSH-response pathway or via a mechanism independent of FSH. Relatively low levels of MEHP disrupt Sertoli cell–gonocyte physical interactions in vitro. This suppresses Sertoli cell proliferation in neonates via mechanisms that are specific to neonatal testes at a time when the very foundations of adult fertility are established [156]. The results also highlight the neonatal period of testicular development as one particularly sensitive to environmental chemicals. Moreover, recent progress in research on the etiology of CIS testicular cancer provides more evidence that the CIS cell is a gonocyte with stem cell potential. This could explain why an adult man can develop a nonseminoma, which itself is considered a neoplastic caricature of embryonic growth. The Sertoli and Leydig cells, which are activated postnatally before and after puberty, may play a critical role for both development of CIS from a gonocyte and subsequent neoplastic progression to form an invasive tumor.

Chapter 22 Gonocyte Development and Differentiation

During the past decade, there has been a reported increase in testicular cancer. This reported increase in testicular tumors is only one indication that male reproductive health is at risk. Some reports suggest that the incidences of undescended testes and hypospadias are on the rise. Other studies suggest that semen quality has deteriorated, at least in some countries. Some epidemiological studies indicate that environmental factors may play a causative role in these observed problems in male reproductive health. During the next few years, genome-wide expression analyses are likely to result in a vastly improved understanding of the genetic programs that must be executed to ensure correct germline development. That understanding will in turn allow a comprehensive analysis of how germline development can be disrupted in the embryo, the fetus, postnatally up to puberty, and in the adult. This knowledge may help us understand what agents place the germline at risk and how to protect the germline during the early stages of development.

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C H A P T E R

23 Hormones and Spermatogonial Development MARVIN L. MEISTRICH, GUNAPALA SHETTY, OLGA U. BOLDEN-TILLER, AND KAREN L. PORTER The University of Texas, M. D. Anderson Cancer Center, Houston, Texas

I. INTRODUCTION II. HORMONE EFFECTS ON NORMAL SPERMATOGENESIS III. LOCALIZATION OF HORMONE RECEPTORS IV. CONTRARY EFFECTS OF HORMONES ON SPERMATOGONIAL DEVELOPMENT V. CELLULAR AND MOLECULAR TARGETS FOR INHIBITION OF SPERMATOGONIAL DEVELOPMENT BY TESTOSTERONE VI. FUTURE DIRECTIONS References

II. HORMONE EFFECTS ON NORMAL SPERMATOGENESIS Two hormones, primarily T but also folliclestimulating hormone (FSH), are required for the complete process of spermatogenesis in normal animals. The two act additively to support survival and differentiation of germ cells [1, 2]. In particular, the progression of round spermatids through elongation and condensation is strongly dependent on T in all mammals. However, the point at which the cells are lost and the relative importance of T and FSH are species dependent [3]. Deprivation of T and FSH in rats and mice reduces the numbers of spermatocytes and round spermatids to levels dependent on the residual concentrations of these hormones; nevertheless, some cells proceed to the round spermatid stage [1, 4]. The effects of T and FSH deprivation on spermatogonial numbers in rodents are moderate; at most a 50–70% decline in type A or B spermatogonia or early spermatocytes when these hormones are absent [5]. Some reports show that T alone can largely restore the numbers of spermatogonia [1, 5], but others indicate that only FSH can restore the numbers [6]. In contrast, in adult primates (both human and monkey), the deprivation of these two hormones, particularly FSH [7, 8], primarily affects the conversion of type A pale to B spermatogonia, resulting in 90% reductions in B spermatogonial numbers, but there are only minor losses in the development of these B spermatogonia to spermatocytes and spermatids [9, 10].

I. INTRODUCTION In this chapter, we focus on recent findings that hormones, primarily testosterone (T), strongly inhibit spermatogonial differentiation in rodents in a variety of pathological situations. We first summarize the effects of hormones on normal spermatogenesis, which are subtle at the level of spermatogonia, and the localization of the hormone receptors in the testis. We later discuss proposed models that involve paracrine or juxtacrine effects from receptor-positive somatic cells to produce major inhibitory effects on spermatogonial development. The most likely candidate for the cell involved in hormone-regulated interaction with the spermatogonia is the Sertoli cell, but other testicular somatic cells are also considered.

SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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Marvin L. Meistrich, Gunapala Shetty, Olga U. Bolden-Tiller, and Karen L. Porter

In contrast, exogenous estrogens, such as estradiol, usually inhibit normal spermatogenesis by suppressing gonadotropins [11] or T biosynthesis [12], or possibly through a gonadotropin- and T-independent effect on spermatogenesis [13]. Results from knockout mice, however, do not indicate direct effects of physiological levels of estrogens on spermatogenesis. The lack of a direct requirement of the germ cells for estrogen is supported by the production of fertile spermatozoa by germ cells from estrogen receptor α (ERα) null mice transplanted into testes of wild-type mice [14]. There are, however, indirect effects on the spermatogenesis in the ERα knockout mice resulting from dilation of the seminiferous tubules [15] in the absence of the estrogen-dependent resorption of tubular fluid in the efferent ductules [16]. The concept that the requirement for estrogens in spermatogenesis is only indirect is supported by the observation of only minor spermatogenic lesions in young adult aromatase-null mice, which do not develop in severity until the mice age [17]. In contrast to the lack of a direct requirement for estradiol for normal spermatogenesis and the inhibition by pharmacological doses, several reports indicated positive actions of estradiol on this process. In vivo, estradiol stimulated spermatogenesis in hypogonadal (hpg) mice [18], and in vitro stimulated proliferation of neonatal rat gonocytes [19] and inhibited apoptosis in human testis [20].

III. LOCALIZATION OF HORMONE RECEPTORS There is general agreement that the androgen receptor (AR), FSH receptor, and ERα are localized in the somatic cells of the testis. Of these, the FSH receptor appears to be specific for Sertoli cells. Therefore, hormones most likely affect spermatogenesis through paracrine and juxtacrine interactions with the somatic cells. The Sertoli cell, because it is in physical contact with the germ cells and because FSH receptor is localized to this cell, is the prime candidate for mediating those interactions, but it is possible that other somatic cells are the target for steroid hormone effects.

A. Androgen Receptor It is generally accepted that germ cells in rodents lack AR [21, 22], although there is one report that late-stage germ cells do contain this receptor [23]. Furthermore, it has been shown by spermatogonial transplantation that when germ cells contain a mutant, nonfunctional androgen receptor gene and the other cells of the testis

have the normal gene, germ cell development proceeds to completion [24]. There is also general agreement that in the normal adult testis the AR is found in the Leydig cells, peritubular myoid cells, vascular smooth muscle cells, and the Sertoli cells. In the Sertoli cells of normal rodents, the levels of AR are highly dependent on the stage of the cycle of the seminiferous epithelium, but after differentiating spermatogenic cells are eliminated, for example by irradiation [25], AR levels are equal in Sertoli cells in all tubules. However, because the stem and early differentiating type A spermatogonia constitute a small percentage of cells in normal animals their AR status has not been clearly described. To describe unusual effects of T on spermatogonial differentiation in various pathological situations, we must first demonstrate the localization of the AR protein in these cases. This was determined by immunohistochemistry and is shown in Figure 23.1A for testes from irradiated rats and in Figure 23.1B for jsd mutant mice. In both species, Sertoli cells within the testes showed a uniform level of staining, Leydig cells displayed greater levels of AR staining, and peritubular myoid cells and some vascular cells stained most intensely for AR. Germ cells, including type A spermatogonia (Figs. 23.1A and B) and the type B spermatogonia and spermatocytes observed in jsd testes (not shown), did not display any AR staining. Some non-Leydig interstitial cells were also negative for AR staining. Because the germ cells lack AR and because the mechanism by which androgens suppress spermatogenesis in irradiated rats and jsd mice is AR dependent, the suppression cannot be the result of androgens acting directly on the germ cells. Instead, this phenomenon appears to result from an indirect mechanism, likely involving Sertoli cells, but possibly involving the peritubular myoid, Leydig, or the vascular smooth muscle cells, all of which possess AR (Fig. 23.1).

B. FSH Receptor It is generally accepted that in normal animals the FSH receptor is specifically localized on the Sertoli cells [26], despite one report of FSH binding to spermatogonia [27]. Thus any effects of FSH on spermatogenesis may be considered to involve juxtacrine and possibly paracrine effects of the Sertoli cell on spermatogonia.

C. Estrogen Receptors In the adult, ERα is localized in the Leydig cells in the mouse [28], the rat [29], and humans [30]. The absence of ERα in germ cells is consistent with the absence of any effect of knocking out ERα on the ability of germ

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IV. CONTRARY EFFECTS OF HORMONES ON SPERMATOGONIAL DEVELOPMENT A. Proliferation-Apoptosis (PAp) Block in Spermatogonial Development

FIGURE 23.1 Immunohistochemical analysis of androgen receptor in testes from irradiated rats and jsd mice (see color plate). (A) Irradiated LBNF1 rat, 15 weeks after 6-Gy irradiation. (B) Jsd mutant mouse, 12 weeks old. Stained cells were marked as follows: SC, Sertoli cells; L, Leydig cells; P, peritubular myoid cells; V, vascular smooth muscle. Unstained cells were marked: A, type A spermatogonia, mI, miscellaneous interstitial cells. Bar = 10 μm.

cells to undergo complete differentiation when transplanted into the testes of normal mice [14]. ERβ expression is more widespread in the testis, but its cellular localization appears to be species dependent and there are discrepancies between reports. Sertoli cells have been reported to express ERβ in all species tested, including rat [31], mouse [22], and human [32]. Most germ cells, including spermatogonia, in these species, also appear to express ERβ [22, 32–34], although two studies did not find ERβ in these cells in the adult mouse [28, 35]. Leydig cells, have been reported to possess ERβ in mouse [22, 35], rat [34], and human [32], but one group did not find ERβ in rat Leydig cells [33]. In any case, there is no evidence that estrogen affects spermatogenesis through ERβ since knocking out the ERβ gene had no effects on spermatogenesis [36].

The stem spermatogonia, designated As, maintain their numbers by self-renewal, and some differentiate to form by sequential divisions Apr (A paired), Aal-4 (A-aligned), and Aal-8 spermatogonia, which go on to produce A1 spermatogonia. This differentiation may be blocked in various ways. One way, which seems to be T dependent, has been defined as the proliferationapoptosis (PAp) block [37]. The early type A spermatogonia proliferate, but their numbers remain relatively constant because of apoptosis [38, 39]. A variety of pathological conditions produces similar testicular histology consistent with a PAp block. In the rat, these include testicular toxicants, such as hexanedione [40], boric acid [41], radiation [42], procarbazine [43], dibromochloropropane [44], and indenopyridines [45]. The type A spermatogonia proliferate in atrophic tubules but do not accumulate, because they continue to be lost by apoptosis many months after the original acute or subchronic exposure. Similarly, atrophic tubules with actively dividing stem type A spermatogonia were also observed in testis cross sections from untreated 27-month-old Brown-Norway rats [46]. In mice, a PAp block in spermatogonial differentiation can be induced by a mutation at the jsd (juvenile spermatogonial depletion) locus [47]. There is an initial wave of spermatogenesis in these mice during the pubertal period, but this is not maintained, so that the adult testis tubules contain only Sertoli cells and type A spermatogonia. As in the toxicant-treated rat models, these spermatogonia proliferate but die by apoptosis instead of differentiating [48]. In all of these cases, the blocks appear to be irreversible if the animals are left untreated. In particular, the blocks in spermatogonial differentiation that occur in jsd mice [49] or after exposure of rats to hexanedione [40] or radiation [42] have persisted during observation periods that exceeded a year.

B. Reversal of Blocks in Spermatogonial Development by Suppression of Testosterone In many of the cases just mentioned, the block in spermatogonial development could be reversed by suppression of T, usually with gonadotropin-releasing

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hormone (GnRH) analogues. The recovery of spermatogenesis in rats can be stimulated with hormone treatment given after irradiation, reversing this PAp block to spermatogonial development [50]. Either GnRH analogues or exogenous T, both of which lower the intratesticular testosterone (ITT) levels, were able to restore spermatogonial development. However, because these treatments suppress ITT, which is required for spermatid differentiation, histological recovery proceeds only to the round spermatid stage. But after stopping suppression of T, spermatids can differentiate into sperm, and because spermatogonial differentiation continued for at least 10 weeks after restoration of ITT in the irradiated rat model, the rats were fertile [51]. In other cases involving PAp blocks to spermatogonial differentiation in the rat, such as hexanedione, procarbazine, and DBCP treatment, maintenance or recovery of spermatogenesis was also enhanced by administering GnRH analogues after the toxicant [52–54]. Although prevention of the indenopyridineinduced block to spermatogonial differentiation was achieved when GnRH analogues were given either before or before and after drug treatment, there was no effect on the maintenance of spermatogenesis when the GnRH analogue was given only after indenopyridine administration [45] (S. A. Hild, personal communication). However, GnRH-analogue treatment did maintain or stimulate recovery of spermatogenesis when given after exposure to two other toxicants, busulfan [55] and heat [56], although the blocks in spermatogonial differentiation were not well characterized. Furthermore, in jsd mice, treatment with GnRH analogues also reversed the PAp block and restored spermatogonial differentiation up to the spermatocyte and round spermatid stages [57, 58]. However, in contrast to the observations in irradiated rats, the increase in spermatogonial and spermatocyte differentiation produced by GnRH-analogue treatment in jsd mice was transient. However, one wave of late spermatids was produced from the differentiating spermatogonia and spermatocytes that developed during the GnRHanalogue treatment. Offspring could be produced, but only by testicular sperm extraction and intracytoplasmic sperm injection [59]. Suppression of ITT with either GnRH-analogue or exogenous testosterone treatment also enhanced the production of differentiated germ cells from stem cells transplanted into either rat or mouse recipients in which spermatogenesis was depleted by busulfan treatment [60–62]. In these studies, however, the hormonal treatment was given both before and after the transplantation so that it was not possible to determine whether the suppression of T enhanced the ability of the transplanted stem cells to initially survive and enter an appropriate niche in the tubules or their subsequent

ability to survive, proliferate, and differentiate. One study indicated that only treatment with a GnRH analogue before, but not after, transplantation was effective in increasing the number and size of colonies [63] supporting the role of T-suppression in the transplantation step but not in the subsequent development. However, in another study, a GnRH analogue given after transplantation stimulated the proliferation and/or survival of the early-stage type A spermatogonia after transplantation, but differentiation could not be assayed in that system [64]. 1. Evidence That Testosterone Is a Major Inhibitory Factor Although treatment of animals with GnRH analogues or T has other effects in addition to the suppression of ITT levels, several lines of evidence indicate that T, acting through the AR, is the major factor that inhibits spermatogonial differentiation in irradiated rats and jsd mice. The most direct evidence was provided by the inhibition of GnRH-analogue or low-dose T-stimulated recovery of spermatogenesis in irradiated rats or jsd mice in a dosedependent manner by exogenous T [58, 65]. Second, the extent of recovery of spermatogenesis in the irradiated rats and in the jsd mice stimulated by different combination treatments of the above hormones showed an excellent negative correlation with ITT levels present during the treatment period [58, 65, 66]. Third, the decline in spermatogenesis in jsd mice coincided with the gradual increase in T levels during the pubertal period. Finally, the reversal of exogenous T-induced inhibition of spermatogonial differentiation by an AR antagonist in GnRH analogue–treated irradiated rats and jsd mice confirmed that T inhibited spermatogonial differentiation in these model systems by acting through the AR. This AR-dependent inhibition was not restricted to T alone; all three androgens tested (dihydrotestosterone, 7α-methyl-19-nortestosterone, and methyltrienolone) inhibited spermatogonial development in irradiated rats [67], suggesting that AR activation by androgens determines the inhibition of spermatogonial differentiation. However, because T is the major testicular androgen present in vivo under normal circumstances and in irradiated rats [67], it is the primary androgen that inhibits spermatogonial differentiation in these models. Furthermore, because these compounds represent 5αreduced, non–5α-reducible, and nonmetabolizable androgens, their inhibitory action cannot be dependent on metabolizing to a different common chemical form. 2. Weak Inhibition by FSH Although, as discussed earlier, T is an inhibitory factor, it is possible that FSH, which is also elevated in

Chapter 23 Hormones and Spermatogonial Development

these pathological models of testicular atrophy and is suppressed by the GnRH-analogue treatment, may also be inhibiting spermatogonial differentiation. As was the case with T, a relatively good inverse correlation was observed between FSH levels in irradiated rats given different hormone treatments and the ability of spermatogonia to differentiate [37]. However, the correlation with FSH could be coincidental because there is a concomitant rise in FSH whenever T is given to rats in which gonadotropin secretion has been suppressed by GnRH analogues [68]. To test for a role of FSH, exogenous FSH was given to jsd mice or irradiated rats while suppressing endogenous FSH and androgen levels with GnRH analogues. Although the FSH did not inhibit spermatogonial differentiation in jsd mice [66], it inhibited the differentiation of spermatogonia in irradiated rats (G. Shetty, unpublished data). However, FSH did not inhibit spermatogonial differentiation as drastically as did exogenous T and, in another experiment, reduction of active FSH levels with an FSH antibody [69] did not enhance differentiation in irradiated rats. Thus, we conclude that FSH can weakly inhibit spermatogonial differentiation in at least one of these pathological models. But in at least one other model, the inhibition is completely due to T. 3. Stimulation by Estradiol We had previously shown that administration of estradiol to GnRH analogue–treated irradiated rats maintained or slightly increased spermatogonial differentiation [67]. We have also obtained data indicating that estradiol alone can strongly stimulate spermatogonial differentiation in both irradiated rats and jsd mice (G. Shetty, unpublished observations). Although these results can be explained in a large part by the potency of estradiol at reducing T levels, some of the data suggest that estradiol may directly stimulate an effect on spermatogonial differentiation in irradiated rats (G. Shuttlesworth, unpublished observations).

V. CELLULAR AND MOLECULAR TARGETS FOR INHIBITION OF SPERMATOGONIAL DEVELOPMENT BY TESTOSTERONE A. Cellular Target for Testosterone Action The four major testicular cell types with welldocumented AR content—the Sertoli, Leydig, peritubular myoid, and vascular smooth muscle cells— must all be considered targets for T-induced inhibition of spermatogonial differentiation. Whereas the Sertoli cell is

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the most obvious target, because it is the only cell in direct contact with the spermatogonia and because several well-documented Sertoli cell juxtacrine factors—glial cell line-derived neurotrophic factor (GDNF), stem cell factor (SCF), and Fas ligand (FasL)—act on the spermatogonia, the other cells cannot be excluded. The possible roles of the other cell types in the T-dependent inhibition of spermatogonial differentiation are discussed briefly in the following subsections. 1. Peritubular Myoid Cells Peritubular myoid cells are separated from the spermatogonia and Sertoli cells only by extracellular matrix forming a layer of the basement membrane. Sertoli cell–peritubular cell interactions were reviewed in detail in Chapter 18 on Sertoli cell–somatic cell interactions. Because the peritubular cells have AR and secrete both extracellular matrix components and growth factors, they can be responsible for the effects of T on spermatogonial differentiation. These peritubular myoid cell products can act either on the spermatogonia directly or on the Sertoli cells to produce secondary factors regulating spermatogonial differentiation. Androgen-modulated factors produced by peritubular cells do affect Sertoli cell function, and this activity has been ascribed to a factor called PModS [70]. In addition, retinol is known to regulate the differentiation of type A spermatogonia [71], and a role for the peritubular myoid cell in the transport of retinol to the seminiferous tubules has been proposed [72], but there is no information as to whether this is affected by androgen levels. 2. Leydig Cells Owing to their production of T, Leydig cells clearly have a role in the T-dependent inhibition of spermatogonial differentiation. However, the question is whether the Leydig cell produces other paracrine factors that respond to T in an AR-dependent autocrine manner and regulate spermatogonial differentiation. Because the regulation may occur through Sertoli cells, the Leydig cell–Sertoli cell interactions, also reviewed in Chapter 18, should be considered. Currently there are no other Leydig cell produced factors, besides T, that are both inhibited by GnRH analogues and could act on the tubules to block the differentiation of spermatogonia. Despite the known interdependence of Sertoli and Leydig cells, very little is known about the specific factors involved. Recently macrophage migration inhibitory factor has been demonstrated to be produced by Leydig cells and to regulate Sertoli cells, but, because it is still produced

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after hypophysectomy [73], it is not a candidate for the T-dependent regulation of spermatogonial differentiation in the models discussed here. Indirect evidence indicates that a Leydig cell paracrine factor is involved in the support of spermatogonial differentiation. Although GnRH-analogue treatment can restore spermatogenesis after hexanedione-induced testicular atrophy, it is unable to do so if the rats are first treated with ethane dimethane sulfonate, which eliminates the Leydig cells [74]. Hence, it appears that a Leydig cell factor, in addition to suppression of T, is required for the restoration of spermatogonial differentiation after hexanedione insult. But these studies do not address the question of whether T regulates a paracrine factor from Leydig cells that would inhibit spermatogonial differentiation. 3. Vascular Smooth Muscle Cell Intratesticular interstitial fluid has been shown to increase following adverse treatments that either induce a PAp block in spermatogonial differentiation or produce testicular atrophy that is reversible by suppression of T. These treatments include exposure to radiation [75], busulfan [55], procarbazine [76], DBCP [77], or hexanedione (K. Boekelheide, personal communication), as well as raising testicular temperature by local heating [78] or by cryptorchidism [79]. This increase in interstitial fluid may have a causative role in the block inasmuch as quantitative data show that the amount of interstitial testicular fluid is extremely well correlated with the inhibition of spermatogonial differentiation (K. L. Porter, unpublished observations). How these toxicants cause the increase in interstitial fluid is not known. These toxicants can act directly to change blood flow, vascular permeability, and vasomotion, all of which contribute to transvascular exchange of fluids [80]. Alternatively, because treatment with LH (or hCG) generally increases interstitial fluid volume by acting on Leydig cells [81] and T can act directly on the vasculature to maintain or increase interstitial fluid volumes [82, 83], the high ITT and LH levels in these models may also be responsible for this edema. We have shown, in irradiated rats treated with combinations of GnRH analogues and steroid hormones, that interstitial fluid volumes are regulated by ITT concentrations (K. L. Porter, unpublished observations). Hence, it is likely that the vascular cells, specifically the vascular smooth muscle cells that express AR, are regulating testicular edema. Testicular edema could in turn block spermatogonial differentiation by some component of the edematous fluid or by causing intratesticular hypoxia, which has been shown to induce germ cell-specific apoptosis in rats [84].

B. Models for Testosterone-Induced Block in Spermatogonial Differentiation Because germ cells lack AR, T must act via paracrine or juxtacrine routes between the cells that contain the receptors for these hormones and the spermatogonia. However, a model is needed to explain why T inhibits spermatogonial differentiation only in the variety of pathological situations described earlier but not in normal rodents. The model chosen to explain this apparent contradiction depends on whether the toxicant or genetic defect directly alters the spermatogonia or the androgenresponsive somatic cell involved. In most cases the target is not known. Although certain toxicants are believed to act primarily on Sertoli cells (e.g., hexanedione) [85] or germ cells (e.g., radiation) [86], it is not possible to prove that the long-term effects are due to action on these cells. Only for the jsd mutation have transplantation experiments shown conclusively that the defect is in the spermatogonia, not the somatic cells [87, 88]. Hence, Figure 23.2, which lays out our model, is divided into two parts representing different cellular targets: Figures 23.2A and B assume the defect lies in the spermatogonia, whereas Figures 23.2C and D assume it lies in the Sertoli cells. The Sertoli cell was used as the example of the androgen-responsive somatic cell because it is the most likely one to exert paracrine effects on germ cells and the only one that can act by juxtacrine mechanisms. In some cases, however, we cannot rule out that the peritubular, Leydig, or vascular smooth muscle cells are involved instead or in addition. The model is further subdivided according to the primary cause of the block. If the primary cause is the lack of a functional signal for the spermatogonia to differentiate (Figs. 23.2A and C), then the observed apoptosis could be secondary to cells remaining undifferentiated for too long. Alternatively, if the primary cause of the block is the induction of apoptosis (Figs. 23.2B and D), then the failure to observe differentiation is secondary to the failure of the cells to survive to an appropriate stage. Thus there are four possibilities. The first two involve alterations in spermatogonia, which could make these cells either more dependent on T-suppressible growth and differentiation factors from the Sertoli cells (Fig. 23.2A) or more sensitive to T-induced proapoptotic factors from the Sertoli cell (Fig. 23.2B). The second two possibilities involve alterations in the Sertoli cells, which could allow T to inhibit expression of normally secreted growth factors (Fig. 23.2C) or to induce expression of pro-apoptotic factors (Fig. 23.2D). Although these models are not meant to represent all possibilities, they do show one feature in common that should stimulate further research in this area. All of these models predict that,

A. Pathological Defect Induced in Spermatogonia; Cause Is Failure of Differentiation

B. Pathological Defect Induced in Spermatogonia; Cause Is Apoptosis

T-inhibited growth factor

Situation

Constitutive growth factor

T-dependent apoptotic factor

Normal T present

Differentiation

Normal No T

Differentiation Defect Development of apoptotic pathway

Defect Loss alternative of growth-factor receptor

Pathological T present

Apoptosis

Pathological No T

Differentiation

Sertoli cell

Normal spermatogonium

C. Pathological Defect Induced in Sertoli Cell; Cause Is Failure of Differentiation Situation

Growth normally constitutive;Factor, T-inhibited in pathological situation

Altered spermatogonium

D. Pathological Defect Induced in Sertoli Cell; Cause Is Apoptosis Apoptotic factor, normally repressed; T-dependent in pathological situation

Normal T present

Normal No T Pathological T present

Outcome

Outcome

Differentiation

Differentiation Defect Growth becomes factor inhibited by T

Defect Apoptotic factor becomes stimulated by T

Apoptosis

Pathological No T

Differentiation

Normal Sertoli cell

Altered Sertoli cell

Spermatogonium

FIGURE 23.2 Models to explain T-dependent inhibition of spermatogonial differentiation in pathological situations in mice and rats but not in normal rodents. Two factors, the cell type in which the defect is induced (spermatogonia in parts A and B; Sertoli cells in parts C and D) and the primary cause of the spermatogonial block (failure to differentiate in parts A and C; apoptosis in parts B and D) are considered, resulting in four possible combinations. (A) Assume a defect is induced in spermatogonia so that they lose one receptor or intracellular component of the signal transduction pathway (square symbol) for a conditionally redundant growth, survival, or differentiation factor pathway. Normal spermatogonia have an alternative pathway to support this step (triangle), but the ligand of this alternative pathway is suppressed by T. (B) Assume a defect is induced in spermatogonia so that they express a receptor or intracellular pathway that makes them sensitive to a T-dependent apoptotic factor (circle) produced by Sertoli cells. (C) Assume a defect is induced in Sertoli cells, resulting in the development of inhibition by T of an essential growth or differentiation factor for spermatogonia. (D) Assume a defect is induced in Sertoli cells resulting in the T-dependent production of an effector for spermatogonial apoptosis.

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in the pathological situation, there should be at least one gene product specifically regulated by T in the target somatic cell and that its regulation will either positively or negatively correlate with the degree of spermatogonial differentiation.

C. Possible Involvement of Juxtacrine/Paracrine Sertoli–Spermatogonia Interactions Because the Sertoli cell is the most likely candidate for mediating hormonal regulation of spermatogonial differentiation, we now summarize the known interactions between the two cell types and discuss their potential relevance for involvement in the T-dependent PAp block of spermatogonial differentiation in the various pathological situations. 1. Stem Cell Factor–c-kit The tyrosine-kinase receptor, c-kit, and its ligand, SCF, are expressed in differentiating spermatogonia [89] and Sertoli cells [90], respectively, and are usually required for spermatogonial differentiation [91, 92]. Mice with mutations in c-kit (W-locus) or SCF (Sl-locus) contain some spermatogonia but they fail to differentiate due to a PAp block as described earlier [48]. This suggests that the SCF–c-kit signaling system might be involved in the PAp block of spermatogonial differentiation in other conditions and its hormonal dependence. Support for this hypothesis was obtained from experiments in which hexanedione-treated rats were shown to primarily express mRNA for the soluble form of SCF, which is less effective at stimulating spermatogonial development than the alternate-spliced mRNA for the membrane-bound form [93], preferentially produced in control rats [94]. Delivery of exogenous soluble SCF to hexanedione-treated rat testes did stimulate the survival and/or proliferation of the type A spermatogonia [94]. Furthermore, treatment of rats with hexanedione-induced testicular atrophy with a GnRH analogue, which induced spermatogonial differentiation, also increased the proportion of SCF in the membrane-bound form [52]. However, in contrast to the results with hexanedione, the ratio of mRNA for the soluble and membranebound forms of SCF was not altered when a PAp block in spermatogonial differentiation was induced by radiation (J. Lee and K. Boekelheide, personal communication). Furthermore, a GnRH analogue given after transplantation stimulated the proliferation and/or survival of the early-stage (c-kit negative) type A spermatogonia transplanted into testes of Sl/Sld mutant

mice, which lack the membrane-bound form of SCF [64]. However, this latter observation does not exclude the possibility that suppression of T can also act on later stage spermatogonia through their c-kit receptor. But the data on hormonal regulation of SCF do not support a role for the SCF–c-kit system in the hormone-dependent block in spermatogonial differentiation. FSH increases the levels of SCF mRNA [95], but, as shown earlier, FSH does not stimulate and can inhibit spermatogonial differentiation in the models with a hormonedependent PAp block. Also T, which is major inhibitor of spermatogonial differentiation in these models, had no effect on levels of SCF mRNA [96]. 2. GDNF–(GFRα-1,Ret) GDNF is produced by Sertoli cells, and spermatogonia express the receptor-signaling complex for GDNF, which includes the Ret receptor tyrosine kinase and GFRα-1 (GDNF family receptor α1) [97]. GDNF positively regulates spermatogonial proliferation but negatively regulates their differentiation, as GDNF+/− mice show a progressive loss of stem cells with age and mice overexpressing GDNF in their spermatogonia show an accumulation of spermatogonia that fail to differentiate [97, 98]. The accumulation of spermatogonia indicates that this block in spermatogonial differentiation is very different from the PAp block. Hence, alterations of GDNF levels produce effects on the spermatogonial differentiation pattern that are different from those consistently observed with toxicants or other mutations such as those in the genes for SCF or c-kit. Furthermore, as was the case with SCF, FSH increases GDNF expression but T has no effect on its expression [98]. Thus alteration in the GNDF–(GFRα-1,Ret) signaling system cannot be a primary cause of the PAp block in spermatogonial development and its reversal by suppression of T. 3. Notch Pathway Data, primarily from Caenorhabditis elegans, have indicated that the Notch receptors and their ligands, Jagged and Delta, are responsible for the balance between mitotic proliferation of spermatogonia and entry into meiosis [99]. Although Notch is expressed in Type A spermatogonia and Jagged and Delta are expressed in Sertoli cells and spermatogonia [100], no data on their functional importance for spermatogonial differentiation is available in mammals. The one study of antibodies to Notch and Jagged with rat testis in tissue culture only reported effects on spermatid development [101]. Thus, there is insufficient information to

Chapter 23 Hormones and Spermatogonial Development

speculate as to whether the Notch pathway is involved in the PAp block in spermatogonial development and its reversal by suppression of T. 4. Fas-Fas Ligand The system consisting of the Fas (APO-1, CD95) transmembrane receptor protein and its ligand FasL has been proposed as a key regulator of germ cell apoptosis in the testis [102]. Although expression of FasL and Fas receptor have been reported to be localized in the Sertoli cells and germ cells, respectively [102], some disagreement about its localization remains [103]. Nevertheless, both Fas receptor and FasL can be upregulated by toxicants that result in germ cell apoptosis [86, 102]. Furthermore, a role for the Fas-FasL system in regulating spermatogonial survival to allow differentiation when a PAp block occurs has been demonstrated by the differentiation of spermatogonia when the Fas receptor gene was eliminated in KitW-v/W-v mutant mice [104]. Regarding the regulation of the Fas-FasL system by T, there are conflicting results; one study showed that suppression of T produced a large reduction in FasL and a small reduction in Fas receptor levels [103], whereas another study showed increases in Fas receptor after suppression of T [105]. Note, however, that the decline in FasL in the first study [103] was in differentiating germ cells. It is possible that the Fas-FasL ligand system may be involved in the PAp block in spermatogonial development and its reversal by suppression of T, but further research is needed to resolve the inconsistencies reported.

IV. FUTURE DIRECTIONS Several areas of investigation appear to be important for resolving questions of how T acts to inhibit spermatogonial differentiation. First, it is important to determine whether or not the Sertoli cell is the primary target for these hormone effects. One approach could be to selectively eliminate the AR in Sertoli or other androgen-responsive somatic cells of the testis using cell type–specific expression of Cre-recombinase in mice with conditional (loxP flanked) null mutation in the AR gene [106]. Second, it would be important to identify the androgen-regulated genes in the target cells that are modulated during suppression of T and the concomitant restoration of spermatogonial differentiation. This can be done with oligonucleotide microarray analysis as was recently done to identify FSH-regulated genes in Sertoli cells [107]. Although it is likely that many genes will be identified, the possibilities can be narrowed down by correlating the regulation of

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these genes with the time courses of reversal of the PAp block in spermatogonial differentiation after different toxicant and hormonal treatments. Finally, after information is obtained from the first two investigations, the pathway in both the hormone-responsive target cell and the spermatogonia by which T either causes apoptosis or failure of differentiation needs to be elucidated.

Acknowledgments We thank Dr. Gail Prins, University of Illinois College of Medicine for providing the AR antiserum, Walter Pagel for editorial advice, and the National Institutes of Health (grants ES-08075 and HD-40397) for support of much of the research reviewed in this chapter.

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11. Ewing, L. L., Desjardins, C., Irby, D. C., and Robaire, B. (1977). Synergistic interaction of testosterone and oestradiol inhibits spermatogenesis in rats. Nature 269, 409–411. 12. Chowdhury, M., Tcholakian, R. K., and Steinberger, E. (1980). Reevaluation of effects of estradiol benzoate on the production of testosterone and luteinizing hormone in the rat. Endocrinology 106, 1311–1316. 13. Meistrich, M. L., Wilson, G., Ye, W.-S., Thrash, C., and Huhtaniemi, I. (1996). Relationship among hormonal treatments, suppression of spermatogenesis, and testicular protection from chemotherapy-induced damage. Endocrinology 137, 3823–3831. 14. Mahato, D., Goulding, E. H., Korach, K. S., and Eddy, E. M. (2000). Spermatogenic cells do not require estrogen receptor-alpha for development or function. Endocrinology 141, 1273–1276. 15. Eddy, E. M., Washburn, T. F., Bunch, D. O., Goulding, E. H., Gladen, B. C., Lubahn, D. B., and Korach, K. S. (1996). Targeted disruption of the estrogen receptor gene in male mice causes alteration of spermatogenesis and infertility. Endocrinology 137, 4796–4805. 16. Hess, R. A., Bunick, D., Lee, K. H., Bahr, J., Taylor, J. A., Korach, K. S., and Lubahn, D. B. (1997). A role for oestrogens in the male reproductive system. Nature 390, 509–512. 17. Robertson, K. M., O’Donnell, L., Simpson, E. R., and Jones, M. E. (2002). The phenotype of the aromatase knockout mouse reveals dietary phytoestrogens impact significantly on testis function. Endocrinology 143, 2913–2921. 18. Ebling, F. J., Brooks, A. N., Cronin, A. S., Ford, H., and Kerr, J. B. (2000). Estrogenic induction of spermatogenesis in the hypogonadal mouse. Endocrinology 141, 2861–2869. 19. Li, H., Papadopoulos, V., Vidic, B., Dym, M., and Culty, M. (1997). Regulation of rat testis gonocyte proliferation by plateletderived growth factor and estradiol: Identification of signaling mechanisms involved. Endocrinology 138, 1289–1298. 20. Pentikainen, V., Erkkila, K., Suomalainen, L., Parvinen, M., and Dunkel, L. (2000). Estradiol acts as a germ cell survival factor in the human testis in vitro. J. Clin. Endocrinol. Metab. 85, 2057–2067. 21. Bremner, W. J., Millar, M. R., Sharpe, R. M., and Saunders, P. T. K. (1994). Immunohistochemical localization of androgen receptors in the rat testis: Evidence for stage-dependent expression and regulation by androgens. Endocrinology 135, 1227–1234. 22. Zhou, Q., Nie, R., Prins, G., Saunders, P., Katzenellenbogen, B., and Hess, R. (2002). Localization of androgen and estrogen receptors in adult male mouse reproductive tract. J. Androl. 23, 870–881. 23. Vornberger, W., Prins, G., Musto, N. A., and Suarez-Quian, C. A. (1994). Androgen receptor distribution in rat testis: New implications for androgen regulation of spermatogenesis. Endocrinology 134, 2307–2316. 24. Johnston, D. S., Russell, L. D., Friel, P. J., and Griswold, M. D. (2001). Murine germ cells do not require functional androgen receptors to complete spermatogenesis following spermatogonial stem cell transplantation. Endocrinology 142, 2405–2408. 25. Maiti, S., Meistrich, M. L., Wilson, G., Shetty, G., Marcelli, M., McPhaul, M. J., Morris, P. L., and Wilkinson, M. F. (2001). Irradiation selectively inhibits expression from the androgendependent Pem homeobox gene promoter in Sertoli cells. Endocrinology 142, 1567–1577. 26. Kliesch, S., Penttila, T. L., Gromoll, J., Saunders, P. T., Nieschlag, E., and Parvinen, M. (1992). FSH receptor mRNA is expressed stage-dependently during rat spermatogenesis. Mol. Cell. Endocrinol. 84, R45–R49. 27. Orth, J., and Christensen, A. (1978). Autoradiographic localization of specifically bound 125I-labeled follicle-stimulating hormone on spermatogonia of the rat testis. Endocrinology 103, 1944–1951.

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C H A P T E R

24 Long-Term Cultures of Mammalian Spermatogonia MARIE-CLAUDE C. HOFMANN

MARTIN DYM

Department of Biology, The University of Dayton, Dayton, Ohio

Department of Cell Biology, Georgetown University Medical Center, Washington, D.C.

spermatogenesis. By 6 days postpartum (dpp), these cells migrate to the basement membrane of the seminiferous cords and become spermatogonial stem cells, which are at the origin of spermatogenesis in the prepubertal and adult animal. Spermatogonial stem cells are unipotent: They are at the origin of only one cell lineage that will ultimately produce spermatozoa. They are capable of both selfrenewal and differentiation, and they can generate sperm production when transplanted into the seminiferous tubules of an infertile male [4, 5]. Currently, our understanding of the molecular mechanisms that control the development of spermatogonial stem cells into mature sperm is limited. Also, the cues that make these stem cells choose between self-renewal and differentiation are unknown. Our poor understanding derives from the limited quantity of cells available for experimentation and the lack of markers uniquely expressed by these cells. Spermatogonial stem cells express surface markers such as α6 and β1 integrins [6–8]. They also express the GFRα-1/Ret receptor system, which is activated by the binding of glial cell line–derived neurotrophic factor (GDNF) [9, 10]. GDNF plays a crucial role in spermatogonial stem cell proliferation since transgenic mice overexpressing this factor show an accumulation of these cells in the seminiferous epithelium [11]. As they differentiate, the spermatogonia lose the expression of GFRα-1 and acquire the surface receptor c-kit, which binds to stem cell factor (SCF, kit ligand) produced by Sertoli cells [12–14]. Other proteins

I. INTRODUCTION II. ISOLATION OF SPERMATOGONIA III. SPERMATOGONIA IN ORGAN CULTURES AND COCULTURES WITH SERTOLI CELLS IV. ESTABLISHMENT OF SPERMATOGONIAL CELL LINES V. PROLIFERATION AND DIFFERENTIATION OF SPERMATOGONIA IN A CONTROLLED ENVIRONMENT VI. CONCLUSION References

I. INTRODUCTION In the mouse, the germline originates from primordial germ cells (PGCs), which are recognizable by 7.5 days postcoitus (dpc) as a small cluster of alkaline phosphatase-positive cells in the extraembryonic mesoderm, close to the primitive streak [1, 2]. These cells enter the hindgut mesoderm and migrate through the dorsal hindgut mesentery to reach the developing fetal gonads (the genital ridges) at around 11.5 dpc. Primordial germ cells proliferate while migrating. Once they reach the male genital ridges, the PGCs are enclosed by the somatic Sertoli cells and then testicular cords are formed. At this point, the PGCs become prospermatogonia or gonocytes [3]. The gonocytes proliferate for a few days and then arrest in the G0/G1 phase of the cell cycle until birth. Within a few days after birth, the gonocytes resume proliferation to initiate SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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expressed by spermatogonial stem cells include the transcription factor Oct-4 [15, 16], the cytoplasmic RNA helicase MVH (homologous of the Drosophila Vasa) [17], and Dazl (homologous of the Drosophila Boule) [18]. Whereas Oct-4 is also expressed in embryonic stem cells (ES cells) and in the female germline, Vasa and Dazl seem to be uniquely expressed in the male germline up to meiosis. In mammals, all premeiotic male germ cells, including the stem cells, are called spermatogonia. Two models of spermatogonial development have been proposed for adult rodents (reviewed in [10]). In the first scheme, the Asingle (As) spermatogonia are considered the stem cells of spermatogenesis (Fig. 24.1) [19, 20]. They can renew themselves or can differentiate into Apaired (Apr) spermatogonia that remain connected by an intercellular bridge. The Apr spermatogonia further divide to form chains of 4, 8, or 16 Aaligned (Aal) spermatogonia. The Aal cells will then differentiate into type A1 spermatogonia. The A1 spermatogonia resume division to form A2 to A4 spermatogonia. Next, A4 cells divide to form intermediate (In) spermatogonia and In spermatogonia divide to produce type B spermatogonia. Finally, type B spermatogonia divide to form primary spermatocytes that will enter the process of meiosis. In the other model, the type As—and possibly some of the Apr—spermatogonia are considered reserve stem cells and are called A0 (Fig. 24.2) [21, 22]. These cells are believed to remain quiescent, or at the most divide very slowly, unless an excessive loss of germ cells stimulates their proliferation, for example, after injury or irradiation [23]. In this model, the A1 through A4 spermatogonia (renewing stem cells) retain their stem cell properties, and the A4 spermatogonia are able either to produce an A1 spermatogonia (dedifferentiation) or go forward and form In and type B spermatogonia. Brawley and Matunis recently demonstrated that

FIGURE 24.1 Self-renewal and differentiation of the germline stem cell according to the model of Huckins [19] and Oakberg [20].

FIGURE 24.2 Spermatogonial self-renewal and differentiation according to the model of Clermont and Bustos-Obregon [21].

dedifferentiation also takes place in the Drosophila germ line, since more differentiated spermatogonia could revert to a stem cell phenotype in the appropriate microenvironment [24]. In adult primates, including humans, the reserve stem cells are also called “Adark” spermatogonia due to their high affinity for toluidine blue. These “Adark” spermatogonia undergo differentiation into renewing stem cells called “Apale” spermatogonia. “Apale” spermatogonia undergo several mitotic divisions to form chains that finally differentiate into type B spermatogonia [25–29]. Spermatogonial stem cells and their progeny are contained in the basal part of the germinal epithelium, in contact with the basement membrane, and they are in close association with the somatic Sertoli cells [30]. We know that spermatogenesis is a complex and highly organized process that is regulated at multiple steps. Regulatory mechanisms mediated by growth factors produced by the Sertoli cells induce or inhibit the proliferation, differentiation, and further development of the germ cells [31, 32]. The ability to isolate, culture, and manipulate spermatogonial cells in vitro would allow us to unravel the molecular mechanisms driving spermatogenesis and to characterize the signaling pathways, inducing spermatogonial differentiation versus self-renewal. This, in turn, could help us understand the origin of certain testicular neoplasias and the causes of male infertility. To research these areas, an in vitro system in which spermatogonial cells could be maintained in long-term cultures would be ideal. The present chapter is a review of the efforts made during the past 40 years to improve the culture conditions of type A spermatogonia, from organ cultures and cultures on feeder layers of Sertoli cells, to the establishment of spermatogonial

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cell lines, and finally to the attempts to culture these cells alone in a controlled environment.

II. ISOLATION OF SPERMATOGONIA Different methods for isolating spermatogonia are now available. The Staput method was originally devised by Miller and Phillips [33] and used by Lam et al. [34] to separate and characterize subpopulations of germ cells. The method was further refined by Bellve et al., allowing the recovery of purified germ cell populations at defined steps of differentiation [35, 36]. In this procedure, adult mouse or rat testes are decapsulated and dissociated by a two-step enzymatic digestion to remove most of the somatic cells. The cells are then separated by sedimentation velocity at unit gravity at 4°C by means of a 2–4% BSA gradient in culture medium. The cells in suspension are first bottomloaded into a separation chamber, and then a gradient of 2–4% BSA is formed under the germ cell layer. The cells are allowed to sediment for a standard period of 2.5 hr, and 35 fractions of 15-mL volume are collected at 90-sec intervals. The cells in each fraction are examined with a phase-contrast microscope. Thus, fractions containing type A spermatogonia can be pooled and spun down by low-speed centrifugation and then resuspended in culture medium. The method has recently been improved to obtain a germ cell population of higher purity [37, 38]. After Staput separation, the spermatogonial fractions are subjected to differential plating for 4 hr at 34°C in a cell culture dish coated with fetal calf serum (FCS). Adherent contaminating Sertoli cells and peritubular myoid cells were thus eliminated. The type A spermatogonia, which remain in suspension, can be collected and washed in culture medium. This technique allows for the isolation of type A spermatogonia with a purity up to 95% (Fig. 24.3). Another improvement consists of using 6-day-old mice (or 9-day-old rat) testes. We recently showed that this age gives the optimal number of As and Apr spermatogonia, as indicated by their expression of the GFRα-1 receptor [39] (Fig. 24.4). The method of centrifugal elutriation to separate germ cells was originally developed by M. Meistrich [40]. In this technique, a single-cell suspension of germ cells prepared by enzymatic digestion is placed into a centrifuge, in an elutriator chamber. The cells are subjected to a fluid flow opposite to their sedimentation. A gradient of flow rates is established and a distinct cell population will move to a position where the fluid flow rate balances exactly its sedimentation rate.

FIGURE 24.3 Phase-contrast micrograph of freshly isolated type A spermatogonia (see color plate). Cells are spherical and contain organelles in a perinuclear location. Nuclei contain chromatin clumps associated with the nucleoli (×780).

The fluid flow rate or the rotor speed can be adjusted so that the sedimentation rate for these particular cells become lower than the fluid flow rate. The cells will then leave the chamber in a centripetal direction and can be collected [41]. After elutriation, the cells can be further purified using equilibrium density centrifugation in Percoll gradients [42, 43]. Percoll gradients have also been used to purify germ cells populations after Staput sedimentation at unit gravity [44]. Percoll gradients alone have also been used to isolate distinct populations of rat and mouse germ cells [45, 46]. The method has been modified to isolate specifically type A spermatogonia from adult rats [47]. In this procedure, vitamin A–deficient (VAD) rat testes

FIGURE 24.4 Immunostaining of type A spermatogonia for the GFRα-1 receptor in a section of seminiferous tubules obtained from a 6-day-old mouse (see color plate). As and Apr spermatogonia show intense staining. Cytoplasmic bridges can be seen between two spermatogonia (stars). N = nucleus (×1000). (From Dettin et al. [39].)

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are decapsulated and the tubules subjected to a twostep enzymatic digestion to eliminate most somatic cells. To further purify the spermatogonial population, the cell suspension is plated on peanut agglutinin-coated dishes to remove residual Sertoli and peritubular myoid cells. The nonadherent cells are then separated on a discontinuous Percoll gradient. This method ensures the isolation of a spermatogonial population of high purity (70–90%). The procedure has been adapted for the isolation of type A spermatogonia from prepubertal rat testes [48]. Percoll isolation can also be used to isolate bovine type A spermatogonia [49, 50]. Spermatogonia from different adult mammalian species can also be isolated using the technique of magnetic beads and an antibody recognizing the c-kit receptor [51]. In this procedure, a single-cell suspension of germ cells is obtained after a two-step enzymatic digestion. The suspension is then labeled with a rabbit anti–c-kit antibody. Magnetic beads coated with goat anti-rabbit IgG are used to retrieve the c-kit–positive cells. However, there is now evidence that mouse spermatocytes express c-kit as well [52, 53]. To isolate c-kit–positive type A spermatogonia, specifically Apr and Aal cells, we recently used the magnetic bead technique with 6-day-old mice [54]. Using this method, the purity of the isolated population was greater than 95%. We also took advantage of the specific expression of the homophilic adhesion molecule Ep-CAM by type A and type B spermatogonia [55] and isolated these cells successfully from mouse adult testes (Fig. 24.5). The method of isolation using magnetic beads is rapid and effective. Although the number of cells recovered is much lower than with the Staput procedure or with elutriation, the purity of the isolated cell populations is extremely high. We tested the viability of the isolated cells by culturing them with Sertoli cell feeder layers. Viable germ cells, still expressing the c-kit receptor, could be visualized after 25 days in vitro. When the

cells were cultured in Sertoli cell tubules on a layer of Matrigel, they incorporated BrdU for at least 1 week [54]. We next adapted the magnetic bead method in order to specifically isolate the germline stem cell (As) from their direct progeny, the Apr and Aal spermatogonia. We know that GFRα-1, the receptor for GDNF, is only expressed by As and possibly Apr spermatogonia. We successfully separated these cells using the Staput method followed by magnetic bead isolation using a GFRα-1 receptor antibody. We further characterized the isolated cells and demonstrated that they express Ret, the tyrosine kinase transmembrane receptor that mediates the intracellular response to GDNF via GFRα-1. In addition, this population contains c-kit–positive and c-kit–negative cells (Fig. 24.6). Thus, using these surface markers, we could characterize more precisely the first steps of spermatogenesis. We observed that the primitive stem cells (As) are GFRα-1+/Ret+/c-kit−, their direct progeny Apr spermatogonia are GFRα-1+/Ret+/c-kit+ and the Aal spermatogonia are GFRα-1−/Ret+/c-kit+ (Fig. 24.7). Therefore, a pure testis stem cell population can be isolated using magnetic beads coated with a GFRα-1 antibody, followed by a subtractive isolation using a c-kit antibody. Direct isolation by subtraction of

FIGURE 24.5 Isolation of type A spermatogonia with the immunomagnetic bead method (see color plate). (A) Type A spermatogonium isolated with beads coated with an Ep-CAM antibody and subsequently stained for c-kit. The spermatogonium is still associated with the magnetic beads (arrowheads). Bright-field microscopy (×600). (B) Negative control without the c-kit primary antibody. Bright-field microscopy (×600). (From van der Wee et al. [54].)

FIGURE 24.6 Selective isolation of undifferentiated type A spermatogonia using magnetic beads coated with a GFRα-1 antibody (see color plate). These cells were further stained for the presence of c-kit or Ret. (A) All cells are GFRα-1+, as shown by the beads still attached (arrowheads) to the Ret+ cells (purple staining). (B) As spermatogonia (GFRα-1+, c-kit–). (C) Apr or Aal spermatogonia (GFRα-1+, c-kit+) (red staining). (D) Negative control (no first antibody).

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FIGURE 24.7 Expression of selected surface receptors during the first steps of spermatogenesis.

c-kit–positive cells is not possible because residual Sertoli cells are also c-kit negative, like the stem cells. The magnetic bead technique allows for the recovery of extremely pure cell populations; however, the yield remains low (about 1/10 of the yield estimated by De Rooij and Russell [10]). Other techniques such as fluorescence-activated cell sorting (FACS) have been used that increase the number of stem cells recovered. Shinohara et al. observed that spermatogonial stem cells preferentially bind to the extracellular matrix component laminin through the α6 and β1 integrins [8]. When mouse testis cells expressing these integrins were selected using magnetic beads or adherence to extracellular matrix proteins, the resulting population contained a fivefold enrichment in stem cells, as detected by testis transplantation experiments. In addition, the authors found that the testes of experimental cryptorchid mice are enriched 25-fold for spermatogonial stem cells [56] in comparison to normal testis. Finally, when testis cells from cryptorchid mice are isolated using parameters such as low intracellular complexity, low expression of αv integrins, no expression of the c-kit receptor, and high expression of α6 integrins, the cell population isolated by FACS was enriched 152- to 166-fold in stem cells [57]. Thus, while enrichment rather than specific isolation is performed using these criteria, FACS is the method of choice to obtain high yields of viable, functional stem cells (1 cell in 30–40 cells). Stem cells in many tissues can be recognized and isolated as a “side population of cells” (SP cells) that are characterized by their ability to efflux rapidly the DNA-binding dye Hoechst 33342 [58, 59]. Recent studies have implicated the ABCG2/Bcrp1 transporter as a Hoechst efflux pump [60]. The dye is excited at 350 nm and its fluorescence is measured at 450 (blue) and 675 nm (red). By using dual-wavelength flow cytometric analysis, the SP population appears as a small population of negative to poorly stained blue fluorescent cells, representing 0.03–0.1% of the total population in the bone marrow, lung, skeletal muscle, brain, mammary gland, and retina [58, 61–65]. A side population (SP) of cells has been recently isolated from

mouse seminiferous tubules using FACS and characterized [66, 67]. The cells represent about 1.3% of the total population. However, the identity of these cells remains controversial. Although one study demonstrated that the testis SP cells do not exhibit the makeup of surface markers typical of spermatogonial stem cells and are unable to repopulate an infertile testis, the other study demonstrates a 13-fold enrichment of stem cell activity and complete spermatogenesis in the recipient testes after SP cell transplantation. Whereas no marker(s) specific for spermatogonial stem cells have been identified to date, our progress in characterizing a particular combination of molecules at their surface now allows us to isolate populations that are pure enough for in vitro studies. Moreover, recent attempts to manipulate primordial germ cells, gonocytes, and spermatogonia in microtiter culture plates have shown that it is possible to analyze signaling pathways even when the size of the population studied is limited [68, 69]. Thus, using specific isolation methods, genetic manipulations in small-scale culture, and subsequent testis transplantations, the biology of spermatogonial stem cells will soon be better understood.

III. SPERMATOGONIA IN ORGAN CULTURES AND COCULTURES WITH SERTOLI CELLS The fact that spermatogonia can be maintained in vitro and that germ cell differentiation in the petri dish is possible when these cells are in close association with Sertoli cells has been recognized since the pioneering studies of Steinberger et al. [70–72]. While in vitro techniques cannot fully replace in vivo conditions, they offer the possibility of studying germ cell development in simplified environments where diverse factors can be tested. Initially, these investigators established organ cultures where fragments of immature rat testicular tissue (~1 mm3) were grown on a 2% strip of agar placed on top of a wire grid platform in a

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60-mm-diameter petri dish [70]. The amount of medium used was sufficient to wet the lower part of the agar strip. After optimization of this culture system, the authors reported gonocyte differentiation and the appearance, growth, and survival of primitive type A spermatogonia for a period as long as 6 months [70]. Primitive type A spermatogonia were able to undergo mitosis for at least 4 weeks [70, 71]. These studies also showed for the first time that pituitary gonadotropins are not directly necessary for the initiation of spermatogenesis or for germ cell differentiation, at least up to the pachytene spermatocyte stage. This culture system was later refined, with the agar strip often replaced by a Millipore filter [73, 74], and it is still used by investigators to study the development of spermatogonia in various conditions and the influence of Sertoli cell–derived factors, in particular when FSH is added to the culture medium [75–79]. In 1976, Eddy and Kahri took a different approach and cultured fragments of seminiferous tubules directly onto the plastic surface of the petri dishes [80]. They observed the remodeling of the fragments into sheets and clusters of cells by phase-contrast, transmission, and scanning electron microscopy. In these cultures, spermatogonia recognized and adhered to Sertoli cells, and they conserved their elongated shape and were connected by typical cytoplasmic bridges, comparable to the in vivo situation (Fig. 24.8) [81, 82]. The ultrastructure of the cells and their association with Sertoli cells was further studied by Palombi et al. [83]. Because the physiological cell–cell interactions are well maintained in these type of cultures, they are now widely used for the study of the influence of hormones and growth factors on spermatogonial proliferation and differentiation, in particular, follicle-stimulating hormone (FSH) and stem cell factor (SCF) [84–88]. For most of these experiments, the technique of transillumination-assisted microdissection has been used to isolate stage-specific segments of seminiferous tubules [89, 90]. Cultures of seminiferous tubules have been now adapted for other species such as monkey and human [91–97]. However, working with selected populations of spermatogonia in vitro is crucial to understanding the mechanisms underlying the first steps of spermatogenesis. This led to the technique developed by Nishimune’s group in Japan [98]. By producing artificially induced cryptorchid testes in adult mice, these investigators obtained seminiferous tubules devoid of germ cells, except for undifferentiated type A spermatogonia (As, Apr, and Aal). These seminiferous tubules were then cultured in organ culture on millipore filters as described earlier. These studies revealed the importance of FSH, retinoids, and soluble factors

FIGURE 24.8 Chain of type A spermatogonia adhering to Sertoli cells in fragment cultures of seminiferous epithelium. The cells were from a 9-day-old rat after 6 days in culture. SEM, ×2300. (From Eddy and Kahri [80].)

for the proliferation and differentiation of undifferentiated type A spermatogonia via Sertoli cell activation [99–101]. Using the same culture system, they also investigated the harmful effects of the Steel mutation (lack of production of SCF) on spermatogonial differentiation in vitro [102]. Absence of differentiating type A spermatogonia and subsequent germ cells could later be obtained by vitamin A deprivation in mice and rats [103, 104]. In addition, the same deficiency is observed in jsd/jsd and in Sl17H mutant mice [105–109]. Thus vitamin A deprivation and mutant mice can be used to study the behavior of undifferentiated spermatogonia (As, Apr, and Aal) in vivo and in vitro, without interference from more differentiated germ cells. Another method to ensure that seminiferous tubules contain only early spermatogonia is to isolate and culture fragments of seminiferous tubules from 3-day-old mice. Using this protocol, Nikolova et al. stressed the importance of leukemia inhibitory factor (LIF) for spermatogonial proliferation [110]. While cultures of seminiferous tubules respect the overall architecture of the germinal epithelium and the cell–cell interaction that takes place in the testis, in vitro systems that are better defined and adapted for the analysis of the cellular and molecular mechanisms controlling spermatogenesis had to be established. Thus, protocols were devised in which the germ cells and Sertoli cells were first dissociated from each other and then allowed to reassociate during culture [111]. Earlier studies by Grund et al. [112] and EscalanteAlcalde and Merchant-Larios [113] showed that dissociated germ cells and somatic cells from newborn rat and fetal mouse gonads can reaggregate and form seminiferous cords in culture. Our own studies with immortalized somatic and spermatogonial cell lines confirmed this reaggregation property in the 10-day-old mouse testis [46] (Fig. 24.9). In general, the formation

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FIGURE 24.9 Reaggregation of immortalized germ cells (asterisks), Sertoli cells, and peritubular cells in vitro (see color plate). Germ cells are always surrounded by the somatic cells producing the basement membrane protein laminin (stained brown with the immunoperoxidase technique).

FIGURE 24.10 Formation of tubules by the Sertoli cell line SF7 on a layer of Matrigel.

of cords and tubules by primary Sertoli cells as well as Sertoli cell lines is facilitated when the cells are cultured on a layer of Matrigel [114–116] (Fig. 24.10). We also have shown that it is possible to use these in vitro sleeves to maintain the proliferation of type A spermatogonia in culture for at least 1 week [52] (Fig. 24.11). Type A spermatogonia, including the stem cells, can be cultured for a short period of time on monolayers of freshly isolated Sertoli cells. However, without the addition of specific growth factors such as epidermal

growth factor (EGF), LIF, forskolin, and oncostatin, self-renewal is very limited [69, 117]. Without these factors, the stem cells will differentiate and produce chains of Aal cells connected by cytoplasmic bridges [118, 119]. They will eventually become spermatocytes [45, 54, 120]. Cultures of type A spermatogonia on Sertoli cell monolayers have been useful for studying apoptosis mediated by Fas or toxicants [121, 122], to assess the effects of activin [123], to study components of the Notch pathway [38], and to assess the

FIGURE 24.11 Incorporation of BrdU by mouse type A spermatogonia in Sertoli cell tubules after 1 week of coculture (see color plate). (A) Sertoli cell tubule containing type A spermatogonia. Phase contrast (×100). (B) Same tubule in fluorescence microscopy showing type A spermatogonia labeled with the red fluorescent dye DiI (×100). (C) Same tubule in fluorescence microscopy showing type A spermatogonia expressing BrdU and revealed with FITC (×100). (From van der Wee et al. [54].)

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role of p53 in the control of apoptosis [69]. These types of cocultures are now used to cultivate human and porcine premeiotic male germ cells in vitro [124, 125]. Several paracrine growth factors produced by Sertoli cells in vivo have been recently identified and their role in spermatogonial survival and proliferation demonstrated. In particular, the transmembrane form of SCF, the ligand for c-kit, supports the survival of primordial germ cells, whereas the soluble form stimulates the proliferation of primordial germ cells [126] and DNA synthesis in type A spermatogonia [127–129]. LIF is a survival and proliferation factor for primordial germ cells and gonocytes [110, 130, 131]. LIF has also been shown to maintain long-term cultures of embryonic stem cells in an undifferentiated stage [132, 133] and might have a similar function in maintaining the stem cell potential of certain spermatogonia [134]. Another factor important for the survival and proliferation of primordial germ cells and possibly spermatogonia in culture is basic fibroblast growth factor (bFGF) [135, 136], which is produced by Sertoli cells. Sertoli cells are also the primary source of testicular inhibin, a paracrine regulator of various cell types [137]. α-Inhibin may also have a role in the regulation of spermatogonial cell number and differentiation [137–139]. Transforming growth factor β (TGFβ), produced by Sertoli cells, has also been linked to germ cell differentiation [111, 140, 141]. Thus, although their function in spermatogenesis has not been clearly elucidated, any of these molecules, alone or in combination, could play a crucial role in spermatogonial proliferation and/or differentiation. Therefore, we established Sertoli cells lines that were screened specifically for the expression of these growth factors [119]. In addition to morphological characteristics of Sertoli cells in light and electron microscopy, these cell lines exhibit a molecular phenotype similar to that of Sertoli cells in vivo. The genes expressed include SGP-2, α-inhibin, GATA-1, and genes coding for major growth and differentiation factors such as SCF, LIF, bFGF, and TGFβ. Moreover, the expression of the orphan nuclear receptor SF-1 by all of the cell lines indicates that transactivation of malespecific genes could occur in these cells, which might influence the development of germ cells in coculture. Although some of these factors are produced by STO fibroblasts, the latter cannot always be used to study somatic cell–germ cell interactions because they do not express Sertoli cell-specific genes such as α-inhibin, SF-1, SGP-2, or GATA-1. When these Sertoli cell lines are used as feeder layers, mouse As spermatogonia differentiate into Apr and Aal cells connected by intercellular bridges, as they would do with freshly isolated Sertoli cells (Fig. 24.12) [119].

FIGURE 24.12 Cocultures of type A spermatogonia on Sertoli cells feeder layers (see color plate). A single-cell suspension of freshly isolated type A spermatogonia was seeded on monolayers of Sertoli cells. After 3 days, signs of spermatogonial differentiation were observed. (A) Paired and aligned type A spermatogonia after 3 days of culture on a Sertoli cell line feeder layer. The germ cells were stained for the c-kit receptor. Intercellular bridges are apparent (arrows) (×400). (B) For comparison, aligned type A spermatogonia after 3 days of culture on primary Sertoli cell feeder layers. The cells were stained with an antibody to LIF receptor and, as expected, both Sertoli cells and germ cells express the protein (×400). (From Hofmann et al. [119].)

Long-term mixed cultures of somatic cells and male germ cells have been recently used. Optimally, the mixed testis cells are cultured over a feeder layer of STO fibroblasts. These culture systems are beneficial for the long-term survival of mouse spermatogonial stem cells in vitro, since the stem cell component of the cell populations could repopulate an infertile testis after 4 months in vitro [142, 143]. Rat spermatogonial stem cells can also survive in these types of culture, but only short term (about 5 days) [144]. Bovine spermatogonial stem cells isolated from newborn bulls can develop into spermatids after several weeks when cocultured with Sertoli cells [145]. More recently, Izadyar et al. established culture conditions in which bovine spermatogonial stem cells could self-renew and differentiate for at least 150 days on a somatic cells feeder layer [146]. The spermatogonia formed paired cells and chains of cells, then large colonies able to produce differentiating cells that eventually acquired the characteristics of spermatids.

Chapter 24 Long-Term Cultures of Mammalian Spermatogonia

In summary, cultures of spermatogonia on somatic cell feeder layers have already allowed investigators to generate much data concerning spermatogonial differentiation. Recent progress includes coculture systems that allow the maintenance and self-renewal of spermatogonial stem cells and systems that allow the proliferation and differentiation of c-kit–positive spermatogonia. These new culture systems are of paramount importance to increase the pool of cells available for experimentation and for the study of spermatogonial differentiation beyond meiosis. However, longterm expansion resulting in a large population of cells able to differentiate in vitro has not yet been achieved. While progress has been made in defining soluble factors essential to their proliferation, investigators still need to find the correct conditions for mimicking the environment provided by the seminiferous epithelium.

IV. ESTABLISHMENT OF SPERMATOGONIAL CELL LINES Over the years, we have seen an increase in the use of primary cultures of Sertoli cells to maintain spermatogonia in culture and to investigate the role of Sertoli cells in spermatogonial proliferation and differentiation. However, this methodology has been limited by the fact that the cell populations are not pure and that the production of Sertoli cell factors eventually stops, leading to the death of the germ cells within 15 days after isolation. Therefore, the establishment of germ cell lines that can survive without feeder layers, but still respond to the appropriate growth factors in vitro, would greatly enhance our ability to study the molecular events leading to germ cell proliferation and differentiation. Moreover, if these lines were to repopulate a mouse testis and to contribute to the recipient’s progeny, these lines would be great tools for genetic manipulations of the germline and the production of transgenic mice. The first germ cell–derived cell lines were obtained from human testicular tumors, called embryonal carcinoma and teratocarcinoma [147–149]. These lines are characterized by their remarkable pluripotentiality. The human embryonal carcinoma (EC) cell line NTERA-2, for example, is able to differentiate into neurons and other cell types when treated with retinoic acid and bone morphogenetic proteins (BMPs) [150]. Another cell line, NCCIT, was established by Shinichi Teshima (National Cancer Institute, Tokyo, Japan) in 1985 from a mediastinal mixed germ cell tumor [151]. The undifferentiated cells are equivalent to a stage intermediate between seminoma and embryonal carcinoma, and they resemble spermatogonia. In response

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to retinoic acid, they will differentiate into derivatives of all three embryonic germ layers (i.e., ectoderm, mesoderm, and endoderm) and extraembryonic cell lineages [152]. Damjanov et al. suggested that this cell line could be a malignant replica of human cleavage stage embryonic cells. Thus, the histogenesis of germ cell tumors (GCTs) reflects embryonic cell differentiation, and derived cell lines are useful as model systems to study the early stages of mammalian embryogenesis [153, 154]. However, they cannot be used to study normal germ cell proliferation and differentiation because of their strong aneuploidy and their malignant phenotype. Stable transfections of immortalizing, nontransforming genes such as the simian virus 40 large T antigen gene (SV40 LTAg), the polyoma virus LTAg gene, or the adenovirus E1A gene have been used to facilitate the production of cell lines from various mouse and human somatic tissues [46, 155, 156]. In 1992, we established several testicular cell lines after stable transfection of the SV40 LTAg gene into cells isolated from a prepubertal 10-day-old mouse testis [46]. The LTAg protein is known to inactivate wild-type Rb and p53, thus promoting entry into S phase and immortalizing the cells [157, 158]. One of the cell lines established appeared to have characteristics of type B spermatogonia and preleptotene spermatocytes and was called GC-1spg (Fig. 24.13). This was the first immortalized, nontumorigenic germ cell line established. The cells expressed the germ cell markers cytochrome ct and LDHC4 isozymes, and have thus facilitated the detailed study of the LDHC4 promoter [159, 160]. Further, they have been used to investigate signal transduction pathways in the early stages of spermatogenesis [161, 162], to study differential gene expression [163], to assess the cytotoxicity of certain environmental chemical compounds [164, 165] and to investigate spermatogonial apoptosis [166].

FIGURE 24.13 Appearance of the germ cell line GC-1spg in phase-contrast microscopy. (From Hofmann et al. [46].)

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However, the germ cell line GC-1spg does not appear to be able to differentiate through meiosis. GC-1spg cells are aneuploid, a phenomenon due to the high expression of LTAg that might have promoted chromosome instability. We attempted to modulate the immortalizing properties of the SV40 LTAg by using the LTAg-binding properties of the p53 protein. It has been shown that an excess of wild-type p53 protein is able to abolish the proliferative function of LTAg when both molecules are expressed in pre-crisis mouse fibroblasts [167, 168]. Using this strategy, we established two other germ cell lines termed GC-2spd and GC-3spd after transfecting primary testicular cells enriched in preleptotene spermatocytes [169, 170]. Flow cytometric analysis of the DNA content of these cells indicated that part of the population was able to differentiate and to become haploid in certain conditions [171]. These results were confirmed by analysis of chromosome spreads [169]. However, upon long-term culture (about 2 years), these cells did not conserve their ability to differentiate [172]. This may be due to cumulative changes induced by the LTAg over time that prohibit reversal of cellular immortalization, as shown in other systems [173]. The nature of these changes is so far unknown. Nevertheless, at the time, these results showed that it is possible to transfect, immortalize, and differentiate mouse testicular germ cells in vitro. The cell line GC-2spd is still used for its spermatocyte characteristics [174–176]. Some years later, the spermatocytic cell line GC-4spc was established by Tascou et al. using the LTAg as an immortalizing agent and the neomycin gene under control of the PGK-2 promoter to select specifically for spermatocytes [177]. This interesting strategy allows for the immortalization of germ cells from a specific stage of development. In addition to PGK-2, the GC4spc cells expressed the A-myb, the TSPY and the proacrosin genes. The authors compared the expression of PGK-2 and TSPY genes in GC-1spg and GC-4 spc (Fig. 24.14) and confirmed that differential expression of these genes corresponded to the stage at which the cells had been immortalized. Whereas a high transcriptional activity of the PGK-2 promoter was observed in the GC-4spc cell line, no activity of this promoter could be detected in the GC-1spg cell line. In contrast, the TSPY promoter showed a higher transcriptional activity in the GC-1spg cell line than in the GC-4spc cell line. Thus, using this strategy, it is now possible to immortalize germ cells “frozen” at different stages of differentiation. More recently, van Pelt et al. established two rat spermatogonial cell lines that they called GC-5spg and GC-6spg [178]. Again, stable transfections of the large T antigen gene were used to immortalize the cells. The

FIGURE 24.14 Differential promoter activity for TSPY and PGK-2 in the GC-1spg and GC-4spg germ cell lines. (From Tascou et al. [177].)

germ cells used were isolated from the testes of VAD rats, thus containing only As, Apr, and Aal spermatogonia beside somatic cells. Cells fractions containing 80% or more type A spermatogonia were used for the transfection experiments, increasing the probability of immortalizing germ cells. The cell lines obtained expressed Hsp90α and Oct-4, which are well known germline markers [15, 16, 179, 180]. The cells were also negative for the expression of c-kit, the receptor for SCF, indicating that they were immortalized at an earlier stage than Aal spermatogonia [181]. The cells could be cultured for 3 years without losing their phenotype. More importantly, cells of both lines could repopulate the testes of irradiated recipient mice after transplantation (Fig. 24.15). These immortalized spermatogonia were able to migrate to the basement membrane and formed clusters on the basal side of Sertoli cells. Although they never formed tumors, indicating that they are not transformed, the cells of both lines were unable to differentiate inside the recipient seminiferous tubules. Despite this progress, the lack of differentiation of GC-5spg and GC-6spg cell lines indicates that the introduction of immortalizing genes of viral origin into germ cells definitely alters their normal physiology. This problem theoretically could be overcome by using mutant conditional immortalizing viral genes. The technique allows the generation of continuously proliferating cell lines capable of differentiation after inactivation of the immortalizing gene. For example, the SV40 mutant temperature-sensitive (ts) gene tsA58 has been used in the generation of a variety of conditionally immortal somatic cell lines, including Sertoli cells [182–186]. However, the temperature at which the viral oncogene is turned off, allowing reversion to

Chapter 24 Long-Term Cultures of Mammalian Spermatogonia

FIGURE 24.15 Tubular cross sections of nude mice 8 weeks after transplantation with an immortalized spermatogonial cell line (see color plate). The nuclear staining show expression of the large T antigen protein by the transplanted germ cells (arrowheads). Sertoli cell nuclei are indicated by asterisks. (From van Pelt et al. [178].)

the normal phenotype, is 39°C, a temperature not suitable for the maturation of mouse spermatogonia that differentiate best at 34°C. To be able to modulate the rate of proliferation of the spermatogonia and to allow them to revert to a normal phenotype if necessary, we placed the immortalizing SV40-LTAg gene under the control of a promoter inducible with ponasterone A, an analogue of ecdysone. We obtained a new germ cell line called C18-4. In the presence of the hormone analogue in the culture medium, the expression of the oncogene was activated; upon its removal, most of the cells stopped proliferating and died. However, after 45–50 passages in culture, the germ cells escaped the hormonal control and continued to express the LTAg even without ponasterone A. Nevertheless, this immortalization strategy allowed us to better modulate the rate of proliferation of these cells in comparison to germ cell lines previously established [46, 169]. Morphologically, the cells resemble type A spermatogonia, with a round nucleus containing little heterochromatin, and a large nucleus-to-cytoplasmic ratio (Fig. 24.16). The cells appear to be adherent in tissue culture and acquire a polygonal shape after spreading. To ascertain that this line is of germ cell origin, we probed the cells for the expression of Dazl by immunocytochemistry, immunoprecipitation, and Northern blotting. In the mouse, the Daz/Dazl gene family is represented only by the autosomally located gene Dazl [187, 188]. In knockout animals, the loss of Dazl causes sterility in both sexes, indicating a predominant role in

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FIGURE 24.16 Appearance of the cell line C18-4 in phasecontrast microscopy.

spermatogenesis and oogenesis [189]. Mouse Dazl is the homologue of the Drosophila gene Boule and the human gene DAZL, and encodes a mRNA-binding protein [188, 190, 191]. Although some of the target mRNAs have recently been identified, it is not understood how mDAZL regulates them [192]. Gene expression studies revealed that Dazl transcription starts in gonocytes as early as 1 day after birth, increases dramatically by day 6, and reaches a plateau by day 10 after birth [18]. Immunocytochemistry for Dazl revealed that, in the mouse, the protein is predominantly found in the nucleus of normal gonocytes and spermatogonia. However, as the cells mature toward meiosis, the protein relocalizes and is mainly found in the cytoplasm of spermatocytes [18]. A recent examination of the testes of adult knockout mice using whole mounts of testicular tubules indicates that the Dazl protein might be involved in the differentiation of Aal into A1 spermatogonia, since only As, Apr, and Aal remain in those animals [193]. We showed that the cell line established express the Dazl transcript and produce the Dazl protein, indicating that it is of germ cell origin. Further, because we have used highly purified spermatogonia from 6-day-old mice as the starting material, and because we found the Dazl protein in the nucleus and in the cytoplasm of the immortalized cells, they are type A spermatogonia. In these cells, Dazl formed dot-like structures as previously described by Ruggiu et al., which could be due in part to oligomerization of the protein with itself (Fig. 24.17) [18, 194]. The germline origin of the C18-4 cells was further confirmed by their expression of the transcription factor Oct-4. To further determine the

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No. of cells (in thousands)

350 GDNF Control

300 250 100 150 100 50 0 0

1

2

3

4

5

6

Time (days)

FIGURE 24.19 Proliferation of the C18-4 germ cell line in response to GDNF.

FIGURE 24.17 Expression of Dazl in the nucleus and cytoplasm of the germ cell line C18-4, visualized by the immunoperoxidase technique (arrows). N=nucleus. (see color plate).

phenotype of this cell line, we probed the cells for a surface receptor that is specifically produced by undifferentiated type A spermatogonia, the GFRα-1 receptor for GDNF. The cell line was positive for GFRα-1 using immunocytochemistry and immunoprecipitation (Fig. 24.18). GDNF is a growth factor that acts on several neuronal populations in the developing brain. It is also acting in the developing kidney. GDNF interacts with the GFRα-1 receptor, which in turn mediates stimulation of the Ret receptor tyrosine kinase [195]. GFRα-1 is expressed by germline stem cells in the testis, whereas GDNF is produced by Sertoli cells [9, 11].

FIGURE 24.18 Expression of the GFRα-1 receptor by the germ cell line C18-4, visualized by the immunoperoxidase technique (see color plate).

The C18-4 cells respond to GDNF by a significant increase in proliferation (Fig. 24.19). However, no signs of differentiation in vitro have been observed yet. Moreover, the cells did not express c-kit and did not respond to SCF by proliferating or differentiating. Thus, because the C18-4 cells express the GFRα-1 and Ret receptors, but not c-kit, we can assume that they derive from a spermatogonial stem cell or As [10, 181]. The cells will need to be transplanted into a recipient infertile mouse testis in order to assess their full differentiation potential. The cells have been further characterized by reverse transcriptase–polymerase chain reaction (RT-PCR) using a screen of 36 genes specifically expressed by spermatogonia in a 6-day-old mouse testis [196]. The cells expressed several genes, such as piwi-like and prame-like, that confirmed their identity as undifferentiated spermatogonia with stem cell characteristics [197, 198]. The C18-4 cell line will be useful for analyzing the Ret signaling pathway in germ cells, for promoter studies of genes specifically expressed in spermatogonia, for understanding how Dazl regulates its target mRNAs, and possibly for studying the function of novel genes identified through expression arrays of primary spermatogonial stem cells. In an attempt to establish nontransformed spermatogonial cell lines that are able to proliferate and differentiate in vitro, we also used a strategy that employed the gene encoding mTERT, the catalytic component of mouse telomerase [199, 200]. Cells from normal somatic tissues exhibit a finite proliferative capacity. A prevailing hypothesis is that as cells divide, the physical ends of linear chromosomes (called telomeres) shorten, eventually causing the cells to age and die. Briefly, telomere shortening is due to the inability of the DNA replication apparatus to complete lagging strand synthesis. This results in a loss of

Chapter 24 Long-Term Cultures of Mammalian Spermatogonia

sequence from the 3’-strand of the DNA molecule in each successive round of DNA replication, eventually eliminating essential genes from the distal regions of the chromosome. Indeed many studies have shown that telomere shortening correlates strongly with cellular aging [200, 201]. However, in tissues that continually remodel or regenerate (such as bone, the hematopoietic system, the epidermis, and the germinal epithelium of the testis), stem cells must continuously divide in order to ensure the lifelong rebuilding of the tissue. To counteract the potentially lethal scenario of telomere shortening, stem cells must utilize a mechanism that compensates for the loss of telomeric DNA. This mechanism has been identified: a specialized ribonucleoprotein, telomerase, adds telomeric DNA onto the ends of linear chromosomes [202, 203]. Telomerase is the major pathway for telomere replication in most eukaryotic organisms. In mammals, telomerase activity is naturally found in committed stem cells and in germline stem cells, but is not found in differentiated cells [204–206]. The absence of telomerase activity from differentiated cells explains the reduction in telomere length in aging cell populations. These data have led to the hypothesis that telomeres function as a molecular clock and may provide a way to measure the life span of specific cells. In addition to stem cells, telomerase activity has been identified in immortalized cell lines and cancer cells [207–209]. The strong correlation of telomerase activity with tumor cells suggests that telomerase plays a role in malignant transformation of cells. Telomerase is one of three essential components required to transform human epithelial and fibroblast cells to a malignant phenotype [210]. Ectopic expression of human telomerase functions cooperatively with two known oncogenes, SV40 large T and H-ras, to create a malignantly transformed cell [211]. An alternative hypothesis states that telomerase functions solely to increase the replicative potential of cells [212]. In this model the presence of telomerase activity maintains telomere length and ensures that a cell will continue to divide, but does not lead to malignant transformation of that cell. For example, a retroviral vector expressing a cloned copy of the human telomerase protein component (hTERT) was used to immortalize normal human fibroblast cells [213, 214]. Interestingly these cells do not show any of the hallmarks of cancer cells, such as altered patterns of growth, karyotype abnormalities, and phenotypic modifications. In these experiments, telomerase activity meets the requirements of an immortalizing agent without changing the cellular characteristics of the cell in which it is expressed. Therefore, maintaining telomere length is a requirement for preventing or slowing down cellular aging, but it does not in itself transform

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a cell into a cancer cell. Recently, mouse fibroblasts have been immortalized using a retroviral vector expressing the gene coding for mTERT [212]. Like their human counterpart, the resulting cell clones did not exhibit any of the changes characteristic of cancer, such as altered patterns of growth, karyotype abnormalities, and phenotypic modifications. We and others have recently demonstrated that the male germline stem cells express high levels of telomerase activity [206, 215]. However, this activity is progressively lost during germ cell differentiation. Furthermore, telomerase deficiency leads to a depletion of male germ cells, indicating a role for telomerase in male germline stem cell maintenance [216]. Thus, to obtain germ cell lines that conserve a normal phenotype and can be maintained in vitro, we decided to stabilize telomere length by overexpressing mTERT in mouse type A spermatogonia. We isolated a population of germ cells containing As, Apr, and possibly Aal spermatogonia from 6-day-old mouse testis using the Staput method described earlier in this chapter [35, 128]. To increase the efficiency of gene transfection, we used a retrovirus system to introduce the mTERT gene into the cells [217]. We were able to obtain six cell lines that have conserved their spermatogonial phenotypes for 3 years now, such as a large cell body and a spherical nucleus with a thin rim of cytoplasm and perinuclear organelles (Fig. 24.20). All cell lines were positive for Oct-4 and Dazl. Further, they all expressed the c-kit receptor, which plays a critical role in type A spermatogonia proliferation and differentiation. We treated one of the cell lines, S4, with SCF for up to 21 days. The cells responded to the treatment by autophosphorylation of the c-kit receptor, followed by differentiation toward meiosis. Specifically, they expressed the testis-specific gene lactate dehydrogenase (LDH-C4), the synaptonemal complex protein 3 (SCP3) and produced crossovers (Fig. 24.21). Further, some of these cells could complete meiosis in vitro because up to 33% of the cell population became haploid. The production

FIGURE 24.20 Morphology of telomerase-immortalized spermatogonia (see color plate). (A) After 2 months of culture, the immortalized cells displayed the typical morphology of type A spermatogonia, such as spherical nuclei and organelles in a perinuclear location. (B) Morphology of freshly isolated type A spermatogonia (control). (From Feng et al. [217].)

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FIGURE 24.21 Crossovers shown by Giemsa staining in a spermatogonial cell line induced by SCF (arrowheads) (see color plate). (From Feng et al. [217].)

of pro-acrosomal granules could be observed, which fused into a larger granule that further flattened into an acrosomal cap (Fig. 24.22). Thus, we have established a spermatogonial cell line able to differentiate and undergo meiosis upon SCF stimulation. In conclusion, cell lines representing different “frozen” stages of germ cell development and cell lines able to differentiate beyond meiosis in a defined environment are promising new tools for studying the molecular mechanisms of germ cell proliferation and differentiation, as well as Sertoli cell–germ cell interactions in the testis.

V. PROLIFERATION AND DIFFERENTIATION OF SPERMATOGONIA IN A CONTROLLED ENVIRONMENT To study cell fate decisions in the early phases of spermatogenesis at the molecular level, an in vitro system must be devised whereby type A spermatogonia, including the testis stem cells, can be cultured for a prolonged period of time. The availability of type A spermatogonia that can be maintained in vitro while retaining their differentiation potential is highly desirable.

FIGURE 24.22 Round spermatids stained with 4’,6’-diamidino-2phenylindole (DAPI) (blue) (see color plate). Different stages of acrosome formation are shown using GFP (arrows). (From Feng et al. [217].)

In vitro culture of these cells is essential for the introduction of foreign genes in order to study spermatogenesis at the molecular level, and for preliminary studies leading to in vitro fertilization and germline gene therapy. However, two difficulties with the coculture systems described earlier in this chapter are the complex interactions between the spermatogonia and Sertoli cells and the fact that germ cell purity is of paramount importance for molecular studies. Although the cell lines meet the criteria of purity, inherent modification of their genome leading to immortalization might limit their use. Therefore, many efforts have been made recently to grow and differentiate spermatogonia in vitro by simply providing the cells with growth factors known to be essential to their behavior in vivo. Time-course analysis of spermatogonial survival in vitro was first performed with rat type A spermatogonia [48]. The cells were cultivated in minimum media (MEM), and the effects of FCS and horse serum (HS) evaluated. In MEM alone, the viability of the cells was reduced to 24% by as little as 24 hr. Addition of FCS or HS increased the viability to 50% after 24 hr. Later, our group attempted to culture porcine type A spermatogonia using different media [37]. In serumfree KSOM, 30% of the cells survived for about 1 week. Moreover, a combination of SCF and granulocyte macrophage-colony stimulating factor (GM-CSF) significantly increased the percentage of viable cells in these cultures. This report described the first attempt at cultivating isolated type A spermatogonia with growth factors in serum-free conditions. More recently, the group of D. de Rooij studied the influence of serum and a collection of growth factors on mouse prepubertal and adult type A spermatogonia [180]. The growth factors used in this study included LIF, bFGF, TGFβ, platelet-derived growth factor (PDGF), GM-CSF, tumor necrosis factor α (TNFα), SCF, and GDNF. Serum was found to stimulate the proliferation of residual somatic cells, thus changing the cellular composition of the cultures. Conversely, serum-free KSOM media containing a combination of the growth factors mentioned above could increase the viability of type A spermatogonia up to 58% by day 3 of culture. More recently, Brinster et al. combined in vitro cultures of stem cells in various conditions with the testicular transplantation technique as a functional assay [4, 5, 218]. Testicular donor cells expressing LacZ were enriched in spermatogonial stem cells and cultured for 7 days in different conditions. In particular, several cell lines treated with mitomycin C were used as feeder layers and the influence of specific growth factors was tested. At the end of the culture period, the

Chapter 24 Long-Term Cultures of Mammalian Spermatogonia

cells were transplanted into busulfan-treated recipients. The feeder layers used were of three types: fibroblast cell lines (some of them expressing isoforms of SCF), Sertoli cell lines, and bone marrow stromal cell lines. The results confirmed that spermatogonial stem cells do not respond to the action of SCF produced by fibroblast cell lines, and also that the main function of Sertoli cells is to support spermatogonial differentiation rather than stem cell self-renewal [219]. Only one of the bone marrow stroma cell lines, OP9, was able to increase the number of colony-producing spermatogonia. In addition to the growth factors tested by De Rooij et al., BPM4, activin A, and FLK2/Flt-3 ligand (FL) were tested in this study. With the exception of GDNF, none of these factors increased the number of stem cell–derived colonies in the recipient mice. On the contrary, BPM4, activin A, and FL significantly reduced the number of colony-forming germ cells. This finding indicates that the latter factors might induce differentiation in culture, thus depleting the number of stem cells available for transplantation. Although these studies defined the factors necessary for germline stem cell self-renewal and differentiation in a more controlled environment, they did not provide for a culture system in which the stem cells could self-renew for a long period of time. Shinohara et al. have now established culture conditions allowing the self-renewal of spermatogonial stem cells for at least 150 days [220]. Starting with a mixture of testis cells enriched in gonocytes, they cultured the cells in StemPro medium supplemented with estradiol, progesterone, EGF, bFGF, LIF, GDNF, and only 1% FCS. In this environment, the germ cells readily proliferated and could eventually be cultured independently of the testicular somatic cells. However, culture on mitomycin–treated mouse embryonic fibroblasts as feeder layers was still necessary. This in vitro– expanded cell population gave rise to an increasing number of colonies when transplanted into the testes of infertile, busulfan-treated mice. Another approach was taken by Hasthorpe’s group [221]. Using a cell cloning technique that they recently established for gonocytes [222], the authors singlecloned type A spermatogonia from 15-day-old mice testes. The cells were cultured in microtiter wells coated with laminin or collagen IV. The basic media contained 20% FCS and 10–4 M β-mercaptoethanol. Colonies developed that could be maintained for at least 1 week on either matrix protein (Fig. 24.23). Addition of several agents such as SCF, TGFβ, PDGF, inhibins, activin, and follistatin had a minor effect on colony formation, thus corroborating the findings of Brinster and De Rooij [180, 218]. Again, a significant inhibition of spermatogonial growth was observed

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FIGURE 24.23 Growth of a pure population of type A spermatogonia in defined culture medium. Bar = 50 μm. (From Hasthorpe [221].)

when the germ cells were cultivated on somatic cell feeder layers. This new culture system, containing pure colony-forming germ cells growing on a layer of ECM proteins, can now be used for direct study and manipulation of type A spermatogonia. Because the cells do not respond to SCF, one can conclude that they are spermatogonial stem cells, and the absence of long-term proliferation might be attributed to the fact that GDNF was not used in this study. In conclusion, optimal environments for the sustained proliferation of type A spermatogonia, including the spermatogonial stem cells, are now characterized. These defined culture systems contain combinations of growth factors that are produced by Sertoli cells in vivo. Modifications of culture conditions will, in the future, determine the role of each growth factor independently. Further, because some of these culture systems amplify a pure population of normal spermatogonia, in vitro studies that investigate the molecular mechanisms of spermatogonial proliferation and differentiation can now be properly standardized.

VI. CONCLUSION In vitro culture and expansion of type A spermatogonia, including spermatogonial stem cells, is of paramount importance for studying the molecular mechanisms that drive the first steps of spermatogenesis. The ability to manipulate germline stem cells in vitro would allow us to unravel the molecular mechanisms driving the first steps of spermatogenesis, and to characterize the signaling pathways, inducing their self-renewal versus differentiation. Transplantation techniques are now available for functional assays after stem cells manipulations. The results of such studies could help us understand the origin of certain testicular neoplasias and the causes of male infertility. In addition, the availability of stem cells that can be expanded and be manipulated in vitro opens up new areas of applications. For example, transgenic mice and rats were produced by retroviral transfections of the LacZ

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reporter gene into spermatogonial stem cells [223]. After transplantation of the manipulated stem cells into the testes of infertile mice, spermatogenesis was restored and expression of the transgene observed in the host germline. Further, the transgene was integrated in the chromosomes of the stem cells and transmitted to the progeny of the recipient. More recently, Marh and colleagues established Sertoli cell–germ cell cocultures where primary spermatocytes developed into haploid spermatid [224]. These spermatids generated normal offspring after nuclear injection into mature mouse oocytes. Thus, in the near future, diploid germ cells might be genetically altered, differentiated, and used to produce transgenic animals by in vitro fertilization. This will be particularly important for animals that do not have good ES cell lines.

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C H A P T E R

25 Transplantation INA DOBRINSKI Center for Animal Transgenesis and Germ Cell Research, New Bolton Center, School of Veterinary Medicine, University of Pennsylvania, Kennett Square, Pennsylvania

of germ cells from fertile donor mice to the testes of infertile recipient mice results in donor-derived spermatogenesis and sperm production by the recipient animal [5]. The use of donor males carrying the bacterial β-galactosidase (lacZ) gene allowed for identification of donor-derived spermatogenesis in the recipient mouse testis and established the fact that donor haplotype is passed on to the offspring by recipient animals [6]. The technique of spermatogonial transplantation in the murine system is outlined in Figure 25.1. Since its first report in 1994, germ cell transplantation has become an invaluable new tool for the elucidation of mechanisms underlying the intricate process of spermatogenesis. The increasing visibility of the technique is reflected by the publication of 20 review articles on the topic since 1998, most notably one by Ralph Brinster in Science [7]. In the years following the initial report of the technique, several important advances have been implicated, some of which are discussed in more detail later. In 1995, Jiang and Short [8] applied the technique to germ cell transplantation between rats and showed that transplanted rat testicular cells formed tubular structures (minitubules) within the seminiferous tubules rather than be integrated into the recipient seminiferous epithelium as described for the mouse. However, interpretation of these results was difficult because no unequivocal marker for donor-derived spermatogenesis was employed. Later studies demonstrated that recipient testis colonization after germ cell transplantation between rats appears to progress similarly as in the mouse [9, 10]. In 1996, production of rat sperm in mouse testes was achieved following xenogeneic spermatogonial

I. GERM CELL TRANSPLANTATION: TECHNIQUE AND FUNCTIONAL ASPECTS II. APPLICATIONS OF GERM CELL TRANSPLANTATION III. TRANSPLANTATION OF TESTIS TISSUE References

I. GERM CELL TRANSPLANTATION: TECHNIQUE AND FUNCTIONAL ASPECTS A. Development of the Technique Spermatogenesis is a continuous, highly organized process comprised of sequential steps of cell proliferation and differentiation resulting in the production of virtually unlimited numbers of spermatozoa throughout the life of the male [1]. The foundation of this system is the spermatogonial stem cell, which has the unique potential for both self-renewal and production of differentiated daughter cells that will ultimately form spermatozoa [2–4]. Among the stem cells in an animal, the spermatogonial stem cell is unique in that it is the only cell in an adult body that divides and can contribute genes to subsequent generations. Because stem cells are ultimately defined by function, unequivocal identification depends on an assay to demonstrate the potential to reconstitute the appropriate body system. This assay became available for spermatogonial stem cells when, in 1994, Dr. Ralph Brinster et al. at the University of Pennsylvania reported that transplantation SERTOLI CELL BIOLOGY Edited by M. K. Skinner and M. D. Griswold

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FIGURE 25.1 Diagram of spermatogonial transplantation in the mouse. (1) Testis tissue from a fertile transgenic donor mouse is digested into a single cell suspension. (2) The cell suspension is concentrated and injected into the seminiferous tubules of an infertile recipient mouse. (3) During the next several months, donor cells will colonize some of the recipient testis seminiferous tubules and produce spermatozoa. The location of transplanted donor cells can be demonstrated in the recipient by a stain that will stain only the donor cells blue. (4) In the most successful transplantations, the recipient male will become fertile and produce spermatozoa with genes of the donor. (5) When the donor genes have been transmitted by spermatozoa of the recipient male to progeny, these offspring contain donor genes in all their tissues and transmit these genes to subsequent generations. Insert: Recipient mouse testis stained with X-Gal. The seminiferous tubules contain donor-derived spermatogenesis. (Reprinted with permission from Brinster and Nagano [97]).

transplantation from rats to mice [11]. This astounding phenomenon provided the opportunity to answer the fundamental question of whether the cell cycle of the germ cell is inherent to the germ cell or under control of the surrounding Sertoli cells. Based on the difference in cycle length between mouse and rat spermatogenesis, it could be unequivocally shown that cycle length is under the inherent control of the germ cells [12]. Also in 1996, Brinster’s group showed that mouse spermatogonial stem cells can be cryopreserved for prolonged periods of time before transplantation and still establish spermatogenesis in the recipient testis [13]. This was a landmark discovery because it provided immediate applicability of the technique to the preservation of male genetic material. Different from cryopreservation of sperm that represent a finite quantity of male germ cells, freezing of male germline stem cells virtually

preserves the entire genetic potential of a given male because, following transplantation, these cells will continue to replicate and undergo meiotic recombination. Also in stark contrast to sperm cryopreservation, for which appropriate protocols have to be developed empirically for each species due to the highly specialized cellular architecture of the sperm, germline stem cells can be frozen successfully in diverse species using a standard cell culture freezing protocol [13–17]. In 1997, a detailed analysis of the technical aspects of germ cell transplantation in the mouse was published [18]. This paper still serves as reference material for investigators interested in adapting the technique. Importantly, the study compared three different approaches for the introduction of donor germ cells into a recipient testis (Fig. 25.2). In the initial reports, germ cells had been introduced by microinjection directly

Chapter 25 Transplantation

FIGURE 25.2

Alternatives for introduction of cell suspensions into mouse seminiferous tubules. (A) Insertion of micropipette directly into seminiferous tubules. The fluid will flow through the tubule, often reaching the rete testis and filling other tubules. (B) Insertion of micropipette into an efferent duct between the testis and the head of the epididymis. The cell suspension will flow into the rete testis and fill most or all of the tubules. (C) Insertion of micropipette into the rete testis located superficially under the tunica. From the rete, the cell suspension has access to all tubules. (Reprinted with permission from Ogawa et al. [18]).

into the recipient testis’ seminiferous tubules. Ogawa et al. [9] achieved equal success introducing cells retrograde through the rete testis or the efferent ducts, and the latter technique has now become the standard approach used by most investigators. When the technique of germ cell transplantation was first introduced, a mixture of all cell types present within the donor seminiferous tubules (all stages of germ cells, Sertoli cells, and some contaminating peritubular cells) were introduced into the recipient testes. The late Dr. Russell and coworkers transplanted testis cells into the tubules of Dominant white spotting (W) mutant mice that are inherently deficient of germ cells due to a defect in the Steel factor/c-kit signaling system. Using elegant morphological [19] and ultrastructural analyses [20, 21], these investigators characterized cellular events following introduction of donor cells into the recipient tubules. It was shown that putative stem cells make contact with recipient Sertoli cells very shortly after transplantation and reach the basement membrane of the recipient tubule within 1 week. All other cells were eliminated from the tubules by fluid movement and phagocytosis by 1 week after transplantation. Therefore, the recipient tubules serve as a filter, selectively retaining spermatogonial stem cells. This intriguing phenomenon of selective homing of

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the stem cells to their niche makes it possible to specifically study stem cell function even in situations in which a heterogeneous mixture of testis cells is transplanted. A companion study utilized xenogeneic spermatogonial transfer between rats and mice to address the question of whether donor-derived spermatogenesis is supported by the recipient Sertoli cells or whether cotransplanted donor Sertoli cells are incorporated into the recipient tubules. Based on ultrastructural differences between rat and mouse Sertoli cells, it could be shown that donor germ cells were exclusively supported by recipient Sertoli cells [22]. Subsequent studies directed at following donor cell fate after transplantation utilized cells isolated from mice carrying a lacZ (e.g., B6;129S-Gtrosa26 from The Jackson Laboratories, Bar Harbor, ME; [23]) or GFP transgene expressed in their germ cells [24]. Transplantation of genetically marked germ cells provided a unique opportunity to study the events of spermatogenesis by following individual stem cells from the moment of transplantation into a recipient testis up to complete sperm production. It was demonstrated that putative stem cells move to the basement membrane of the seminiferous tubules shortly after transplantation, where they begin to proliferate horizontally and form a monolayer network along the basement membrane for about the first month after transplantation. Beginning 1 month after transplantation, when a network of spermatogonia lining the tubule has been established, germ cells in the center of the network enter into further differentiation toward the lumen of the tubule, ultimately resulting in complete spermatogenesis [24, 25]. The temporal and spatial sequence of these steps appears to be dependent on a specific distribution and density of germ cells along the tubule. Subsequently, transplantation of marked germ cells also aided in elucidating the development of the spermatogenic stages in the testis [26]. Although much attention was initially focused on the fate of donor cells in the recipient testis, it quickly became evident that the environment of the recipient testis plays a crucial role in the dynamics and efficiency of donor cell colonization. Initial studies used mice that were inherently devoid of germ cells like the W mutant mouse. Any spermatogenesis observed in these animals after transplantation is derived from the donor cells. Alternatively, endogenous spermatogenesis in recipient animals can be depleted by irradiation [27]. Pretreatment of adult wild-type mice with the alkylating agent busulfan results in ablation of endogenous germ cells [28], a strategy now widely used in preparing recipient mice [6, 18, 29]. This approach, however, cannot be easily adapted to other species due to species-specific sensitivity to busulfan, notably its

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suppressive effect on bone marrow function and testicular health [9, 30]. Adverse effects of busulfan on testicular health can be partially overcome by treatment with estradiol and suppression of testosterone through gonadotropin-releasing hormone (GnRH) agonists [9, 29, 31]. Treatment of pregnant females with busulfan and subsequent use of the male offspring as recipients provides a promising, alternative approach [30]. Studies exploring different recipient animal models have also been directed at determining the effect of recipient age on the success of transplantation. The immature mouse testis provides a superior microenvironment for donor cell colonization compared to the adult testis [32]. The seminiferous tubules in prepubertal animals even without cytotoxic treatment contain only Sertoli cells and primitive germ cells, and the Sertoli cell tight junctions are not completely formed, facilitating the use of these animals as germ cell recipients. These properties of the prepubertal recipient testis were successfully exploited in germ cell transplantation in nonrodent species [33, 34]. Another aspect to be considered for the success of germ cell transplantation is the immunological status of the recipient. In most studies in rodents, syngeneic animals are used as donors and recipients or recipient animals are chosen from immunocompromised strains. Recent studies demonstrated that allogeneic spermatogenesis is possible in rodents, however, only when recipient animals received immunosuppressive treatment [10, 35]. In stark contrast to these reports, germ cell transplantation was successful in pigs and goats between unrelated, fully immunocompetent individuals [33, 34]. The testis is considered to be an immune privileged site, but it is unclear why transplantation between unrelated, immunocompetent animals is possible in domestic animal species but not in rodents. Nonetheless, this makes the technique infinitely more applicable in large animal species as discussed later. Most of the earlier reports on germ cell transplantation in rodents focused on qualitative aspects of recipient testis colonization. However, more quantitative analysis was a prerequisite for using this bioassay system to study treatment effects on donor cell colonization. These studies became possible when we established an image analysis approach that allows the quantification of colonization foci (colonies) and extent of colonization of recipient testes by donor stem cells [36]. As outlined earlier, the recipient testicular environment directly influences the efficiency of donor cell colonization. Studies on recolonization of testes after destruction of spermatogenesis by chemotherapy have shown that suppression of testosterone production improved spermatogenic recovery in rats [37].

Accordingly, quantitative analysis of germ cell transplantation experiments in mice demonstrated that suppression of testosterone production in recipient animals prior to and after germ cell transplantation by treatment with GnRH agonists enhanced donorderived colonization of the testes [31, 38]. The mechanism behind this observation is not entirely clear but it appears to be independent of the stem cell factor (SCF)/c-kit signaling pathway [39].

B. Bioassay for Stem Cell Potential Transplantation of germ cells into a recipient testis to monitor donor-derived spermatogenesis presently is the only unequivocal bioassay for stem cell function. After the fundamental aspects of the technique had been established, it became possible to more closely study the stem cell niche in the testis and to characterize putative spermatogonial stem cells. A summary of studies using germ cell transplantation as a bioassay for stem cell potential is outlined in Table 25.1. 1. Enrichment for Stem Cells It has been estimated that there are only about 2 × 104 stem cells in 108 cells of a mouse testis [4, 40]. To study the biology of these stem cells, it is desirable to obtain populations of cells enriched in spermatogonial stem cells from testis cell preparations. Initial work focused on potential similarities between spermatogonial stem cells and better characterized stem cell populations, mostly hematopoietic stem cells. Screening for cell surface markers potentially expressed on spermatogonial stem cells showed that selection of germ cells for expression of α6 and β1 integrin in the absence of c-kit receptor resulted in enrichment of cells capable of colonizing a recipient testis [41]. Because exposure to core body temperature results in the loss of all differentiated germ cells in a testis, it was hypothesized and subsequently demonstrated that collection of cells from experimentally induced cryptorchid testes and after induced testicular hyperthermia resulted in a significant relative enrichment for spermatogonial stem cells recovered from a mouse testis [42, 43]. Combining cell collection from cryptorchid testis with selection for surface molecules known to be present on putative stem cells, it became possible to harvest a cell population 166-fold enriched in stem cells compared to the total population of cells isolated from mouse seminiferous tubules [44]. More recently, researchers tested whether side population cells isolated from the testes were enriched in putative stem cells as described for hematopoietic stem cells [45, 46]. Side population cells are identified

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Chapter 25 Transplantation TABLE 25.1 Use of Germ Cell Transplantation to Characterize Mutants with a Phenotype of Disrupted Spermatogenesis Mutant strain (description)

Characterization of defect by reciprocal germ cell transplantation

Sl/Sld and W/Wv (defects in Steel factor–c-kit signaling system)

Germ cells from infertile Sl/Sld donor mouse can restore fertility in infertile W/Wv recipient mouse

[64]

jsd (juvenile spermatogonial depletion)

Defect causing progressive loss of spermatogenesis is inherent to germ cells

[65, 66]

ERKO (estrogen receptor α knock out)

Germ cells do not require functional estrogen receptor

[67]

Reference

ARKO (androgen receptor knock out)

Germ cells do not require functional androgen receptor

[69]

GDNF (overexpression of glial cell line–derived neurotrophic factor)

Phenotype of increased spermatogonial proliferation versus decreased differentiation is inherent to germ cells

[27]

CREM (cAMP response element modulator)

Germ cell differentiation is dependent on CREM function

[70]

As-mutant rat

Defect of germ cells and blood–testis barrier

[71]

by flow cytometry based on their ability to actively exclude a Hoechst dye. Although results were initially controversial, it appears that side population cells isolated from testicular cells are enriched in stem cells [47, 98, 99]. At the time of writing, experiments using germ cell transplantation have established that a cell population expressing α6 and β1 integrins, CD24, and Thy-1 but not Sca-1, αV integrin, c-kit, CD34, and major histocompatibility complex (MHC) class I, especially when isolated from cryptorchid testes, is likely to contain a high percentage of spermatogonial stem cells [47]. In a different approach, expression of a surface marker transgene directed by the Stra8 promoter allowed purification of a subpopulation of spermatogonia that were shown by transplantation experiments to be 700 times enriched in stem cells [48]. Another study used transgenic donor mice expressing GFP under the control of an 18-kb genomic fragment of the Oct-4 transcription factor, presumed to be expressed in undifferentiated spermatogonia from prepubertal mice to select donor cells by GFP expression, for subsequent transplantation and analysis for expression of candidate stem cell markers. Results supported the notion that putative stem cells from adult mice do not express c-kit. However, in the immature donor animal, even c-kit-positive gonocytes could colonize a recipient testis [49]. This indicates that gene expression in potential stem cells might change with testicular maturation. As mentioned earlier, young animals provide a more efficient recipient testicular environment. Accordingly, how donor age affects colonization efficiency was investigated. Neonatal mouse testes contain a higher relative concentration of stem cells than adult testes and form larger donor-derived spermatogenic

colonies after transplantation [32]. However, the absolute number of stem cells in the testis increases with age, making it more practical to collect a donor cell population from mice having undergone induced cryptorchidism as adults. Another important fundamental question raised by germ cell transplantation is whether a single stem cell can form a spermatogenic colony and what the homing or transplantation efficiency is. These questions have been addressed by experiments using serial dilution of donor cells [36] as well as serial transplantation experiments in which marked cells are collected from a recipient animal and transplanted again to subsequent recipients in a closely controlled quantitative manner. These experiments supported the “one stem cell—one colony” hypothesis and established a homing efficiency of 4–12% [50–52]. 2. Germ Cell Culture Culture of germ cells in general and putative stem cells in particular have long been elusive and it was generally accepted that stem cells could not be maintained in culture for more than a few days. To study and potentially manipulate male germline stem cells in vitro, an efficient culture system is of paramount importance. Availability of the transplantation assay system made it possible to develop and improve culture conditions for spermatogonial stem cells. Coculture with embryonic fibroblasts, L fibroblasts, or OP9 bone marrow stroma cells but not Sertoli cell lines, and addition of several growth factors known to be beneficial for other stem cell types such as embryonic stem cells and primordial germ cells [e.g., glial cell line–derived neurotrophic factor (GDNF), leukemia inhibitory factor

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(LIF), epidermal growth factor (EGF), and basic fibroblast growth factor (bFGF)], are now employed to successfully maintain mouse germline stem cells in culture for prolonged periods of time [53–55]. Stem cells isolated from newborn donors are more likely to proliferate in culture, whereas culture conditions directed at preventing germ cell differentiation appear to be most efficient in maintaining adult stem cells in vitro. A method to isolate and culture bovine type A spermatogonia has also been recently described [56]. Improving culture conditions for primary male germline stem cells is still under intense study. In the quest to understand and manipulate male germline stem cell biology, the availability of immortalized lines of germ cells would also be of immense potential. Due to the limitations of unequivocal identification, purification, and culture of the primary cells, progress in this area has been slow. Two transformed cell lines have been reported, one from rat germ cells and one from mouse type A spermatogonia, and transplantation of these cells has been used to investigate their stem cell potential [57, 58]. 3. Stem Cell Self-Renewal versus Differentiation Researchers have long wanted to know more about the molecular mechanisms governing self-renewal proliferation versus differentiation of spermatogonial stem cells. Through consecutive germ cell transplantation in Steel and W mutant mice, it was established that c-kit signaling is necessary for maintenance of germ cell differentiation but not for proliferation of type A spermatogonia [59]. GDNF has attracted special attention for its potential to maintain stem cells in an undifferentiated state. Its receptor, GFRα-1, is present on a subpopulation of spermatogonia in the mouse testis [60]. In mice overexpressing GDNF, undifferentiated spermatogonia are the predominant germ cell type in the testis [60], and transplantation of GDNFoverexpressing spermatogonia to infertile recipients maintained the phenotype, pointing to a cell-intrinsic function of GDNF in promoting proliferation over differentiation [27]. In a similar approach, transplantation of germ cells from mice ectopically expressing bcl2 in spermatogonia demonstrated that a member of the bcl2 family is involved in inhibition of spermatogonial apoptosis and growth but does not affect differentiation [61]. Collectively, these studies illustrate the utility of germ cell transplantation to increase our understanding of stem cell self-renewal and differentiation. 4. Germ Cell Transplantation in Rats Although not yet as widely established as in mice, germ cell transplantation is increasingly used in rats to

study unique aspects of spermatogenesis that are not as easily accessible in the mouse. Orwig et al. characterized functional and morphological features of rat spermatogonial stem cells [62, 63]. These studies demonstrated that gonocytes, primitive germ cells present in newborn animals and presumed to be the precursors of spermatogonial stem cells, can be divided by morphological criteria into subpopulations with different levels of stem cell potential [63]. Germ cell transplantation as a bioassay to study stem cell biology is now firmly established, and it is expected that this system will continue to further our understanding of male germline stem cells.

II. APPLICATIONS OF GERM CELL TRANSPLANTATION A. Characterization of Defects in Spermatogenesis The process of spermatogenesis is based on a complex interplay between germ cells and the supporting Sertoli cells. When a phenotype of male infertility with a defect in spermatogenesis is observed, it is often difficult to determine whether the defect affects germ cells, Sertoli cells, or both cell types. Beginning in 2000, germ cell transplantation in rodents has been increasingly employed to investigate this question. Initially, we demonstrated that transplantation of germ cells from infertile Steel mutant mice (Sertoli cell defect) to infertile W/Wv mutant mice (germ cell defect) could restore fertility [64]. This was the first report of restoration of fertility with donor cells collected from an infertile donor transplanted to an infertile recipient. Equally important, the fact that donor cells were collected from adult Steel mutant mice demonstrated that stem cells can retain their potential to support spermatogenesis for a long period of time even in the absence of adequate environmental stimuli. This finding is promising for potential applications of this approach to adult patients presented with infertility problems. In the preceding experiments, the molecular mechanisms underlying the defects in spermatogenesis were known. However, to characterize an unknown defect, standard experimental design includes reciprocal transplantation of germ cells from affected donors to wild-type testes and vice versa (Fig. 25.3). If donor cells from affected animals successfully colonize wildtype testes, the defect is localized to the somatic compartment of the testis. In that case donor cells from fertile wild-type animals will not be able to support spermatogenesis in affected recipient animals. If on the other hand affected donor cells fail to develop in

Chapter 25 Transplantation

B. Germ Cell Transplantation to Study Sertoli Cell Function

Phenotype of male infertility

Reciprocal germ cell transplantation

Donor germ cells: affected male Recipient: normal

Restoration of spermatogenesis

Yes

No

Donor germ cells: normal male Recipient: affected

Restoration of spermatogenesis

Yes

477

No

Germ cell defect

Sertoli cell defect

FIGURE 25.3 Application of germ cell transplantation to characterize defects in spermatogenesis. Reciprocal transplantation between affected animals and normal, fertile controls can determine the cellular localization of the defect. If germ cells from an affected animal can restore spermatogenesis in a normal testicular environment, but normal germ cells cannot sustain spermatogenesis in the affected animal, the defect is localized in the Sertoli cells. Inversely, if normal germ cells can restore spermatogenesis in the affected animal, but affected germ cells cannot support spermatogenesis in the normal testis, the defect is localized in the germ cells of the affected animal.

wild-type testes and wild-type germ cells can colonize affected testes, the defect is localized to the germ cells. Using this approach, it was shown that the defect associated with the juvenile spermatogonial depletion (jsd) mutation was inherent to the germ cells [65, 66], that germ cells do not require functional estrogen receptors [67, 68] or androgen receptors [69] for development, and that germ cell differentiation is regulated by GDNF and the cAMP response element modulator (CREM) function [27, 70]. In the rat, the testicular phenotype of spermatogenic arrest at the pachytene stage observed in the as-mutant was shown by germ cell transplantation studies to entail a defect in germ cells and the blood– testis barrier [71]. The genetic mutation underlying this defect has recently been elucidated [72]. A list summarizing the use of germ cell transplantation to localize spermatogenic defects to the germ cells or Sertoli cells is shown in Table 25.2. This list is continuously growing as new mouse models with male infertility phenotypes become available.

Aside from investigations to determine whether a particular defect localizes to germ cells or Sertoli cells as outlined earlier, the transplantation technique has also been applied to the study of Sertoli cell function. Xenotransplantation of rat germ cells to mouse testes established that Sertoli cells did not control the germ cell cycle but that their function appeared to be controlled by the germ cells [12]. Although not strictly speaking germ cell transplantation, a target gene can be delivered to Sertoli cells by introduction of lentiviral or adenoviral vectors directly into the seminiferous tubules. This approach was used to restore Sertoli cell function and spermatogenesis in infertile Steel mutant mice [73,74], showing that gene therapy could be targeted to Sertoli cells to restore fertility. The delicate interaction of Sertoli cells and germ cells is of fundamental importance to spermatogenesis. If this interaction is disturbed, as for example in xenotransplantation approaches or due to an inherent defect, cotransplantation of germ cells and Sertoli cells from a fertile donor might restore spermatogenesis [75]. Whereas Sertoli cells from adult testes are terminally differentiated and mitotically quiescent, Sertoli cells isolated from perinatal mouse testes are still rapidly dividing and can form tubular structures (minitubules) within the recipient testes that in some cases can support spermatogenesis. However, to make the approach of Sertoli cell transplantation more efficient, the ablation of endogenous Sertoli cells by microinjection of cadmium into the seminiferous tubules is necessary, making its practical application difficult.

C. Xenotransplantation of Germ Cells In 1996, production of rat sperm in mouse testes was achieved following xenogeneic spermatogonial transplantation from rats to mice. Some endogenous spermatogenesis recovered in the immunodeficient mouse host leading to the production of both rat and mouse sperm in the same testis (Fig. 25.4) [11]. This exciting observation opened up the possibility that spermatogenesis from a wide range of donor species could be achieved in the mouse host. For its obvious practical potential, cross-species germ cell transplantation was explored further. Rat and mouse are phylogenetically distant by about 11 million years. The next area of investigation involved whether xenogeneic spermatogenesis in mouse testis could occur from a phylogenetically more distant rodent donor species. Hamster and mouse are separated by 15 million years, a phylogenetic

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TABLE 25.2 Germ Cell Transplantation as Bioassay for Stem Cell Potential Topic investigated

Main findings

References

Pattern of colonization after transplantation

Proliferation of spermatogonia along the basement membrane precedes initiation of germ cell differentiation in a defined pattern

[19, 24–26]

Cell cycle control

The germ cell, not the Sertoli cell, determines the length of the cell cycle

[12]

Germ cell culture

Mouse spermatogonial stem cells can survive in culture for prolonged periods of time

[53–55]

Endocrine effects on donor cell colonization

Suppression of testosterone supports colonization and supports proliferation of germ cells in the jsd mutant mouse model

[31, 38, 92, 93]

Characterization of stem cells

Germ cells expressing α6- integrin, CD24, Thy1, but not αV-integrin, c-kit, MHC-I, Sca1, CD34 have stem cell potential

[41, 47]

Enrichment of stem cells

Induced cryptorchidism, testicular hyperthermia, and selection for surface markers result in a germ cell population enriched for stem cells

[42–44]

Characterization of a stem cell niche in the testis

SCF/c-kit signaling is necessary for spermatogonial differentiation but not proliferation; The immature testis provides a superior environment for colonization with donor stem cells; Distribution of spermatogonia in the mouse is not random; Donor cells can compete with endogenous stem cells for the stem cell niche

[59] [32, 94] [95] [94]

Rat (SV40 transformation); Mouse (telomerase overexpression)

[57] [58]

Cell lines

FIGURE 25.4 Xenogeneic rat spermatogenesis in mouse testes: (A, B) normal mouse sperm and ( C, D) normal rat sperm. Note the characteristic difference in head shape and length of the sperm tail between mouse and rat spermatozoa. (E, F) Epididymal sperm recovered from a recipient mouse that had received transplantation of rat germ cells 110 days previously. One in 39 sperm produced by this mouse showed the distinct morphology of rat sperm. (Reprinted with permission from Clouthier et al. [11]).

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Chapter 25 Transplantation

distance similar to that between cattle and goats and three times the 5 million years that separate human and chimpanzee. Hamster spermatogenesis occurred successfully in the mouse host [76], although with lower efficiency than xenogeneic rat spermatogenesis. However, further experiments showed that with increasing phylogenetic distance between donor and recipient species, complete spermatogenesis could no longer be achieved in the mouse testis. Transplantation of germ cells from nonrodent donors ranging from rabbits (about 60 million years distant to mice) and dogs to pigs, bulls, horses, and ultimately nonhuman primates and humans resulted in colonization of the mouse testis, but spermatogenesis became arrested at the stage of spermatogonial expansion [14–17]. A report of human spermatogenesis occurring in mouse or rat testes has so far not been substantiated. The experiments reported to date are summarized in Table 25.3. Although the initial steps of germ cell recognition by the Sertoli cells, localization to the basement membrane and initiation of spermatogonial proliferation, appear to be conserved between such diverse species, it was hypothesized that with increasing phylogenetic distance between donor and recipient species the recipient testicular environment (comprised of Sertoli cells and paracrine factors) becomes unable to support spermatogenic differentiation and meiosis. This incompatibility of donor germ cells and recipient testicular environment could be overcome by cotransplantation of germ cells and Sertoli cells as mentioned earlier or by testis tissue transplantation as described later. Although xenogeneic spermatogonial transplantation did not have the envisioned immediate practical application, it nonetheless provides a bioassay for stem cell potential of germ

cells isolated from other species [14, 15, 77] and an intriguing tool for the study of the cellular and noncellular requirements of spermatogenesis in diverse species.

D. Germ Cell Transplantation in Species Other Than Rodents Although most germ cell transplantation experiments utilize rodent models, germ cell transplantation technology has recently been applied to nonrodent species including pigs, goats, and rhesus monkeys [33, 34, 78, 79, 100] (Table 25.4). The rationale behind this work is twofold. In nonhuman primates, in humans, and perhaps also in other large animal species, germ cell transplantation could serve to restore male fertility after an insult to the testis. Specifically, one could preserve germ cells prior to irradiation or chemotherapy treatment for cancer because these treatments often lead to temporary or permanent destruction of spermatogenesis. Reintroduction of autologous germ cells could then restore fertility in the patient once the illness has been overcome. A report in the monkey demonstrated this approach in principle [79, 100], and its application in humans has been discussed [80, 81], but no definitive data have been reported. Reintroduction of donor cells collected before treatment carries the risk that cancerous cells could also be reintroduced into the patient as demonstrated in leukemic rats [82]. Therefore, the safety of this approach for human applications is questionable at this time. Cell sorting technology could potentially be employed to overcome this risk. Germ cell transplantation has an advantage over the only approach currently available, cryopreservation of sperm prior to treatment, in that it could be

TABLE 25.3 Xenotransplantation of Germ Cells Donor species Rat

Recipient species Mouse

Donor-derived spermatogenesis Complete

Reference [11, 22]

Hamster

Mouse

Complete

[76]

Rabbit, dog

Mouse

Spermatogonial proliferation but no further differentiation

[14]

Bull

Mouse

Spermatogonial proliferation but no further differentiation

[15, 77]

Pig, horse

Mouse

Spermatogonial proliferation but no further differentiation

[15]

Rhesus monkey

Mouse

Spermatogonial proliferation but no further differentiation

[16]

Human

Mouse

No colonization; Spermatogonial proliferation but no further differentiation

[96] [17]

Mouse

Rat

Complete

[9]

Mouse

Pig

No colonization

[33]

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TABLE 25.4 Germ Cell Transplantation in Species Other Than Rodents Species

Donor-derived spermatogenesis

Transmission of donor haplotype to offspring

References

Rat (homologous)

Complete

No Not tested Yes

[8] [9] [10]

Monkey (autologous)

Complete

Not tested

[79, 100]

Pig (homologous)

Complete

Not tested

[33]

Goat (homologous)

Complete

Yes

[34]

applied to prepubertal patients where sperm cannot be obtained or to adult patients rendered azoospermic or teratozoospermic by disease. The technique is of great interest in domestic or endangered animals for its potential to preserve genetic material from immature males that is lost before they reach puberty such that preservation of sperm is not an option. It has also been discussed as an agricultural tool to store genetic material from castrated male pigs (barrows) used in performance testing in lieu of using littermates of the top-quality animals for breeding. Recently, potential applications in cattle have been postulated in which the high-quality genetics of Bos taurus bulls could be delivered to the production herd via Bos indicus recipient animals that are more adapted to certain environments. Another important potential application of the technique would be transgenesis through the male germ line. Application of germ cell transplantation technology to nonrodent species has been difficult. Due to differences in testicular anatomy and physiology, germ cells cannot be delivered by the same technique as in rodents. Instead, we developed a technique combining ultrasound-guided cannulation of the centrally located rete testis with delivery of germ cells by gravity flow [33, 78]. With this technique, we succeeded in transplanting donor cells from transgenic donor goats into the testes of immunocompetent, prepubertal recipient animals. Once these goats became sexually mature, they produced sperm carrying the donor haplotype and transmitted the donor genetic makeup to the offspring. This provided proof-of-principle that germ cell transplantation results in donor-derived sperm production and fertility in a large animal species [34]. Importantly, donors and recipients were unrelated and immunocompetent. Transplantation between unrelated immunocompetent animals has not been achieved in rodents. Future experiments are now directed at improving the efficiency of germ cell transplantation in domestic animals by enrichment for stem cells and suppression of recipient spermatogenesis as discussed earlier.

E. Germ Cell Transplantation and Transgenesis In recent years, transplantation of transfected germ cells has been investigated as an alternative means to generate transgenic animals through the manipulation of the male germline. Although this is potentially a very powerful approach, several factors make it an exceedingly difficult task. Efficient, targeted introduction of a gene of interest into male germline stem cells before transplantation requires high numbers of relatively pure starting populations of germline stem cells and optimized culture systems that allow the selection and expansion of transgenic cells before transplantation. Progress is being made in the areas of enrichment of cells for putative stem cells and stem cell culture as outlined earlier. However, the low proliferating activity of stem cells has been an additional major obstacle, and, therefore, progress toward achieving the goal of efficient transgenesis through the male germline has been slow. Because putative stem cells are considered to be replicating very slowly or not at all when cultured, viral gene therapy vectors that integrate into nonreplicating cells are an obvious choice for gene transfer. Some success has been reported in generating transgenic mice and rats by retroviral or lentiviral transduction of germ cells prior to transplantation [83–87]. In one report, the transgene was stably integrated and passed on to subsequent generations without gene silencing [84]. However, for the production of transgeneic domestic animals destined to produce pharmaceutical proteins in their milk, approaches using nonviral transfection are desirable and currently under investigation. The practical applicability of transgenesis through germ cell transplantation is ultimately dependent on improvements in efficiency of stem cell selection, transfection, and culture for each target species. Efficient culture systems that allow maintenance and expansion of stem cells are of utmost importance because this would allow for gene targeting approaches and cell selection prior to transplantation.

Chapter 25 Transplantation

Transgenesis through the male germline using transplantation of transfected germ cells has tremendous potential in species like the rat and domestic animals where embryonic stem cell technology is not available. Current options to generate transgenic animals other than mice include pronuclear microinjection of DNA and nuclear transfer technology using transgenic donor nuclei, as well as several other more isolated approaches, such as sperm-mediated DNA transfer and intracytoplasmic co-injection of sperm and DNA. However, all currently available technology is fraught with low efficiency and developmental abnormalities in the few resulting offspring, making the approach of using germ cell transplantation a potentially very valuable alternative.

III. TRANSPLANTATION OF TESTIS TISSUE Many complex aspects of testis function in humans and large animals have remained elusive due to the lack of suitable in vitro or in vivo models. Supporting spermatogenesis in an in vitro system has so far remained fairly inefficient. As outlined earlier, transplantation of isolated male germ cells represents a very powerful approach for the study of spermatogenesis as well as for preservation and manipulation of the male germline. However, the technique cannot be easily adapted between diverse species. Xenogeneic germ cell transplantation did not result in complete sperm production in species other than rodents, probably due to an incompatibility between donor germ cells and recipient Sertoli cells and testicular environment. Cotransplantation of the donor germ cells with their surrounding testicular tissue into a mouse host would preserve the testicular integrity and still allow experimentation in a small rodent. Therefore, we developed ectopic xenografting of testicular tissue under the back skin of immunodeficient mice as an alternative approach for the maintenance and propagation of male germ cells that can be more readily applied to different mammalian species. Testis tissue xenografting maintains the structural integrity of the testicular tissue and provides the accessibility essential for the study and manipulation of testis function as well as for male germline preservation. Through this approach, we showed that xenografting of testis tissue from newborn pigs and goats resulted in production of normal, functional sperm in a mouse host (Fig. 25.5) [88]. This was the first report of complete, functional xenogeneic spermatogenesis in species other than rodents, and it also represented the first time that sperm could be obtained from neonatal donors.

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Subsequently, ectopic xenografting of testicular tissue was also reported from hamsters or marmoset monkeys to mice [89]. Sperm recovered from allografts (mouse) and xenografts (pig, goat) supported pronuclear formation when injected into mouse oocytes. Mouse sperm recovered from allografts resulted in normal, fertile progeny when injected into mouse oocytes followed by embryo transfer [90]. Mouse and rabbit sperm recovered from testis grafts developing in the testis of recipient mice also resulted in the birth of live pups when used for ICSI into homologous oocytes [91]. Pig sperm recovered from ectopic xenografts were used for ICSI into pig oocytes and supported development to the blastocyst stage. Embryo transfer experiments will have to be performed to provide proof that fertilization using xenogeneic sperm can also result in normal development of offspring. The onset of spermatogenesis in xenografted pig testis tissue occurred slightly earlier than in the donor species [88], making it likely that sperm production through xenografting can be significantly accelerated in slow-maturing species. We now have evidence in rhesus macaques that testicular maturation and sperm production can indeed be accelerated by xenografting of testis tissue [101]. The onset of puberty appears to be limited by the lack of adequate gonadotrophic support in the immature animal, not by immaturity of the gonadal tissue. Xenografting into a castrated mouse host initially provides high levels of FSH as the required stimulus for testicular development. Testis tissue grafting presents a unique opportunity to study the effects of exogenous factors on spermatogenesis in an in vivo culture system. Additional manipulation of the endocrine milieu in the host could significantly shorten the time span required for sperm production from genetically superior males. On the female side, shortening of the generation interval to accelerate genetic progress has been achieved by in vitro fertilization (IVF) of oocytes recovered from sexually immature calves. To date, however, significant acceleration of sperm production has not been possible in male farm animals. Similar to isolated germ cells, testicular tissue can be stored frozen prior to grafting and with retention of its developmental potential [88, 89]. Cryopreserved testis tissue has two major advantages: It contains spermatogonial stem cells with an unlimited potential for self-renewal, providing a potentially inexhaustible source of male germ cells after grafting, and it makes xenografting of testis tissue applicable to clinical and field situations where transplantation has to be performed at a later date. Due to their small size, testis tissue grafts appear to be accessible to viral and nonviral vectors for the introduction of genetic material [102]. Transfection of germ

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FIGURE 25.5 Ectopic xenografting of testis tissue from newborn piglets to nude mice. (A) Small pieces of pig testis tissue (0.5–1 mm in diameter) were grafted under the back skin of nude mice. (B) The grafts had expanded to 4–8 mm in diameter by 10 weeks after grafting. (C) A porcine sperm recovered from a xenograft 27 weeks after grafting. Bars = 5 mm (A, B) or 20 μm (C). (Reprinted with permission from Honaramooz et al. [88]).

cells prior to grafting provides a unique tool to study the genetic regulation of spermatogenesis since it can be used to induce or disrupt specific genes followed by close observation of spermatogenesis without the necessity of performing experiments in the target species. In addition, once a gene of interest is introduced into the germ cell of immature donor testis tissue, subsequent sperm production after xenografting will result in the production of transgenic sperm that can be used for in vitro embryo production. Transduction of germ cells in testis tissue prior to xenografting might represent an alternative approach for the generation of transgenic sperm that is more easily applicable to different domestic animal species and would require fewer large animals. Ectopic testis tissue grafting represents a new option for male germline preservation. Because it provides a potentially inexhaustible source of male gametes even from immature gonads, grafting of fresh or preserved testis tissue offers an invaluable tool for the conservation of endangered species or valuable livestock by allowing sperm production from immature males. Unlike autologous transplantation of isolated

germ cells to restore fertility in a patient following cancer therapy, xenologous grafting and use of the resultant sperm for assisted fertilization will eliminate the potential risk of tumor cell transmission. The accessibility of the tissue in the mouse host makes it possible to manipulate spermatogenesis and steroidogenesis in a controlled manner that is not feasible in the donor animal and certainly not in humans. This in turn will allow analysis of the effects of toxicants and potential male contraceptives on testis function in the target species. Finally, grafting of testis tissue from experimental animal strains will provide a previously unavailable tool to study germ cell development and even to produce gametes from animals with poor viability, such as neonatal lethal transgenic, mutant, or cloned animals.

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Index

A Aaligned (Aal), 306 Apaired (Apr), 306, 308 Astem (As), 306–308 ABP, 97, 208 Activin, 229–230. See also Inhibins and activins. ActRI, 228, 229t ActRII, 228, 229t ADAM-TS family, 124t, 127–128 ADAM-TS4, 124t, 128 Adrenoleukodystrophy (ALD), 391–392 Affymetrix, 104 Affymetrix Rat U34A GeneChip, 98 Aggregation of Sertoli cells, 53 Aging, 33 Ague, Jacqueline, 271 AKT, 307 Ala189Val, 158 Ala419Thr, 158 ALCAM, 425 ALD, 391–392 Alkylphenols, 332 Allan, Charles M., 171 α-inhibin, 390 α-macroglobulins, 141 α-Tubulin, 31f, 32f α2-antiplasmin, 130 α2-macroglobulin (α2-MG), 111, 127t, 130, 141–142, 164–165, 239 Altered germ cell attachment, 346–347 ALV, 429 AMH. See Müllerian inhibiting substance (MIS). Amphibians. See Fishes and amphibians. Amyloid, 187 Anamniotes. See Fishes and amphibians. Androgen-binding protein (ABP), 208 Androgen receptor (AR), 261–262, 438 Androgen regulation, 199–216 ABP, 208 AR expression, 200–202, 219 AR mutations/knockouts, 205–207 future prospects, 211–213 genomic vs. nongenomic effects, 211 germ cell transplantation, 209–211 Pem, 207–208 protein synthesis/secretion, 209 recovery of spermatogenesis, 211 SCARKO mouse, 205–207 Sertoli cell proliferation, 203 spermatogenesis, 203–207 spermatogonial development, 209–211 STF, 208–209 Androgens, 61 ANKYRIN, 266f

Ankyrin, 266 Anti-Müllerian hormone (AMH). See Müllerian inhibiting substance (MIS). AP-1 transcription factor complex, 258 Apical cytoskeletal support, 347 Apical sloughing, 349–350 Apoptosis, 350–351 AR, 261–262, 438. See also Androgen regulation. AR expression, 200–202, 219 Architis, 399–401 Arg273Cys, 158 ARKO, 475t Armadillo, 312 As-mutant rat, 475t ASC-17D cells, 330t, 336–337 Asp224Val, 158 Aspartylyglycosuria, 187 Atanassova, Nina, 213 Authors. See Contributors. Avian leukosis virus (ALV), 429 Axl, 17t AZF, 387 AZF deletions, 387 Azoospermia, 403 Azoospermia factor (AZF), 387

B Bambi, 228t, 239 Bartke, Andrzej, 81 Basic fibroblast growth factor. See bFGF. Basic helix–loop–helix (bHLH) transcription factors, 253t, 254t, 259–260 Basic leucine zipper (bZIP) transcription factors, 253t, 254t, 256–259 Bclw, 17t BDNF, 111 Benomyl, 355 β1 integrin, 420 Betaglycan, 228t, 235, 238 Beta scaffold factors with minor groove contacts, 253t, 255t, 266–268 bFGF, 110, 322 bHLH transcription factors, 253t, 254t, 259–260 Bilateral testicular tumors, 187 Bioassay of the stem cell potential, 474–476 Biological process ontology, 99–100, 100f Black bear, 82. See also Seasonal breeders. Blood-testis barrier, 32–33 Blood-testis barrier defect, 347 BMP, 233–234, 308 BMP4, 233, 308–309, 419 BMP7, 234 BMP8A, 234 BMP8b, 234, 308, 419

487

BMPRI, 228, 229t BMPRII, 228, 229t Boekelheide, Kim, 345 Bolden-Tiller, Olga U., 437 Bone morphogenetic proteins (BMPs), 233–234, 308 Bott, Rebecca, 62 Bouma, Jerry, 71 Brain derived neurotrophic factor (BDNF), 111 Brinster, Ralph, 471 Brown, Terry, 145 Brown marsupial mouse, 81. See also Seasonal breeders. Bunsen burner, 4 bZIP-bHLH family of transcription f actors, 260 bZIP transcription factors, 253t, 254t, 256–259

C C. elegans, 304–305, 309–311 C/EBP transcription factors, 256–257 c-jun, 258 C-kit, 165, 309, 444. See also SCF/c-kit system. C-kit/PI3-K signaling, 307f c-myc, 260 C18-4 cell line, 459–460 CAH, 187, 392–393 CAM, 204 cAMP, 159, 292 cAMP-evoked events, 159 Cancer, 427–429 Cancer Genome Anatomy Project database, 145 CANNTG, 270 Carbendazim, 31, 355–357 Carney’s complex (CNC), 389, 390 Cathepsin, 124t, 133–144 biological functions, 134 biosynthesis, 133–134 cathepsin A, 124t, 139–140 cathepsin B, 124t, 138 cathepsin D, 124t, 140 cathepsin K, 124t, 138 cathepsin L, 124t, 135–138, 142–144 cathepsin S, 124t, 138–139 cathepsin V, 124t inhibitors, 140–142 CBP, 268 CCAAT/enhancer protein (C/EBP) transcription factors, 256–257 Cdk5, 333 CDKIs, 222 Cdks, 222 Cell adhesion molecule (CAM), 204 Cell biology of testis, 317, 318f

488 Cell-cell interactions, 301–342 cell lines, 329–342 Sertoli cell-somatic cell interactions, 317–328 spermatogonial stem cell fate, 303–315 Cell lines. See Sertoli cell lines. Cellular component—cell ontology, 100–101, 100f Centrifugal elutriation, 451 cfos, 258 Changes in distribution, quantity, or biochemical properties, 352–353 Charron, Martin, 121 Chaudhary, Jaideep, 251 Chemotherapy, 403–404 Chern, Jeannie, 145 ChIP on a chip, 271 Chromosome distribution of Sertoli cell expressed genes, 103–104 Ciliary neurotrophic factor (CNTF), 113, 421, 426 Cip/Kip family of CDKIs, 222 CIS cell, 430 Cisplatin, 367–368 Claudin-1, 208 Claudin 11, 125 Claudins, 208 Clinical conditions. See Conditions affecting Sertoli cells. Cloning FSHR, 283 Cloud, Joseph G., 71 Clusterin, 17t CNC, 389, 390 CNTF, 113, 421, 426 Co-Smads, 228, 229t Coelomic epithelium, 51 Colchicine, 368 Collagen IV, 127t Common α mutations, 157 Common α-subunit, 155 Computer software (gene array analysis software), 99 Conditions affecting Sertoli cells, 383–413 ALD, 391–392 architis, 399–401 CAH, 392–393 chemotherapy, 403–404 cryptorchidism, 397–398 diabetes, 401–403 disorders of sexual differentiation, 384–386 fragile X syndrome, 390–391 FSH gene and receptor mutations, 388, 389t hypothyroidism, 397–398 Klinefelter’s syndrome, 393–395 markers of Sertoli cell function, 383–384 McCune Albright syndrome, 395 MGD, 384 PMDS, 384–386 radiation, 403–404 SCOS, 388–389 Sertoli cell neoplasia, 389–390 TDS, 386–387 testicular torsion, 401 TH, 384 variocele, 398–399 vasectomy, 401 Y-chromosome deletions, 387 Congenital adrenal hyperplasia (CAH), 187, 392–393 Connexin 43 (Cx43), 224, 423–424 Contributors Allan, Charles M., 171 Bartke, Andrzej, 81 Boekelheide, Kim, 345

Index Bolden-Tiller, Olga U., 437 Bouma, Jerry, 71 Charron, Martin, 121 Chaudhary, Jaideep, 251 Cloud, Joseph G., 71 Cooke, Paul S., 217 Cupp, Andrea S., 43 de Miguel, Maria P., 417 Dobrinski, Ina, 471 Donovan, Peter J., 417 Dym, Martin, 303, 449 Feng, Lixin, 303 França, Luiz R., 3, 19, 217 Griswold, Michael D., 15, 95 Handelsman, David J., 171 Heckert, Leslie L., 281 Hess, Rex A., 3, 19 Hikim, Amiya P. Sinha, 81 Hofmann, Marie-Claude C., 449 Holsberger, Denise R., 217 Huhtaniemi, Ilpo, 155 Johnson, Kamin J., 345 Loveland, Kate L., 227 McLean, Derek, 95 Meistrich, Marvin L., 437 Porter, Karen L., 437 Richburg, John H., 345 Roberts, Kenneth P., 329 Robertson, David M., 227 Salameh, Wael A., 383 Sharpe, Richard M., 199 Shetty, Gunapala, 437 Sinha Hikim, Amiya P., 81 Skinner, Michael K., 43, 107, 251, 317 Swerdloff, Ronald S., 383 Toppari, Jorma, 155 Wright, William W., 121 Cooke, Paul S., 217 COUP-TF, 295 Cre, 159, 292 CREB, 159, 257 CREL, 266f CREM, 159, 257–258, 475t CRES3, 141 Crim1, 228t, 239 Cryopreserved testis tissue, 481 Cryptorchidism, 34, 397–398 Cultures. See Long-term cultures and mammalian spermatogonia. Cupp, Andrea S., 43 Cx43, 224, 423–424 CXCR4, 420 Cyclic protein 2, 17t Cyclin-dependent kinase inhibitors (CDKIs), 222 Cyclin-dependent kinases (Cdks), 222 Cyclins, 222 Cys2 His2 zinc-finger domains, 255t, 264–265 Cys4 zinc fingers, 264 Cystatin C, 141 Cystatin SC, 141 Cystatin superfamily, 140–141 Cystatin TE-1, 141 Cystatins, 140–141 Cytokines, 111 Cytoplasm, 26–32 cytoskeleton, 28–32 endoplasmic reticulum, 27–28 Golgi, 28 liposomes/multivesicular bodies, 28 mitochondria, 26–27 Cytoskeleton, 28–32

D D. melanogaster, 305, 310–311 DAX-1, 49, 263 Dazl, 459–460 DBCP, 210 de Gendt, Karel, 213 De Miguel, Maria P., 417 Decreased seminiferous tubule fluid secretion, 351–352 Deer, 182–183 Defects in spermatogenesis, 476–477 DEHP, 218, 363, 430 Delta 1, 309–310 Desert hedgehog, 17t Development of functional testis, 58–62 gonadotropins, 61–62 Leydig cell differentiation, 59 NGF family, 60–61 PDGF, 61 Sertoli cell proliferation, 59 steroid hormones end receptors, 61 TGFα/EGF, 60 TGFβ family, 60 thyroid hormone, 61 Development of bipotential gonad, 47–50 Dhh, 57, 320 Dhh/Ptch-1 signaling, 424 Di-(2-ethylhexyl)phthalate (DEHP), 218, 363, 430 Diabetes, 401–403 Dibromochloropropane (DBCP), 210 Diethylhexaphthalate (DEHP), 218 Dinoseb, 361 Disorders/diseases. See Conditions affecting Sertoli cells. Distal tip cell (DTC), 304, 310–311 DM genes, 49 Dmrt genes, 49 Dmrt1, 49, 268–269 dmrt1Y, 72 dmY, 72 Dobrinski, Ina, 471 Donovan, Peter J., 417 DPP class (BMP), 233 Drosophila, 305, 310–311 DTC, 304, 310–311 Dym, Martin, 303, 449 Dysfunction. See Conditions affecting Sertoli cells; Toxicants.

E E2, 75 E-T switch, 188–189 E2-2, 259 E2f, 289–290 E12, 259 E47, 259 E-box consensus site (CANNTG), 270 ovine, 295 rat FSHR promoter, 287–288 EC cells, 427 Ectopic secretion of hCG, 187 Ectoplasmic specialization, 28–30, 122 EDS, 203, 204 EGF, 60, 108t, 109 EGFR, 311 Embryonic Sertoli cell differentiation, 43–70 overview, 43–44 Sry expression, 48, 49t, 57

Index stage 1 (genital ridge development), 44–47 stage 2 (development of bipotential gonad), 47–50 stage 3 (sex determination), 50–52 stage 4 (seminiferous cord formation), 52–58. See also Seminiferous cord formation. stage 5 (development of functional testes), 58–62. See also Development of functional testes. Emx2, 46–47 Endoglin, 235 Endoplasmic reticulum, 27–28 Endothelial cell, 323 Endothelial cell migration, 58 Enrichment for stem cells, 474–475 Environmental cell-cell interactions defined, 318, 318t Sertoli cell-Leydig cell interactions, 319 Epidermal growth factor (EGF), 60, 108t, 109 Epidermal growth factor receptor (EGFR), 311 ERα, 262, 438 ERβ, 439 ERβ1, 262 ERβ2, 262 ERKO, 262, 475t Erythropoietin, 113 ES cell lines, 330t, 338 eSRS21, 75 Estradiol-to-testosterone (E-T) switch, 188–189 Estrogen, 56, 61, 113 Estrogen receptors, 56, 262, 438–439 E-T switch, 188–189 Ethane dimethane sulfonate (EDS), 203–204 Ethylene glycol monomethyl ether, 369 Eukaryotic promoter, 252f. See also Promoter. Evans, Herbert, 171 Experimental approaches to studying dysfunction, 354–355 Extracellular matrix proteins, 47

F FACS, 452–453 Family 2 cystatins, 140–141 Fas system, 362–363 Fas ligand (FasL), 332, 362–363, 445 Fas-FasL system, 445 FAST-1, 228 FBF-1, 311 FBF-2, 311 Feng, Lixin, 303 FGF, 108t, 110 FGF-9, 55 FGF R1, 127t Fibrinase, 124t Fibrinolysin, 124t Fibroblast growth factor (FGF), 108t, 110 Fibroblast growth factor 9 (FGF-9), 55 Fibronectin, 127t 15P-1 cells, 330t, 334–335 Fishes and amphibians, 71–79 concluding remarks, 77 meiosis, 76–77 Sertoli cell origin, 72–74 Sertoli cell proliferation, 75 spermatogenesis, 74–75 testicular recrudescence, 73–74 testis structure, 72 5-hydroxybenzimidazole, 355

Fluorescence-activated cell sorting (FACS), 452–453 FMRP, 391 FOG-2, 52, 264 Follicle-stimulating hormone. See FSH. Follistatin, 228t, 239 Forskolin, 57 42GPA-9 cells, 330t, 335–336 45T-1 cells, 330t, 335 Fragile X mental retardation protein (FMRP), 391 Fragile X syndrome, 187, 390–391 França, Luiz R., 3, 19, 217 FS288, 239 FS315, 239 FSH, 155, 440–441 estrogen, 113 fishes and amphibians, 74, 76 functional testis, 61–62 historical overview, 171–172 in vivo actions, 171–197. See also In vivo FSH actions. inhibin/activin, 112, 230–231 MIS, 113 modification of action by spermatogenic cells, 163–165 protein subunits, 155–159 seasonal breeders, 83, 84 FSH action in vivo. See In vivo FSH actions. FSH binding, 163–164 FSH deficiency, 185–187 FSH promoter, 97 FSH-R3, 161 FSH receptor, 17t. See FSHR. FSH receptor gene, 97 FSH-secreting pituitary macroadenomas, 187 FSH-stimulated cAMP production, 164 FSH-stimulated protein synthesis, 164–165 FSHβ, 155 FSHβ mutations, 157 FSHR, 157–158, 281–299, 438 activation, 159 alternate splicing of message, 161–162 biology, 281–283 cloning, 283 constitutive activity, 159–161 desensitization/downregulation, 162–163 evolution, 283–284 genome analysis advancements, 284 mutations, 158–159 promoter, 284–295. See also FSHR promoter. signal transduction, 159 FSHR promoter, 284–295 general features, 284–287 human promoter, 294 murine promoter, 294 ovine promoter, 294–295 rat promoter, 287–294 Future directions, 17–18 FXR2P, 391 FXRIP, 391 Fyn tyrosine kinase, 17t

G Gs, 159 Gs?-GTP, 159 Galbo, Jeanne, 371 γ-Diketones, 357–361 γ-Tubulin, 360 GATA family of transcription factors, 52, 239–241, 264

489 GATA-1, 17t, 264 GATA-4, 52, 264 GATA-4 knockin allele (GATA-4ki), 52 GATA-4 null mutants, 52 GATA-6, 264 Gβ, 159 GC-1spg, 457–458 GC-2spd, 458 GC-3spd, 458 GC-4spc, 458 GC-5spg, 458 GC-6spg, 458 GC-box binding transcription factors, 264–265 GDNF. See Glial cell line-derived neurotrophic factor (GDNF). Gelatinase A, 124t Gelatinase B, 124t Gene array analysis, 97–99 Gene array analysis software, 99 Gene expression, 96. See also Sertoli cell gene expression and protein secretion. Gene expression (mammalian germline), 429–430 Gene Expression Omnibus (GEO), 104 Gene grouping and family nomenclature, 145 Gene ontology analysis, 99–103 Gene Ontology (GO) Consortium, 99 Genes high expressed by/unique to Sertoli cells, 96–97 GeneSifter, 99 GeneSpring 5.0, 99 Genital ridge, 43 Genital ridge development, 44–47 Genome analysis advancements, 284 Genomes, 104 Germ cell apoptosis, 351 Germ cell culture, 475–476 Germ cell-Sertoli cell interactions, 48 Germ cell transplantation, 16, 354, 471–485 applications, 476–481 bioassay of the stem cell potential, 474–476 defects in spermatogenesis, 476–477 development of the technique, 471–474 enrichment for stem cells, 474–475 germ cell culture, 475–476 one stem cell - one colony hypothesis, 475 rats, 476 species other than rodents, 479–480 stem cell self-renewal vs. differentiation, 476 studying Sertoli cell function, 477 transgenesis, 480–481 xenotransplantation of germ cells, 477–479 Germ line stem cells, 311–312 GFRα-1, 237, 444, 452 GFRα-2, 237 Glial cell line-derived neurotrophic factor (GDNF), 112, 307–308, 444 function in testis, 237–238 physiological action, 108t receptor, 108t size, 108t spermatogonial cell lines, 460 structure/synthesis, 236–237 transplantation, 460, 475t Glp-1, 310–311 Glutaraldehyde, 21 Glycogen, 34 Golden (Syrian) hamster, 82, 85. See also Seasonal breeders. Golgi, Camillo, 3 Golgi apparatus, 28

490

Index

Gonad formation. See Embryonic Sertoli cell differentiation. Gonadotropin. See FSH; LH. Gonocyte development and differentiation, 417–435 formation of PGCs, 419–422 gene expression (mammalian germline), 429–430 from gonocyte to spermatogonia, 425–427 history of germline, 418–419 from PGCs to gonocyte, 422–425 testicular cancer, 427–429 Greep, Roy, 171 Griffin, Jill, 62 Griswold, Michael D., 3, 15, 95, 254–255, 271 Growth factors, 108–112 cytokines, 111 defined, 107 FGF, 110 GDNF, 112. See also Glial cell line-derived neurotrophic factor (GDNF). IGFs, 108–109 KL/SCF, 111–112 neurotropins, 110–111 TGFα, 109 TGFβ, 109–110

H H-kininogens, 140 HA, 333 Hamster, 82, 85, 89, 181–182. See also Seasonal breeders. Handelsman, David J., 171 Hasthorpe’s group, 463 hCG, 185–187 HEB, 259 Heckert, Leslie L., 281 Helix–turn–helix motif, 253t, 255t, 265–266 Hepatocyte growth factor (HGF), 55 Hermaphroditism, 384 Hess, Rex A., 3, 19 HFH-11, 266 HGF, 55 High-mobility group (HMG) box, 48 High mobility group (HMG) proteins, 267–268 Hikim, Amiya P. Sinha, 81 Historical overview, 3–13, 19 FSH actions, 171–172 germ cell transplants, 471–474 germline, 418–419 Sertoli. See Sertoli, Enrico. timeline, 8t transition to modern Sertoli cell, 10 HMG1, 267 Hofmann, Marie-Claude C., 308, 449 Holsberger, Denise R., 217 Homeobox genes, 265 Hormones, 107, 112–113 estrogen, 113 inhibin/activin, 112 MIS/AMH, 112–113 Hormones and spermatogonial development, 437–448 AR, 438 cellular target for testosterone action, 441–442 contrary effects of hormones, 439–440 estradiol, 441 estrogen receptors, 438–439 Fas-FasL, 445 FSH, 440–441 FSH receptor, 438

future directions, 445 GDNF, 444 hormone effects on normal spermatogenesis, 437–438 Notch pathway, 444–445 PAp block, 439 SCF-c-kit system, 444 testosterone, 440 testosterone-induced block, 442–443 Horse, 182–183 hTERT, 461 Huhtaniemi, Ilpo, 155 Human FSHR promoter, 294 Humic acid (HA), 333 Hypothyroidism, 397–398

I I-Smads, 228, 229t ICER, 159, 258 Id proteins, 259 Id1, 259 Id2, 259 Id3, 259 Id4, 259 Idiopathic hypogonadotrophic hypogonadism (IHH), 185 IFN-α, 111 IGF, 108–109, 108t IGF-I, 76, 108–109, 108t, 320 IGFBP3, 127t Igfr1, 56–57 IHH, 185 IL-1α, 111 IL-1β, 111 IL-6, 111 Ile160Thr, 158 ILs, 111 In vitro culture and expansion. See Long-term cultures and mammalian spermatogonia. In vitro techniques, 354 In vivo FSH actions, 171–197 deer, 182–183 E-T switch, 188–189 hamster, 181–182 historical overview, 171–172 horse, 182–183 humans, 184–188 monkey, 179t, 183–184 mouse, 174–177, 178t nonhuman primates, 179t, 183–184 nonseasonal breeders, 172–181 pig, 177, 180–181 rat, 172–174, 178t seasonal breeders, 181–183 sleep, 182 Indazole-3-carboxylic acids, 368–369 Indenopyridine, 369 Indifferent gonad, 47 Inhibin A, 164, 229 Inhibin-α, 17t, 390 Inhibin B, 164, 229, 383–384 Inhibins and activins, 112, 164, 229–232, 238–239 activin structure/synthesis, 229–230 clinical significance, 239–240 FSH, 230–231 germ cells, 232 inhibin structure/synthesis, 229 postmitotic Sertoli cells, 232 regulation of synthesis, 230–231 Sertoli cell proliferation, 231–232 synthesis patterns, 230

testis, 231–232 Inhibitory Smads, 228, 229t Inr, 290 Institute for Genomic Research (TIGR), 104 Insufficient apical cytoskeletal support, 347 Insulin-like growth factor (IGF), 108–109, 108t Insulin-like growth factor I (IGF-I), 76, 108–109, 108t, 320 Insulin-like growth factor II (IGF-II), 108t, 109 Insulin-like growth factor receptor 1 (Igfr1), 56–57 Insulin receptor (Ir), 56–57 Insulin receptor family, 56–57 Insulin-receptor-related receptor (Irr), 56–57 Interferon alpha (IFN-α), 111 Interleukin 1 alpha (IL-1α), 111 Interleukin 1 beta (IL-1β), 111 Interleukin 6 (IL-6), 111 Interleukins (ILs), 111 Intermediate region, 35 Internet web sites. See Web sites. Interpro, 145 Interstitial release of Sertoli cell proteins, 353–354 Irr, 56–57 Isolated FSH deficiency, 185–187 Isolation of spermatogonia, 451–453

J Jagged 1, 309–310 JAK-Stat, 311 Johnson, Kamin J., 345 Jost, Alfred, 62 jsd, 475t JunB, 258 JunD, 258 Juvenile hypothyroidism, 187

K K252A, 56 Kallmann’s syndrome, 185 Kininogens, 140 Kit ligand (KL), 48, 108t, 111–112, 420–421 Klinefelter’s syndrome, 393–395 Knockout of genes, 16–17 Kölliker, Albert, 5

L L-kininogens, 140 Lactate dehydrogenase A (LDHA), 165 Lag-2, 310–311 Laminin, 127t Laminin α5 chain, 53 Langur monkeys, 89. See also Seasonal breeders. Latent TGFβ binding protein (LTBP), 235 LCCCST, 389, 390 LDHA, 165 Leptin, 309, 312 “Let my children go,” 12 Leu601Val, 158 Leukemia inhibitory factor (LIF), 113, 322, 421, 454–456 Leydig cell, 319, 441–442 Leydig cell differentiation, 58f, 59 Leydig cells, 45 LH fishes and amphibians, 74 functional testis, 61–62 seasonal breeders, 83–84 Lhr, 293

491

Index Lhx1, 45 Lhx9, 46, 53, 265 LIM homeodomain (Lhx9), 265 Lim1, 46 Lipids, 33, 113 LNGFR, 110 Long-term cultures and mammalian spermatogonia isolation of spermatogonia, 451–453 proliferation/differentiation of spermatogonia, 462–463 spermatogonia in cultures/cocultures, 453–457 spermatogonial cell lines, 457–462 Loveland, Kate L., 227 LTAg, 457–459 LTBP, 235 Luteinizing hormone. See LH. Lymphocytes, 323 Lysosomal carboxypeptidase A, 124t Lysosomes, 28

M Mab4B6.3E10, 425 Macroorchidism, 187 Macrophage-Leydig cell interaction, 323 Magnetic bead technique, 452 Major acute phase protein, 140 Matrix metalloproteinase (MMP), 122–127 Matrix metalloproteinase-2 (MMP2), 124t, 125–126 Matrix metalloproteinase-9 (MMP9), 124t, 126 McCune-Albright syndrome, 187, 395 McGeer, Laura, 213 McLean, Derek, 95, 254–255, 271 MEHP, 363f, 364, 430 Meistrich, Marvin L., 437 MEM, 462 Membrane-type matrix metalloproteinase 1 (MT1-MMP), 124t, 125 Mer tyrosine kinases, 17t Merops, 145 Mesenchymal cell-epithelial cell interaction, 321 Mesonephric cell migration, 53–54 Mesonephros, 50 Metabolic insult, 347 Metzincins, 122–129 MGD, 384 Mice FSH, 174–177, 178t SCARKO, 205–207 spermatogonial transplantation, 472t TGFβ superfamily signaling components, 234t Microarrays, 97 Microtubule-dependent transport defects, 347–348 Microtubules, 31–32 Migration inhibiting factor (MIF), 111 Minimum media (MEM), 462 MIS promoter, 270 Mitochondria, 26–27 Mixed gonadal dysgenesis (MGD), 384 MMP, 122–127 MMP inhibitors (TIMPS), 124, 126 MMP2, 124t, 125–126 MMP9, 124t, 126 MMP-14, 124t Molecular, 101, 102t Monkey, 179t, 183–184 Mono-(2-ethylhexyl)phthalate (MEHP), 363f, 364, 430

Mother cells, 20 Mouse. See Mice. Mouse Genome Informatics, 104 MSC-1 cell, 330t, 333–334 MSC-1 cell line, 96 MT1-MMP, 124t, 125 mTERT, 461 mTOR, 309 Müllerian duct, 44–45, 58 Müllerian inhibiting substance (MIS), 45, 51, 112–113, 219–220, 232–233 Multivesicular bodies, 28 MuLVs, 430 Murine Fshr promoter, 294

N n-butylisocyanate, 355 N-cadherin, 204 N-CAM, 237 n-Hexane, 358 NCBI, 104 NCCIT, 457 NDFα, 322 NDFβ, 322 Neonatal testosterone surge, 200 Nerve growth factor (NGF), 110–111 Neu differentiation factors (NDF), 322 Neurotrophin-3 (NT-3), 55–56, 61, 111, 322, 424 Neurotrophin growth factor family, 55–56, 60–61, 108t, 110–111 Neurotrophin ligands, 55 Neurturin (NRTN), 236–237 NFαB, 266–267 NGF, 55, 61, 110–111 93RS2 cells, 330t, 338 Nishimune’s group (Japan), 454 Nitric oxide (NO), 401 Nitroaromatics, 361–363 Nitrobenzene, 361 NO, 401 Nonhuman primates, 179t, 183–184 Nonseasonal breeders, 172–181 Notch pathway, 444–445 Notch receptor system, 309–310 NRTN, 236–237 NT-3, 55–56, 61, 111, 322, 424 NTERA-2, 457 Nuclear receptors, 253t, 254t, 260–265 androgen receptor, 261–262 Cys2 His2 zinc-finger domains, 264–265 Cys4 zinc fingers, 264 estrogen receptors, 262 orphan nuclear receptors, 263–264 retinoic acid receptors, 262 thyroid hormone receptors, 262–263 Nuclear TR, 223 Nucleus/nucleolus, 21–26 Nutritional cell-cell interactions defined, 318, 318t Sertoli cell-Leydig cell interactions, 319

O Oehl, Eusebio, 3, 9 1-benzyl-1H-indazole-3-carboxylic acids, 368–369 1,3-dinitrobenzene, 361–363 One stem cell - one colony hypothesis, 475 Ontology classification of gene expression, 99–103 Orphan nuclear receptors, 263–264 Overview, 43–44 Ovine E box, 295

Ovine FSHR promoter, 294–295

P p21Cip1, 222–223 p27Kip1, 222–223 p70S6K, 307 p75NTR, 56 p100, 266f p105, 266f p300, 268 Paal-Knorr condensation, 358 Pagel, Walter, 445 PAI-1, 130–132 PAI-2, 130 PAp block, 439 Pathophysiology. See Sertoli cell pathophysiology. PCI, 130 PDGF-B, 55 PDGF-C, 55 PDGFRα, 55 PDGFRβ, 55 Pem, 97, 265 Pem-androgen regulated homeobox, 17t Percoll isolation, 451–452 Peritubular cells, 59 Peritubular myoid cell, 317, 321–323, 441 Permissive view, 16–17 Peroxisome proliferator-activated receptors (PPARs), 366–367 Persistence of Müllerian duct derivatives syndrome (PMDS), 384–386 Peutz-Jeghers syndrome (PJS), 390 Pfam, 145 PGC migration, 47 PGCs. See Gonocyte development and differentiation. PGD2, 48, 233 Phosphatidylserine synthase 2, 17t Phthalates, 363–367 Phthalic acid esters, 363 PI3-K, 307 PI3 kinase, 57 PIAS1, 261 Pig, 177, 180–181 PJS, 390 Plasmin substrates, 130 Plasminogen, 127t Plasminogen activator inhibitor 1 (PAI-1), 130–132 Plasminogen activator inhibitor 2 (PAI-2), 130 Plasminogen system, 129–133 Platelet derived growth factor (PDGF), 55, 61 PMDS, 384–386 PModS, 321 Polarization of Sertoli cells, 53 Porter, Karen L., 437 Postmitotic Sertoli cells, 232 PPARα, 367 PPARs, 366–367 Pre-Sertoli cells, 21 Precocious puberty, 187 Presenilin 1, 309 Primary spermatocytes, 75 Primordial germ cell migration, 47 Primordial germ cells (PGCs). See Gonocyte development and differentiation. Prins, Gail, 445 PRKARIA, 390 Pro-IL-1β, 127t Pro-MMP-2, 127t Pro-TGFβ, 127t

492

Index

Pro-TNFα, 127t Pro348Ala, 158 Pro519Thr, 158 Procollagen I N-endopeptidase, 124t Proliferation-Apoptosis (PAp) block, 439 Proliferation/differentiation of spermatogonia, 462–463 Promoter eukaryotic, 252 FSHR, 287–295 human, 294 MIS, 270 murine, 294 ovine, 294–295 rat, 287–294 SF-1, 270 TATA box-containing, 252 TATA-less, 252 Propylthiouracil (PTU), 220–221 Prostaglandin D2 (PGD2), 48, 233 Proteases/protease inhibitors, 121–152 ADAM-TS family, 127–128 cathepsins, 133–142. See also Cathepsins. chromosomal localization, 124t future directions, 144–145 metzincins, 122–129 MMPs, 122–127 nomenclature, 124t plasminogen system, 129–133 proteolysis, 121–122 seminiferous epithelium, 122, 123f terminology, 122 web sites, 145 Protective protein, 124t Protein C inhibitor (PCI), 130 Protein kinase A (PKA), 57, 159, 160f, 161f Protein secretion. See Sertoli cell gene expression and protein secretion. Proteinase nomenclature, 124t Proteolysis, 121–122 PTEN, 427

R R-Smad/Smad4 complex, 309 Radiation, 403–404 Raf, 311 Rapamycin, 307 RAR, 262 RARα, 262 RARβ, 262 RARγ, 262 Rat FSH, 172–178t FSHR promoter, 287–294 germ cell transplantation, 476 spermatogenic arrest, 477 Rat FSHR promoter, 287–294 CAMP, 292 cell specificity, 287 chromatin structure, 293–294 E box, 287–288 E2f, 289–290 GATA, 290 Inr, 290 methylation, 293 promoter function, 287 proximal promoter, 287–294 Sf-1, 290–292 transgenic mouse, 292–293 USF proteins, 289 Rat U34 GeneChip, 98

RE-3, 295 Receptor/second messenger alterations, 348 Receptor signaling, 306–310 Receptor Smads (R-Smads), 228, 229t, 309 Reciprocal proenzyme activation, 129 Regulatory cell-cell interactions defined, 318–319, 318t Sertoli cell-Leydig cell interactions, 319–320, 320t Sertoli cell-peritubular cell interactions, 321 Regulatory factors, 107. See also Sertoli cell secreted regulatory factors. REL proteins, 266f RELA, 266f RELB, 266f RER, 27, 33 Ret, 444. See also Glial cell line-derived neurotrophic factor (GDNF). Reticuloendotheliosis family (REL) of proteins, 266f Retinoic acid receptors (RAR), 262 Richburg, John H., 345 RNFN, 262 Roberts, Kenneth P., 329 Robertson, David M., 227 Rough endoplasmic reticulum (RER), 27, 33 Russell, Lonnie D., 3, 11–12, 15, 90, 473 rWIN, 266 RXR, 262 RXRβ, 262

S S14-1 cells, 330t, 337 SAGE, 429 Salameh, Wael A., 383 Saunders, Philippa, 213 Sawhney, Pragati, 371 SCARKO mouse, 205–207 SCF, 48, 108t, 111–112, 306–307, 444 SCF/c-kit system, 165, 309, 444 SCOS, 388–389 Seasonal breeders, 81–92 cell size, 85–86 FSH actions, 181–183 gonadal regression/recrudescence, 85 LH/FSH, 83–84 number of germ cells, 84 photoperiodic control, 82–84 plasma membrane surface area, 86 SER/RER, 86–88 spermatogenesis, 86–90 surface-to-volume ratio, 85 testis size, 84 Self-renewal and differentiation of germline stem cell, 450f Seminiferous cord formation, 52–58 endothelial cell migration/vasculature development, 58 estrogen/estrogen receptor, 56 FGF-9, 55 genes regulating initial transition/ aggregation, 53 growth factors, 54 HGF, 55 insulin receptor family, 56–57 mesonephric cell migration, 53–54 MIS/AMH, 58 morphological changes, 53 NGF family, 55–56 pattern of gene expression, 57–58 PDGF/PDGFRα, 55

signal transduction pathways, 57 Seminiferous epithelium, 122, 123f Seminiferous growth factor (SGF), 113 Seminiferous tubule fluid (STF), 208–209 Seminiferous tubule fluid secretion, 351–352 Seminiferous tubule/rete testis, 35 Serial amplification of gene ends (SAGE), 429 Sertoli, Enrico, 3–10, 20 completing the manuscript, 8–9 discovery, 5–8 laboratory, 4–5 microscope, 4 post-graduation, 9–10 Sertoli cell aggregation, 53 cytoplasm, 26–32 form/function, 19–21 mammals, in, 303–304 nucleus/nucleolus, 21–26 origin, 50–51 polarization, 53 proliferation, 59, 203, 231–232 type A/B, 21 Sertoli cell aging, 33 Sertoli cell apoptosis, 350–351 Sertoli cell barrier, 32–33 Sertoli cell cDNA library (Unigene database), 145 Sertoli cell cycle, 33 Sertoli cell cytoplasm, 26–32. See also Cytoplasm. Sertoli cell development, 41–92 embryonic Sertoli cell differentiation, 43–70 fishes and amphibians, 71–79 seasonal breeders, 81–92 Sertoli cell differentiation, 58f. See also Embryonic Sertoli cell differentiation. Sertoli cell dysfunction. See Conditions affecting Sertoli cells; Toxicants. Sertoli cell endocrinology and signal transduction, 153–247 androgen regulation, 199–216 FSH regulation at molecular/cellular levels, 155–169 in vivo FSH actions, 171–197 TGFB superfamily, 227–247 thyroid hormone regulation, 217–226 Sertoli cell function and gene expression, 93–152 protease/protease inhibitors, 121–152 Sertoli cell gene expression and protein secretion, 95–106 Sertoli cell secreted regulatory factors, 107–120 Sertoli cell gene expression and protein secretion, 95–106 chromosome distribution, 103–104 gene array analysis, 97–99 gene ontology analysis, 99–103 genes high expressed by/unique to Sertoli cells, 96–97 overview of gene expression, 96 web sites, 104 Sertoli cell-germ cell interactions, 48 Sertoli cell-Leydig cell interactions, 319–320 Sertoli cell lines, 329–342 15P-1 cells, 334–335 42GPA-9 cells, 335–336 45T-1 cells, 335 93RS2 cells, 338 ASC-17D cells, 336–337 ES cell lines, 338 MSC-1 cells, 333–334 overview, 330t S14-1 cells, 337

493

Index SerW3 cells, 338 SF-7 cells, 336 SG5-2 cells, 338 SK11 cells, 337 SM cell lines, 338 SMAT-1 cells, 337–338 TM4 cells, 330–333 TR-ST cells, 336 TTE-3 cells, 337 WL cell lines, 338 Sertoli cell markers, 383–384 Sertoli cell membrane modifications, 34 Sertoli cell mitochondria, 26–27 Sertoli cell neoplasia, 389–390 Sertoli cell nucleus, 21–30 Sertoli cell numbers, 354 Sertoli cell only syndrome (SCOS), 388–389 Sertoli cell origin, 50–51 Sertoli cell pathophysiology, 343–413 conditions affecting Sertoli cells, 383–413 toxicants, 345–382 Sertoli cell-peritubular cell interactions, 320–323 Sertoli cell proliferation, 59, 231–232 Sertoli cell secreted growth factor (SCSGF), 113 Sertoli cell secreted regulatory factors, 107–120 growth factors, 108–112. See also Growth factors. hormones, 107, 112–113 lipids, 113 overview, 107–108 SCSGF, 113 SGF, 113 uncharacterized factors, 113 Sertoli cell-somatic cell interactions, 317–328 categorization of cell-cell interactions, 318–319 cell biology, 317, 318f environmental interactions. See Environmental cell-cell interactions. macrophage-Leydig cell interaction, 323 nutritional interactions. See Regulatory cell-cell interactions. other interactions, 323 Sertoli cell-Leydig cell interactions, 319–320 Sertoli cell-peritubular cell interactions, 320–323 Sertoli cell toxicants, 345–382 altered germ cell attachment, 346–347 apical sloughing, 349–350 blood-testis barrier defect, 347 carbendazim, 355–357 changes in distribution, quantity, or biochemical properties, 352–353 cisplatin, 367–368 colchicine, 368 decreased seminiferous tubule fluid secretion, 351–352 definition, 345–346 experimental approaches to studying dysfunction, 354–355 γ-diketones, 357–361 germ cell apoptosis, 351 indazole-3-carboxylic acids, 369–370 indenopyridine, 369 insufficient apical cytoskeletal support, 347 interstitial release of Sertoli cell proteins, 353–354 MAA, 369 manifestations, 349–355 mechanisms, 346–348 metabolic insult, 347 microtubule-dependent transport defects, 347–348 nitroaromatics, 361–363

phthalates, 363–367 receptor/second messenger alterations, 348 reversibility, 370–371 Sertoli cell apoptosis, 350–351 Sertoli cell numbers, 354 shedding, 349–350 tri-o-cresyl phosphate, 369–370 vacuolation, 349 Sertoli cell transcriptional regulation, 250–299 structure/regulation of FSH receptor gene, 281–299 transcription factors, 251–280 Sertoli cell transplants, 354–355. See also Transplantation. Sertoli cell volume, 19 SerW3 cells, 330t, 338 Setchell, Brian P., 3, 11, 304 17α,20β-dihydroxyprogesterone, 76 Sex determination, 50–52 SF-1, 46, 49, 263, 290–292 SF-1 promoter, 270 SF-7 cells, 330t, 336 SG5-2 cells, 330t, 338 SGF, 113 SGP-2, 17t Shaper, Joel, 145 Sharpe, Richard M., 199 Shedding, 349–350 Shetty, Gunapala, 437 Shima, Jim, 271 SI/SId, 475t Siberian hamster, 82, 85, 89. See also Seasonal breeders. Side population of cells, 453 Sinha Hikim, Amiya P., 81 SIP-1, 267 60A class (BMP), 233 SK11 cells, 330t, 337 Skinner, Michael K., 43, 107, 251, 317 SLBS-3, 294 Sleep, 182 Sloughing, 349–350 SM cell lines, 330t, 338 Smad1, 309 Smad2, 309 Smad3, 309 Smad4, 228, 229t, 309 Smad5, 309 Smad6, 228, 229t Smad7, 228, 229t Smad8, 309 Small testes, 187 SMAT-1 cells, 330t, 337–338 Smith, Phillip, 171 Smooth endoplasmic reticulum, 27–28 SNURF, 262 Software (gene array analysis software), 99 Sox family of transcription factors, 267–268 Sox3, 267 Sox8, 51 Sox9, 51, 53, 267–268 SP cells, 453 SP1 family of GC-box binding transcription factors, 264–265 Spermatogenesis, 471 androgen regulation, 203–207 defects, 476–477 fishes/amphibians, 74–75 germ cell transplantation, 476–477 hormone effects, 437–438 recovery of, 211 seasonal breeders, 86–90

Spermatogonial cell lines, 457–462 Spermatogonial renewal and differentiation, 306 Spermatogonial self-renewal and differentiation, 450f Spermatogonial stem cell fate, 303–315 C. alegens, 304–305, 310–311 Drosophila, 305, 310–311 germ line stem cells, 311–312 receptor signaling, 306–310 stem cell renewal/differentiation, 306 Spermatogonial stem cells, 415–485 gonocyte developments/differentiation, 417–435 hormones and spermatogonial development, 437–448 long-term cultures of mammalian spermatogonia, 449–470 transplantation, 471–485 Spermatogonial transplantation, 471, 472f. See also Transplantation. Sry, 48, 50, 267 Sry expression, 48, 49t, 57 SSCT, 389 SSH, 429 Staput method, 98, 451 STAT-1, 255t, 268 STAT-3, 255t, 268, 309 STAT-5, 255t, 268 STATs, 255t, 268 Stefins, 140 Stem cell factor (SCF), 48, 108t, 111–112, 165, 306–307, 444 Stem cell renewal/differentiation, 306, 476 Stem cells. See Spermatogonial stem cells. Steroid hormones end receptors, 61 STF, 208–209 STF secretion, 351–352 Straight tubules, 35 Subcutaneous testicular xenotransplants, 354 Suppression subtraction hybridization (SSH), 429 SV40 LTAg, 457–459 Swerdloff, Ronald S., 383 Syrian hamster, 82, 85. See also Seasonal breeders.

T T3, 332 T antigen, 333 T-kininogen, 140 Tan, Karen, 213 TATA box-containing promoters, 252 TATA-less promoters, 252 TDS, 386–387 Telomerase, 461 Telomere shortening, 461 TER, 236 Teratocarcinomas, 427 Teratomas, 427 Terminal segment, 35 Testatin, 141 Testibumin, 165 Testicular cancer, 427–429 Testicular dysgenesis syndrome (TDS), 386–387 Testicular irradiation, 403–404 Testicular macrophage, 323 Testicular torsion, 401 Testis, 317 Testis cDNA library (Cancer Genome Anatomy Project database), 145 Testis cell-cell interactions. See Sertoli cell-somatic cell interactions. Testis development. See Embryonic Sertoli cell differentiation.

494 Testis tissue xenografting, 481–482 Testis volume, 187 Testosterone, 59, 437, 440. See also Hormones and spermatogonial development. Testosterone-induced block, 442 TGFα, 60, 108t, 109, 320, 322 TGFβ superfamily, 60, 108t, 109–110, 227–247, 309 antagonists, 228t, 238–239 BMPs, 233–234 GDNF, 236–238 inhibins/activins. See Inhibins and activins. Leydig cells, 320 members, 227 MIS, 232–233 overview, 227–228 peritubular cells, 322 regulation of TGFβ synthesis, 236 testis, 235–236, 240–241 TGFβ proteins, 235–236 TGFβ1, 60, 109, 235–236 TGFβ2, 60, 109, 235–236 TGFβ3, 60, 109, 235–236 TGFβRI, 60 TGFβRIII, 60 TGFRIII (betaglycan), 228t, 235, 238 TH, 384 3β-HSD, 59 Thyroid hormone, 61, 218, 395–397 Thyroid hormone receptors, 262–263 Thyroid hormone regulation, 217–226 AR expression, 219 humans, 220–221 MIS production, 219–220 p27Kip1/p21Cip1, 222–223 PTU, 220–221 questions to be answered, 223–224 Sertoli cell development, 218–219 species other than rats and mice, 220–221 T3, 219–224 TRS, 223 TRx1 mRNA expression, 221–222 Timeline, 8t TIMP-1, 126 TIMP-2, 126 TIMP-3, 126 Tissue inhibitors of metalloproteinases (TIMPs), 124, 126 Tissue-type plasminogen activator (tPA), 124t, 129–132 TM4 cells, 330–333 TNFα, 266 Toppari, Jorma, 155 Toxicants. See Sertoli cell toxicants. tPA, 124t, 129–132 TR-ST cells, 330t, 336 TRα1, 223, 224, 395 TRα1 mRNA expression, 221–222 TRα2, 395 Transcription factors, 251–280 beta scaffold factors with minor groove contact, 266–268 bHLH, 259–260

Index bZIP, 256–259 bZIP-bHLH, 260 categorization, 253t CBP, 268 defined, 251 developmental stage of Sertoli cells, 256t Dmrt1, 268–269 future directions, 270–271 helix–turn–helix motif, 265–266 mechanisms of modulation, 252–253 MIS promoter, 270 nuclear receptors, 254t, 260–265. See also Nuclear receptors. promoter. See Promoter. SF-1 promoter, 270 STATs, 268 transferrin, 269 zinc-finger domains, 260–265 Transcriptional regulation. See Sertoli cell transcriptional regulation. Transcriptome, 253 Transepithelial resistance (TER), 236 Transferrin, 17t, 269 Transforming growth factor. See TGFβ superfamily. Transforming growth factor α. See TGFα. Transgenesis, 480–481 Transient differentiating cells, 306 Transitional zone, 35 Transplantation, 354–355, 471–485 germ cell. See Germ cell transplantation. Sertoli cell transplants, 354–355 spermatogonial, 471, 472f subcutaneous testicular xenotransplants, 354 of testis tissue, 481–482 TRβ1, 223–224 TRβ2, 223 Tri-o-cresyl phosphate, 369–370 Trident, 266 Tripartite nucleolus, 21–22, 26f trkA, 56, 111 trkC, 56 TrnR1, 237 TrnR2, 237 True hermaphroditism (TH), 384 TTE-3 cells, 330t, 337 Tubulin, 360 Tubulobulbar complexes, 30–32 Tubulus rectus, 35 Tumors, 389–390 TUNEL, 351, 359 tvA, 429 2-methoxyacetic acid (MAA), 369 2,4,6-trinitrotoluene, 361 2,5-hexanedione, 357–361 Type A Sertoli cells, 21 Type A spermatogonia, 306–310, 454–456 Type B Sertoli cells, 21 Type B spermatogonia, 306 Tyro 3, 17t

U U34 GeneChip, 98 Undifferentiated spermatogonia, 306 Unigene database, 145 Unilateral compensatory hypertrophy, 187 uPAR, 130 Upd, 311 Urinary plasminogen activator, 124t URL addresses. See Web sites. Urokinase, 124t, 129–132 USF proteins, 260, 289 USF1, 260 USF2, 260

V Vacuolation, 349 Val61, 157 Variocele, 398–399 Vascular smooth muscle cell, 442 Vasculature development, 58 Vasectomy, 401 Verhoeven, Guido, 213 Vimentin staining, 31f Viscacha, 88–89. See also Seasonal breeders. Vogl, A. Wayne, 28, 30

W Water buffalo, 34 Web sites databases, 145 gene expression, 104 proteases/protease inhibitors, 145 Weizmann Institute of Science—GeneCards, 104 Wellcome Trust Sanger Institute, 104 WH proteins, 266 Wilms tumor gene protein/Wilms’ tumor 1 (WT1), 202, 265 WIN-1, 266 Winged helix (WH) proteins, 266 WL cell lines, 330t, 338 Wnt4, 49–50 Wolffian duct, 44 Wright, Pamela, 145 Wright, William W., 121 Wrobel, Karl-Heinz, 31f, 35 WT1, 202, 265 Wt1, 46, 48–49

X Xenotransplantation of germ cells, 477–479 XXY patients, 393–395

Z Z-score, 101, 102t Zinc, 353, 366 Zinc-finger domains, 253t, 254t, 260–265 Zirkin, Barry, 145