Male Infertility: Diagnosis and Treatment

medwedi.ru

Male Infertility Diagnosis and Treatment

DEDICATION

This book is dedicated to our wives, Sanderina Kruger and Laura Oehninger, who were always there over the last decades, inspiring us to achieve and to contribute.

Male Infertility Diagnosis and Treatment

Editors

Sergio C Oehninger

MD PhD

Professor, Departments of Obstetrics and Gynecology, and Urology and Division Director, The Jones Institute for Reproductive Medicine Eastern Virginia Medical School, Norfolk, Virginia USA

Thinus F Kruger

MD FRCOG

Professor and Chairperson Department of Obstetrics & Gynaecology, and Reproductive Biology Unit Tygerberg Academic Hospital and Stellenbosch University, Tygerberg South Africa

© 2007 Informa UK Ltd First published in the United Kingdom in 2007 by Informa UK Ltd, 4 Park Square, Milton Park, Abingdon, Oxon OX14 4RN. Informa Healthcare is a trading division of Informa UK Ltd. Registered Office: 37/41 Mortimer Street, London, W1T 3JH. Registered in England and Wales Number 1072954. Tel.: +44 (0)20 7017 6000 Fax: +44 (0)20 7017 6699 E-mail: [emailprotected] Website: www.informahealthcare.com All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording, or otherwise, without the prior permission of the publisher or in accordance with the provisions of the Copyright, Designs and Patents Act 1988 or under the terms of any licence permitting limited copying issued by the Copyright Licensing Agency, 90 Tottenham Court Road, London W1P 0LP. Although every effort has been made to ensure that all owners of copyright material have been acknowledged in this publication, we would be glad to acknowledge in subsequent reprints or editions any omissions brought to our attention. A CIP record for this book is available from the British Library. Library of Congress Cataloging-in-Publication Data Data available on application ISBN10: 0-415-39742-1 ISBN13: 978-0-415-39742-1 Distributed in North and South America by Taylor & Francis 6000 Broken Sound Parkway, NW, (Suite 300) Boca Raton, FL 33487, USA Within Continental USA Tel.: 1(800)272 7737; Fax: 1(800)374 3401 Outside Continental USA Tel.: (561)994 0555; Fax: (561)361 6018 E-mail: [emailprotected] Distributed in the rest of the world by Thomson Publishing Services Cheriton House North Way Andover, Hampshire SP10 5BE, UK Tel.: +44 (0)1264 332424 E-mail: [emailprotected] Composition by Parthenon Publishing Printed and bound in India by Replika Press Pvt. Ltd.

Contents Acknowledgments

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Foreword

Preface

List of Contributors

xix

Color section

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Section 1 – Basic concepts: sperm physiology and pathology 1. Anatomy and molecular morphology of the spermatozoon Christiaan F Hoogendijk, Thinus F Kruger, Roelof Menkveld

2. Physiology and pathophysiology of sperm motility Michaela Luconi, Elisabetta Baldi, Gustavo F Doncel

3. The pathophysiology and genetics of human male reproduction Christiaan F Hoogendijk, Ralf Henkel

4. Contribution of the male gamete to fertilization and embryogenesis Gerardo Barroso, Sergio Oehninger

5. Genome architecture in human sperm cells: possible implications for male infertility and prediction of pregnancy outcome Olga Mudrak, Andrei Zalensky

6. Sperm pathology: pathogenic mechanisms and fertility potential in assisted reproduction Hector E Chemes, Vanesa Y Rawe

7. Testicular dysgenesis syndrome: biological and clinical significance Niels Jørgensen, Camilla Asklund, Katrine Bay, Niels E Skakkebæk

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Section 2 – Diagnosis of male infertility 8. Evaluation of the subfertile male Agnaldo P Cedenho

117

9. The basic semen analysis Roelof Menkveld

141

10. Advances in automated sperm morphology evaluation Kevin Coetzee, Thinus F Kruger

171

11. Sperm morphology training and quality control programs are essential for clinically relevant results Daniel R Franken, Thinus F Kruger

181

12. Role of acrosome index in prediction of fertilization outcome Roelof Menkveld

187

13. Acrosome reaction: physiology and its value in clinical practice Daniel R Franken, Hadley S Bastiaan, Sergio Oehninger

195

14. Sperm–zona pellucida binding assays Sergio Oehninger, Murat Arslan, Daniel R Franken

209

15. Detection of DNA damage in sperm Ralf Henkel

225

16. Chromosomal and genetic abnormalities in male infertility Pasquale Patrizio, Jose Sepúlveda, Sepideh Mehri

239

17. Reactive oxygen species and their impact on fertility R John Aitken, Liga E Bennetts

255

18. How do we define male subfertility and what is the prevalence in the general population? T Igno Siebert, F Haynes van der Merwe, Thinus F Kruger, Willem Ombelet

269

19. DNA fragmentation and its influence on fertilization and pregnancy outcome Ralf Henkel

277

20. The impact of the paternal factor on embryo quality and development: the embryologist’s point of view Marie-Lena Windt

291

Section 3 – Therapeutic alternatives for male infertility 21. Clinical management of male infertility Murat Arslan, Sergio Oehninger, Thinus F Kruger

305

22. Urological interventions for the treatment of male infertility Victor M Brugh, Donald F Lynch Jr

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CONTENTS

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23. Medical treatment of male infertility Gerhard Haidl

333

24. Male tract infections: diagnosis and treatment Frank H Comhaire, Ahmed MA Mahmoud

345

25. Sperm-washing techniques for the HIV-infected male: rationale and experience Gary S Nakhuda, Mark V Sauer

351

26. Treatment of HIV-discordant couples: the Italian experience Augusto E Semprini, Lital Hollander

363

27. Artificial insemination using homologous and donor semen Willem Ombelet, Martine Nijs

375

28. Intracytoplasmic sperm injection: current status of the technique and outcome André Van Steirteghem

393

29. Sperm retrieval techniques for intracytoplasmic sperm injection Valérie Vernaeve, Herman Tournaye

401

30. Hyaluronic acid binding by sperm: andrology evaluation of male fertility and sperm selection for intracytoplasmic sperm injection Gabor Huszar, Attila Jakab, Ciler Celik-Ozenci, G Leyla Sati

413

31. In vitro maturation of spermatozoa Rosália Sá, Mário Sousa, Nieves Cremades, Cláudia Alves, Joaquina Silva, Alberto Barros

425

32. New developments in the evaluation and management of the infertile male Darius A Paduch, Marc Goldstein, Zev Rosenwaks

453

Index

461

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Acknowledgments We have been able to complete this work thanks to the devoted efforts of a few assistants. We wish to acknowledge the editorial assistance of Helena Krüger (from Tygerberg), who made a significant contribution to the textbook, but died sadly on 14 October 2005. We sincerely appreciate the excellent help of Madaleine du Toit, who took over the responsibilities for Helena. Irene Foy (from the Jones Institute) is thanked for her secretarial contributions. We also wish to acknowledge clinicians, scientists and laboratory personnel of the Reproductive

Biology Research Laboratory at the Department of Obstetrics and Gynaecology, Tygerberg Hospital, Stellenbosch University; the Vincent Palotti Hospital, Cape Town, Republic of South Africa; and the Jones Institute for Reproductive Medicine, Department of Obstetrics & Gynecology, Eastern Virginia Medical School, Norfolk, VA, USA. We are truly indebted to all contributors for their enthusiasm in making this project a success.

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Foreword In about one-half of all couples who are plagued by infertility, the male partner has a deficiency in his sperm. Infertility in the male has two very peculiar characteristics. First, even though details of the pathology of the sperm deficiency are not at all understood in most cases, there is a very good therapeutic modality which overcomes these problems and is still able to transmit the male partner’s genetic message to the next generation. This therapeutic modality is, of course, intracytoplasmic sperm injection (ICSI). This successful therapy has made it seem less urgent to investigate the pathophysiology of male infertility. This is unfortunate, as there is an inner concern and some evidence that ICSI may transmit to succeeding generations the seeds of an increased incidence of sperm defects. Section 1 and several chapters of Sections 2 and 3 of this book tell us what is known about this area and thus serve as a launching pad for the further necessary investigation of the pathophysiology of sperm deficiencies. These chapters also alert the clinician to our ignorance of the molecular details of at least some sperm problems, which may lead to the passing of these defects to the next generation by ICSI. There is no doubt, however, that ICSI is one of the major breakthrough ‘blockbuster’ treatments resulting in the enjoyment of children for couples who otherwise would not be able to. The second peculiar characteristic of male infertility is that it is often diagnosed by a most unlikely specialist – the gynecologist – simply because it is this

specialist who is most likely to be consulted first by those who are infertile. Thus, it is not surprising that the editors of this book are gynecologists who have specialized in problems of reproduction and superspecialized in problems of male infertility. Hence has come into existence the subspecialty of andrology, which has found a home most often within the broad field of obstetrics and gynecology. Special problems of infertility in the male are treated by the urologist, and in some countries by dermatologists, but the therapy of last resort, i.e. ICSI, is in the hands of reproductive endocrinologists who have at their fingertips the technology of in vitro fertilization (IVF). It is noteworthy that Male Infertility: Diagnosis and Treatment is a synthesis of current knowledge about human andrology, and comes from two departments of obstetrics and gynecology where it was realized, even before the era of IVF, that a new perspective was required if true progress was to be made in solving the problems of male infertility. Notwithstanding these considerations, the editors have assembled an outstanding list of contributors who thoroughly overview the approach to male infertility not only from the perspective of the reproductive endocrinologist but also from the urologist, dermatologist and medical scientist points of view. Andrology is by no means a matured discipline, as indicated above. However, this book is a superb summary of our current understanding of the art and science of this dynamic approach to the solution of a major portion of infertility. Howard W Jones Jr MD Professor Emeritus, The Jones Institute for Reproductive Medicine Department of Obstetrics & Gynecology Eastern Virginia Medical School Norfolk, VA USA

Preface Physicians dealing with childless couples are well aware of the high incidence of male infertility. Recent estimates indicate that a male factor is present in up to 40–50% of cases consulting for infertility. While the causes of male infertility are multiple, the therapeutic options have traditionally been more limited. Urological and medical interventions have been, and continue to be, successfully implemented in defined clinical scenarios. But, undisputedly, the explosive growth and efficiency of assisted reproductive technologies (ART) has changed the direction of the field of andrology. Without any doubt, the development of intracytoplasmic sperm injection (ICSI) constituted a significant advancement not only in the treatment of infertility but also in nurturing further development of the discipline of clinical andrology. As a microtechnique to assist fertilization, ICSI has allowed men with severely compromised semen parameters (patients with oligo-astheno-teratozoospermia, alone or in combination, presenting with antisperm antibodies and even with obstructive or non-obstructive azoospermia) to achieve their desire to establish a family. Spermatozoa are highly differentiated cells that have an essential function to fertilize the oocyte, leading to embryo development. Functionally competent sperm cells are the result of the complex processes of spermatogenesis that involve cell differentiation, multiplication (mitosis), acquisition of the haploid stage (meiosis) and a dramatic metamorphosis (spermiogenesis). Spermatozoa are released into

the epididymis (spermiation), where further maturational, structural, biochemical and functional changes (capacitation) take place. Gametogenesis and seminiferous tubule functions occur under strict endocrine and paracrine control. To fertilize the oocyte successfully, the spermatozoon must be able to perform the critical functions of migration, recognition and binding to the zona pellucida, penetration of the zona pellucida, binding to the oolemma, activation of the oocyte, nuclear decondensation and participation in pronuclear formation leading to syngamy. This complex sequence of events leads to multiple potential opportunities for errors and interference by a multitude of pathogenic mechanisms. Current treatment options for male infertility include a large number of urological procedures (reconstructive surgery in cases of ductal obstruction, correction of varicocele and others), medical– pharmacological interventions (use of hormones, antibiotics), low-complexity assisted reproductive procedures (such as intrauterine insemination therapy) and the more advanced and complex ART. However, despite that contemporary therapies have enhanced the opportunities for conception in couples suffering from male infertility, often these solutions are raised in the absence of a defined etiological or pathophysiological diagnosis. Male infertility is unfortunately still considered ‘idiopathic’ in a large proportion of cases. The first in vitro fertilization (IVF) child in the world, Louise Brown, was born in Bourn Hall, UK in 1978. She was followed by the first IVF birth in xi

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Australia in 1980; in Norfolk, USA in 1981 (Elizabeth Carr); in continental Europe in 1982; and in 1984 in Tygerberg, South Africa (reviewed in Fauser and Edwards 2005)1. Since the early 1980s, the efficiency of IVF has improved dramatically, with clinical pregnancy rates per transfer cycle increasing from the mid-teens to 30–50%, according to the individual prognosis group. This accomplishment has been achieved by continuing efforts resulting in improved ovarian stimulation protocols, optimized gametes and embryo in vitro culture conditions, superior techniques of oocyte retrieval and embryo transfer, and development of more efficient embryo cryopreservation programs. The field of andrology has grown exponentially in parallel to the developments in ART. A few of the most significant milestones and some relevant clinical papers are worth highlighting: • Manual for the examination of semen (WHO 1980, fourth revised edition 1999)2; • First paper on IVF and male infertility (Wood 1984)3; • Aneuploidy in human sperm using fluorescence in situ hybridization (FISH) (Joseph et al. 1984)4; • Chromosomal abnormalities in human sperm (Martin 1985)5; • Male factor and IVF: first years of Norfolk experience (Van Uem et al. 1985)6; • First human pregnancy by IVF with epididymal sperm in obstructive azoospermia (Temple– Smith et al. 1985)7; • Sperm morphology as a prognostic factor for IVF (Kruger et al. 1986)9;

• First pregnancies following preimplantation genetic diagnosis (PGD) from biopsied embryos sexed by Y-specific DNA amplification (Handyside et al. 1990)12; • ICSI: first pregnancies (Palermo et al. 1992)13; • Place of ICSI in the management of male infertility (Oehninger 2001)14; • Pregnancy after testicular sperm aspiration/ ICSI (Schoysman et al. 1993)15; • Microsurgical epididymal sperm aspiration/ ICSI and congenital absence of the vas deferens (Tournaye et al. 1994)16; • The essential partnership between diagnostic andrology and ART (Mortimer 1994)17; • Intrauterine insemination for male subfertility (Ombelet et al. 1995)18; • Pregnancies after ICSI with testicular sperm (Silber et al. 1995)19; • Pregnancies after ICSI with testicular sperm in non-obstructive azoospermia (Devroey et al. 1995)20; • Deletions of the Y chromosome and severe oligospermia (Reijo et al. 1996)21; • Infertility in ICSI-derived sons (Kent-First et al. 1996)22; • Prospective follow-up study of ICSI children (Bonduelle et al. 1996)23; • Thresholds for semen parameters in fertile versus subfertile populations (Ombelet et al. 1997)24; • Approaching the next millennium: management of andrology diagnosis in the ICSI era (Oehninger et al. 1997)25;

• IVF and epididymal aspiration in congenital absence of the vas deferens (Silber et al. 1987)8;

• Consensus workshop on diagnostic andrology (European Society of Human Reproduction and Embryology, ESHRE) (Fraser et al. 1997)26;

• Description and definition of the Tygerberg Strict Criteria (R Menkveld 1987 – PD thesis);

• Detection of aneuploidy in human sperm using FISH (14 chromosomes) (Pang et al. 1999)27;

• Births after microsurgical sperm aspiration/ IVF in men with congenital absence of the vas deferens (Patrizio et al. 1988)10;

• Forging a partnership between total quality management and the andrology laboratory (De Jonge 2000)28;

• Definition of male factor in ART (Acosta et al. 1989)11;

• A meta-analysis of sperm function tests (Oehninger et al. 2000)29;

PREFACE

• Testicular dysgenesis syndrome (Skakkebaek et al. 2001)30; • ICSI should not be the treatment of choice for all cases of in vitro conception (Oehninger and Gosden 2002)31; • Multiple gestations in ART: an ongoing epidemic (Adashi et al. 2003)32; • Identification of the subfertile male in the general population: suggested new thresholds (van der Merwe et al. 2005)33. The overall objective of this book is to deliver information in an approachable fashion about the most common pathogenic mechanisms involved in male infertility and the state-of-the-art diagnostic tools, and a detailed description of the current therapeutic options available for the infertile man. The organization of the book follows these goals. Objective evidence, supported by a thorough and updated list of references, is presented in each individual chapter. The contributing authors have presented easy-toread chapters and the outlined information should be readily understood by a variety of readers, including medical and postgraduate students, physicians and scientists interested in reproduction. Indeed, the main expectation is that a wide range of generalists and specialists (andrologists, reproductive endocrinologists, urologists, obstetricians and gynecologists, primary-care practitioners) will benefit from the information presented herein. It was not our aim to present a manual with recipes of screening tests or techniques, but rather to examine the rationale behind clinical management, always supported by evidence-based medicine. Notwithstanding these considerations, methods have been succinctly mentioned and the interested reader can access more technical details through the extensive cited bibliography. We were fortunate to assemble an outstanding and international group of contributors: six of the seven continents are represented (Europe, North and South America, Africa, Australia and Asia). This multidisciplinary group of authors includes clinicians and scientists who have had a significant impact as pioneers and/or have made distinguished contributions to the field of male infertility.

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Section 1 critically discusses ‘Basic concepts: sperm physiology and pathology’. In Chapter 1, CF Hoogendijk, TF Kruger and R Menkveld (from South Africa) provide a synopsis of the ‘Functional anatomy and molecular morphology of the spermatozoon’. The authors outline the basic anatomy of the human spermatozoon through a light- and electron-microscopic approach. In addition, they introduce the concepts of chromosomal arrangement and the high degree of organization of the sperm nuclear chromatin. In Chapter 2, M Luconi and E Baldi (from Italy) and GF Doncel (from the USA) present ‘The physiology and pathophysiology of sperm motility’. The authors describe with accuracy the mechanochemical basis of sperm movement, placing special emphasis on the regulatory factors involved in the acquisition and maintenance of sperm motility, hyperactivation and chemotaxis. The authors also discuss the molecular defects associated with asthenozoospermia, a sperm pathology that represents one of the main causes of male infertility, as well as systemic and in vitro therapeutic approaches for this condition. In Chapter 3, CF Hoogendijk and R Henkel (from Germany, now South Africa) delineate ‘The pathophysiology and genetics of human male reproduction’. This chapter reviews in detail the genetic controls that are operative at different steps of spermatogenesis, the nuclear chromatin organization levels and the role of spermatozoa in early embryogenesis. In Chapter 4, G Barroso (from Mexico) and S Oehninger (from the USA) describe the ‘Contribution of the male gamete to fertilization and embryogenesis’. A large body of evidence demonstrates that: (1) the fertilizing spermatozoon plays a significant part in bringing about the development of the zygote, with its contributions being well beyond the delivery of the paternal DNA; and (2) infertile men with or without altered ‘classic’ semen parameters may have associated sperm dysfunctions that can result in aberrant embryogenesis. This review focuses on examination of the paternal effects that become manifest before and after the major activation of embryonic gene expression. In Chapter 5, O Mudrak and A Zalensky (from the USA) present innovative work on ‘Genome

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architecture in human sperm cells: possible implications for male infertility and prediction of pregnancy outcome’. The concepts of chromosome territories, architecture, compactness and position, telomeres localization and the dynamic modifications during fertilization in the normal and abnormal situations are elegantly set forth. In Chapter 6, HE Chemes and VY Rawe (from Argentina) describe ‘Sperm pathology: pathogenic mechanisms and fertility potential in assisted reproduction’. The authors define sperm pathology as the discipline that characterizes structural and functional deficiencies in abnormal spermatozoa. They accurately detail phenotypes associated with sperm motility and morphology disturbances and the impact of non-specific anomalies and systematic defects of genetic origin. In Chapter 7, N Jørgensen, C Asklund, K Bay and NE Skakkebæk (from Denmark) present ‘Testicular dysgenesis syndrome: biological and clinical significance’. It is proposed that testicular cancer, hypospadias, cryptorchidism and low sperm counts are symptoms of a disease complex, the testicular dysgenesis syndrome (TDS), with a common origin in fetal life. The knowledge of the etiology of TDS is still rather limited, but environmental and life-style factors are suggested as contributing factors. The authors present a sophisticated description of how genetic polymorphisms or aberrations may render some individuals particularly susceptible to these exogenous factors. Section 2 discusses the ‘Diagnosis of male infertility’. Notwithstanding the major impact of IVF and ICSI, the approach to the assessment and treatment of male infertility is much more than simply ART. An exhaustive anamnesis and a thorough physical examination of the male partner are of paramount importance in the initial screening of the infertile couple. The cornerstone of the andrological evaluation in all cases is repeated semen analysis. A urological, endocrine, genetic and/or imaging workup should be implemented as appropriate. In Chapter 8, AP Cedenho (from Brazil) describes the ‘Evaluation of the subfertile male’. This chapter thoroughly delineates the clinical assessment of the male partner consulting for infertility, and how the work-up should be further individualized

according to the findings of the anamnesis and physical examination. In Chapter 9, R Menkveld provides an excellent state-of-the-art contribution on the ‘The basic semen analysis’, including laboratory performance, interpretation of results and quality-control guidelines. In Chapter 10, K Coetzee (from New Zealand) and TF Kruger present their extensive experience in ‘Advances in automated sperm morphology evaluation’. Automated systems have the power to increase the objectivity, precision and reproducibility of sperm morphology evaluations. As attractive as this option may seem, not many automated systems have been introduced into routine andrology laboratories. The majority of systems currently in operation are used in more experimental situations, because of the objective biological resolution of the systems. In Chapter 11, DR Franken (from South Africa) and TF Kruger give a powerful insight into why ‘Sperm morphology training and quality-control programs are essential for clinically relevant results’. The authors present prospective studies that clearly illustrate that an external quality-control program can be successfully implemented on condition that continuous monitoring is part of the program. In Chapter 12, R Menkveld updates ‘The role of the acrosome index in prediction of fertilization outcome’. Evidence is presented supporting the view that careful assessment of acrosome morphology provides extended information on the sperm fertilizing capacity. In Chapter 13, DR Franken, HS Bastiaan (from South Africa) and S Oehninger give a thorough presentation of the ‘Acrosome reaction: physiology and its value in clinical practice’. A simple and novel microassay using minimal volumes of solubilized zona pellucida is highlighted. The authors demonstrate that the use of a calcium ionophore or the natural solubilized zona pellucida in combination with fluorescent lectins constitute validated assays for assessment of the induced acrosome reaction in live sperm. The authors conclude that such tests should therefore be implemented in the functional evaluation of sperm from subfertile men, in order to guide clinical management properly. In Chapter 14, S Oehninger, M Arslan (from Turkey) and DR Franken provide a detailed

PREFACE

overview of ‘Sperm–zona pellucida binding assays’. Clinical data have demonstrated that successful sperm–zona pellucida binding is essential for the achievement of in vitro fertilization, and that abnormalities of this binding step are frequently present in subfertile men. Human sperm–zona pellucida interaction under in vitro conditions reflects multiple sperm functions, including the acquisition and completion of capacitation, recognition and binding to specific zona pellucida receptors and induction of the physiological acrosome reaction. The authors provide unequivocal evidence supportive of the use of sperm–zona pellucida binding assays in the clinical setting. In Chapter 15, R Henkel (from Germany, now South Africa) outlines ‘Detection of DNA damage in sperm’. The author describes a variety of techniques developed to examine sperm DNA, and presents a compelling view that testing for DNA integrity and damage should be introduced into the routine andrological laboratory work-up. In Chapter 16, P Patrizio, J Sepúlveda and S Mehri (from the USA) accurately review the ‘Chromosomal and genetic abnormalities in male infertility’. The authors outline a multitude of genetic and chromosomal aberrations diagnosed in infertile men, as well as detection methods and clinical significance. Based on the evaluated data, the authors outline a defined algorithm for genetic evaluation of the infertile male/infertile couple prior to and after ICSI. In Chapter 17, RJ Aitken and LE Bennetts (from Australia) elegantly describe ‘Reactive oxygen species and their impact on fertility’. The authors unequivocally demonstrate that excessive production or exposure to reactive oxygen species is both statistically and causally associated with defective sperm function and DNA damage. In Chapter 18, TI Siebert (from South Africa), FH van der Merwe (from South Africa), TF Kruger (from South Africa) and W Ombelet (from Belgium) outline ‘How do we define male subfertility and what is the prevalence in the general population?’. The authors critically discuss present standards for the definition of male subfertility/ infertility and their drawbacks, and introduce new thresholds based upon worldwide-derived experience.

In Chapter 19, R Henkel (from Germany, now South Africa) presents detailed information on ‘DNA fragmentation and its influence on fertilization and pregnancy outcome’. Over the past few years, the interest of scientists and clinicians has focused on the influence and involvement of sperm DNA fragmentation on and in fertility, as this parameter may have a serious impact on fertilization and pregnancy. The author thoroughly describes the potential mechanisms that may lead to DNA damage during spermatogenesis and sperm maturation. In Chapter 20, M-L Windt (from South Africa) extends these concepts with a detailed analysis of ‘The impact of the paternal factor on embryo quality and development: the embryologist’s point of view’. The author delineates the limitations of current methodologies used in the IVF laboratory to assess the impact of the male factor and to select embryos for transfer. Many studies have focused on embryo selection, and, especially since singleembryo transfer has become a goal in many countries, methods for selection of the genetically normal spermatozoon with the potential to contribute to normal embryo development are under current and active investigation. Section 3 delineates the ‘Therapeutic alternatives for male infertility’. In Chapter 21, M Arslan, S Oehninger and TF Kruger carry out a thorough description of the ‘Clinical management of male infertility’. The authors examine the causes and diagnostic and therapeutic management of the most common clinical scenarios, with emphasis on isolated and combined oligoastheno-teratozoospermia. The chapter provides defined avenues to be pursued following a state-ofthe-art diagnostic screening. In Chapter 22, VM Brugh and DF Lynch (from the USA) present an update on ‘Urological interventions for the treatment of male infertility’. This team of urologists elegantly describes varicocele repair, cryptorchidism and orchiopexy, disorders of ejaculation, ductal obstruction, vasovasostomy versus ICSI, congenital bilateral absence of the vas deferens and testis biopsy techniques. In Chapter 23, G Haidl (from Germany) outlines ‘Medical treatment of male infertility’. The author carefully presents medical options based on objective evidence as related to: (1) specific treat-

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ment (cases where hormonal supplementation is indicated in the form of gonadotropins, gonadotropin releasing hormone (GnRH), androgens, treatment of emission and ejaculatory disturbances and anti-infectious agents); and (2) empirical treatment (use of antiestrogens, aromatase inhibitors, purified/recombinant follicle stimulating hormone (FSH), antioxidants, carnitines, mast-cell blockers, phosphodiesterase inhibitors, zinc salts, kallidinogenase, adrenoceptor antagonists and antiphlogistic treatment). In Chapter 24, FH Comhaire and AMA Mahmoud (from Belgium) share their extensive experience on ‘Male tract infections: diagnosis and treatment’. The understanding of the link between infection of the accessory sex glands and reduced male fertility is scientifically acquired and diagnostic tools are available, but results of antibiotic treatment in terms of fertility remain disappointing. The latter is probably due to the irreversibility of functional damage caused by chronic infection/inflammation. The authors stress that prevention, early diagnosis and adequate treatment of infections of the male tract, both trivial and sexually transmitted, are of pivotal importance. In Chapter 25, GS Nakhuda and MV Sauer (from the USA) describe ‘Sperm-washing techniques for the HIV-infected male: rationale and experience’. The authors review the clinical aspects of providing fertility care for HIV-positive men and their uninfected female partners, focusing on the technical facets of sperm processing and options available for treatment. In Chapter 26, AE Semprini and L Hollander present their extensive observations on ‘Treatment of HIV-discordant couples – the Italian experience’, and discuss the evidence regarding human immunodeficiency virus (HIV) transmission and safe parenthood in men infected with HIV. Reproductive counseling and semen washing with ART are the milestones in offering reproductive assistance to these individuals. In Chapter 27, W Ombelet and M Nijs (from Belgium) outline the current status of ‘Artificial insemination using homologous and donor semen’. The authors argue that there is clear evidence in the literature that this low-complexity therapy can be offered as a first-line treatment in most cases of mild and moderate male-factor infertility, resulting in

acceptable pregnancy rates, before starting more invasive and more expensive techniques of assisted reproduction such as IVF and ICSI. A detailed description of indications, techniques, results and cost-efficiency is presented. In Chapter 28, A van Steirteghem (from Belgium) reviews ‘Intracytoplasmic sperm injection: current status of the technique and outcome’. Based on the pioneer work performed at his center, the author discusses the indications for and technique of ICSI, the outcome and children’s health (including pregnancy complications, major malformations, possible causes of adverse outcome and multiple pregnancies). In Chapter 29, V Vernaeve (from Spain) and H Tournaye (from Belgium) examine the techniques and indications of ‘Sperm retrieval for intracytoplasmic sperm injection’. The authors present a sophisticated description of surgical sperm retrieval in patients with obstructive and non-obstructive azoospermia, and predictive factors for success and outcome. They present an in-depth discussion of clinical questions, including testicular sperm extraction (TESE) by open biopsy or by percutaneous fine needle aspiration, multiple testicular biopsies or a single testicular biopsy, microsurgical or conventional testicular sperm extraction, how many TESE procedures and adverse effects of testicular sperm extractions. In Chapter 30, G Huszar, A Jakab, C CelikOzenci and GL Sati (from the USA) elegantly describe ‘Hyaluronic acid binding by human sperm: andrology evaluation of male fertility and sperm selection for intracytoplasmic sperm injection’. This group of authors introduces the novel concept of an association between a testis-expressed chaperone protein, sperm cellular maturity and function, including fertilizing potential, and frequencies of aneuploidy in human spermatozoa. In Chapter 31, R Sa, M Sousa, N Cremades, C Alves, J Silva and A Barros (from Portugal) outline ‘In vitro maturation of spermatozoa’. At present, the major goal of somatic cell–germ cell coculture systems is to establish a minimum of conditions that can artificially keep alive a more or less functional epithelium for a reasonable period of time. This group of investigators share their extensive experience with experimental studies of animal and human spermiogenesis in vitro. The objectives are directed not only to produce gametes in vitro for those cases

PREFACE

where no spermatids are found, but also to enable a more controlled study of the mechanism of action of toxins, hormones and signal molecules on the seminiferous epithelium. As a corollary, Chapter 32 by DA Paduch, M Goldstein and Z Rosenwaks (from the USA) presents a view to the future, with ‘New developments in the evaluation and management of the infertile male’. The authors highlight the significance of the following topics: (1) advances of genetics in male infertility; (2) the reproductive health of survivors of childhood and adult malignancies; (3) hormonal manipulation in the treatment of idiopathic infertility; (4) the use of alternative and integrative medicine in male infertility; and (5) surgical treatment of male infertility. The authors conclude that, ‘Over the next decade further developments in our understanding of the genetics and physiology of male reproduction, advances in stem cell research

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and better ways of measuring outcomes of surgical techniques, combined with novel therapeutic options, will allow us to offer treatment to patients who are considered sterile by today’s standards.’ We are enthusiastic about the book in its content and presentation of the state-of-the-art of the discipline of andrology. We also remain hopeful that extended cellular–molecular–genetic investigations of the processes of human spermatogenesis and sperm capacitation and interaction with the female gamete, as well as the paternal contributions to embryogenesis, will lead to improved therapies to alleviate human infertility further. As the human genome project and the area of proteomics/metabonomics and translational research advance, their results and those of studies performed in combination with more classic reproductive biology– endocrinology techniques will bring us near to the achievement of these goals. Sergio C Oehninger MD PhD Thinus F Kruger MD FRCOG

REFERENCES 1. Fauser BC, Edwards RG. The early days of IVF. Hum Reprod Update 2005; 11: 437 2. World Health Organization. WHO Laboratory Manual for the Examination of Human Semen and Sperm–Cervical Mucus Interaction, 4th edn. Cambridge: Cambridge University Press, 1999 3. Wood C. Selection of patients. In Wood C, Trounson A, eds. Clinical In Vitro Fertilization. Philadelphia: Springer-Verlag, 1984: 31 4. Joseph AM, Gosden JR, Chandley AC. Estimation of aneuploidy levels in human spermatozoa using chromosome specific probes and in situ hybridization. Hum Genet 1984; 66: 234 5. Martin RH. Chromosomal abnormalities in human sperm. Basic Life Sci 1985; 36: 91 6. Van Uem JF, et al. Male factor evaluation in in vitro fertilization: Norfolk experience. Fertil Steril 1985; 44: 375 7. Temple-Smith PD, et al. Human pregnancy by in vitro fertilization (IVF) using sperm aspirated from the epididymis. J In Vitro Fert Embryo Transf 1985; 2: 119 8. Silber S, et al. New treatment for infertility due to congenital absence of vas deferens. Lancet 1987; 2: 850

9. Kruger TF, et al. Sperm morphologic features as a prognostic factor in in vitro fertilization. Fertil Steril 1986; 46: 1118 10. Patrizio P, et al. Two births after microsurgical sperm aspiration in congenital absence of vas deferens. Lancet 1988; 2: 1364 11. Acosta A, Oehniger S, et al. Assisted reproduction and treatment of the male factor. Obstet Gynecol Surv 1989; 44: 1 12. Handyside A, et al. Pregnancies from biopsied preimplantation embryos sexed by Y-specific DNA amplification. Nature 1990; 334: 768 13. Palermo A, et al. Pregnancies after intracytoplasmic injection of single spermatozoon into an oocyte. Lancet 1992; 34: 17 14. Oehninger S. Place of intracytoplasmic sperm injection in management of male infertility. Lancet 2001; 357: 2068 15. Schoysman R, et al. Pregnancy after fertilisation with human testicular spermatozoa. Lancet 1993; 342: 1237 16. Tournaye H, et al. Microsurgical epididymal sperm aspiration and intracytoplasmic sperm injection: a new effective approach to infertility as a result of congenital

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18.

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22. 23.

24.

25.

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bilateral absence of the vas deferens. Fertil Steril 1994; 61: 1445 Mortimer D. The essential partnership between diagnostic andrology and modern assisted reproductive technologies. Hum Reprod 1994; 9: 1209 Ombelet W, Puttemans P, Bosmans E. Intrauterine insemination: a first-step procedure in the algorithm of male subfertility treatment. Hum Reprod 1995; 10 (Suppl 1): 90 Silber SJ, et al. High fertilization and pregnancy rate after intracytoplasmic sperm injection with spermatozoa obtained from testicle biopsy. Hum Reprod 1995; 10: 148 Devroey P, et al. Pregnancies after testicular sperm extraction and intracytoplasmic sperm injection in nonobstructive azoospermia. Hum Reprod 1995; 10: 1457 Reijo R, et al. Severe oligozoospermia resulting from deletions of azoospermia factor gene on Y chromosome. Lancet 1996; 347: 1290 Kent-First MG, et al. Infertility in intracytoplasmicsperm-injection-derived sons. Lancet 1996; 348: 332 Bonduelle M, et al. Prospective follow-up study of 877 children born after intracytoplasmic sperm injection (ICSI), with ejaculated epididymal and testicular spermatozoa and after replacement of cryopreserved embryos obtained after ICSI. Hum Reprod 1996; 11 (Suppl 4): 131 Ombelet W, et al. Semen parameters in a fertile versus subfertile population: a need for change in the interpretation of semen testing. Hum Reprod 1997; 12: 987 Oehninger S, Franken D, Kruger T. Approaching the next millennium: how should we manage andrology

26.

27.

28.

29.

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diagnosis in the intracytoplasmic sperm injection era? Fertil Steril 1997; 67: 434 Fraser L, et al. Consensus workshop on advanced diagnostic andrology techniques. ESHRE Andrology Special Interest Group. Hum Reprod 1997; 12: 873 Pang MG, et al. Detection of aneuploidy for chromosomes 4, 6, 7, 8, 9, 10, 11, 12, 13, 17, 18, 21, X and Y by fluorescence in-situ hybridization in spermatozoa from nine patients with oligoasthenoteratozoospermia undergoing intracytoplasmic sperm injection. Hum Reprod 1999; 14: 1266 De Jonge C. Commentary: forging a partnership between total quality management and the andrology laboratory. J Androl 2000; 21: 203 Oehninger S, et al. Sperm function assays and their predictive value for fertilization outcome in IVF therapy: a meta-analysis. Hum Reprod Update 2000; 6: 160 Skakkebaek NE, Rajpert-De Meyts E, Main KM. Testicular dysgenesis syndrome: an increasingly common developmental disorder with environmental aspects. Hum Reprod 2001; 16: 972 Oehninger S, Gosden RG. Should ICSI be the treatment of choice for all cases of in-vitro conception? No, not in light of the scientific data. Hum Reprod 2002; 17: 2237 Adashi EY, et al. Infertility therapy-associated multiple pregnancies (births): an ongoing epidemic. Reprod Biomed Online 2003; 7: 515 van der Merwe FH, et al. The use of semen parameters to identify the subfertile male in the general population. Gynecol Obstet Invest 2005; 59: 86

Contributors R. John Aitken PhD ScD FRSE ARC Centre of Excellence in Biotechnology and Development and Discipline of Biological Sciences University of Newcastle Callaghan, NSW Australia

Camilla Asklund MD University Department of Growth and Reproduction Rigshospitalet Copenhagen Denmark

Cláudia Alves BSc Department of Genetics Faculty of Medicine University of Porto Portugal

Elisabetta Baldi PhD Associate Professor in Clinical Pathology ‘DENOthe’ Andrology Unit Department of Clinical Physiopathology University of Florence Florence Italy

Murat Arslan MD Assistant Professor, Department of Obstetrics and Gynecology Mersin University, Mersin, Turkey and The Jones Institute for Reproductive Medicine, Department of Obstetrics & Gynecology Eastern Virginia Medical School Norfolk, VA USA

Alberto Barros MD PhD Cathedratic Professor and Director, Department of Genetics Faculty of Medicine University of Porto Centre for Reproductive Genetics A Barros Porto Portugal

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Gerardo Barroso MD Professor, Departamento de Obstetricia y Ginecologia, and Director de la División de Reproducción Asistida Instituto Nacional de Perinatologia México DF México Hadley S Bastiaan PhD Reproductive Biology Unit Obstetrics and Gynaecology Department Tygerberg Hospital and Stellenbosch University Tygerberg South Africa Katrine Bay MSc University Department of Growth and Reproduction Rigshospitalet Copenhagen Denmark Liga E. Bennetts Discipline of Biological Sciences University of Newcastle Callaghan, NSW Australia Victor M Brugh III MD Assistant Professor, Department of Urology Eastern Virginia School of Medicine and Consultant Urologist The Jones Institute for Reproductive Medicine Norfolk, VA USA

Agnaldo P Cedenho MD Professor, Laboratory of Human Reproduction Division of Urology Paulista School of Medicine Federal University of São Paulo UNIFESP São Paulo Brazil Ciler Celik-Ozenci PhD The Sperm Physiology Laboratory Department of Obstetrics and Gynecology Yale University School of Medicine New Haven, CT USA Hector E Chemes MD PhD Laboratory of Testicular Physiology and Pathology Center for Research in Endocrinology National Research Council (CONICET) Buenos Aires Children’s Hospital, Buenos Aires Argentina Kevin Coetzee PhD Fertility Associates Ltd Newtown, Wellington New Zealand Frank H Comhaire MD Professor, Center for Medical and Urological Andrology and Reproductive Endocrinology University Hospital Ghent Ghent Belgium Nieves Cremades BSc Chief Embryologist, IVF Unit Department of Gynecology General University Hospital of Alicante Spain

CONTRIBUTORS

Gustavo F Doncel MD PhD Professor of Obstetrics and Gynecology and Director, CONRAD Preclinical Research Department of Obstetrics and Gynecology Eastern Virginia Medical School Norfolk, VA USA Daniel R Franken PhD Professor, Department of Obstetrics and Gynaecology Tygerberg Hospital Tygerberg South Africa Marc Goldstein MD Professor of Urology and Professor of Reproductive Medicine Department of Urology Weill Medical College of Cornell University New York, NY; The Population Council Center for Biomedical Research New York, NY and Center for Reproductive Medicine and Infertility Weill Medical College of Cornell University New York, NY USA Gerhard Haidl MD PhD Department of Dermatology/Andrology Unit University of Bonn Bonn Germany Ralf Henkel PhD Department of Urology Friedrich Schiller University Jena Germany

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Lital Hollander BSc Clinica Ostetrica e Ginecologica Università di Milano Milan Italy Christiaan F Hoogendijk MSc Reproductive Biology Unit Department of Obstetrics and Gynaecology Tygerberg Hospital University of Stellenbosch Tygerberg South Africa Gabor Huszar MD Professor, The Sperm Physiology Laboratory Department of Obstetrics and Gynecology Yale University School of Medicine New Haven, CT USA Attila Jakab MD The Sperm Physiology Laboratory Department of Obstetrics and Gynecology Yale University School of Medicine New Haven, CT USA Niels Jørgensen MD PhD Certified Clinical Andrologist Specialist in Medical Endocrinology and Consultant University Department of Growth and Reproduction Rigshospitalet Copenhagen Denmark Michaela Luconi PhD Associate Professor, ‘DENOthe’ Andrology Unit Department of Clinical Physiopathology University of Florence Florence Italy

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Donald F Lynch Jr MD Professor and Chairman, Department of Urology and Professor of Obstetrics and Gynecology Eastern Virginia Medical School Norfolk, VA USA Ahmed MA Mahmoud MD PhD Center for Medical and Urological Andrology and Reproductive Endocrinology University Hospital Ghent Ghent Belgium Sepideh Mehri MD Research Fellow Yale Fertility Center Yale University New Haven, CT USA Roelof Menkveld PhD Andrology Laboratory Reproductive Biology Unit Department of Obstetrics and Gynaecology Tygerberg Hospital and Stellenbosch University Tygerberg South Africa Olga Mudrak The Jones Institute for Reproductive Medicine Norfolk, VA USA and Institute of Cytology Russian Academy of Sciences St Petersburg Russia

Gary S Nakhuda MD Assistant Professor, Division of Reproductive Endocrinology Department of Obstetrics and Gynecology College of Physicians and Surgeons Columbia University New York, NY USA Martine Nijs Mas Sc Department of Obstetrics and Gynecology Genk Institute of Fertility St Jans Hospital Genk Belgium Willem Ombelet MD PhD Professor, Department of Obstetrics and Gynecology Genk Institute of Fertility Technology ZOL Campus St Jan Genk Belgium Darius A Paduch MD PhD Assistant Professor of Urology and Assistant Professor of Reproductive Medicine Department of Urology Weill Medical College of Cornell University New York, NY; The Population Council Center for Biomedical Research New York, NY and Center for Reproductive Medicine and Infertility Weill Medical College of Cornell University New York, NY USA

CONTRIBUTORS

Pasquale Patrizio MD Professor of Obstetrics and Gynecology and Director Yale Fertility Center Yale University New Haven, CT USA Vanesa Y Rawe Laboratory of Biology, Research and Special Studies Center of Studies in Gynecology and Reproduction CEGyR Buenos Aires Argentina Zev Rosenwaks MD Professor of Obstetrics and Gynecology and Revlon Distinguished Professor of Reproductive Medicine Center for Reproductive Medicine and Infertility Weill Medical College of Cornell University New York, NY USA Rosália Sá BSc Lab Cell Biology Institute of Biomedical Sciences Abel Salazar and Department of Genetics Faculty of Medicine University of Porto Porto Portugal G Leyla Sati MS The Sperm Physiology Laboratory Department of Obstetrics and Gynecology Yale University School of Medicine New Haven, CT USA

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Mark V Sauer MD Professor and Vice Chairman, Department of Obstetrics and Gynecology Columbia University and Chief, Division of Reproductive Endocrinology College of Physicians and Surgeons Columbia University New York, NY USA Augusto E Semprini MD Clinica Ostetrica e Ginecologica Università di Milano Milan Italy Jose Sepúlveda MD Clinical Assistant Professor, Instituto Estudio Concepcion Humana Monterrey, México and Yale Fertility Center Yale University New Haven, CT USA T Igno Siebert MD Department of Obstetrics and Gynaecology Stellenbosch University Tygerberg South Africa Joaquina Silva MD Chief Embryologist, Centre for Reproductive Genetics A Barros Porto Portugal Niels E Skakkebaek MD PhD Professor, University Department of Growth and Reproduction Rigshospitalet Copenhagen Denmark

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Mário Sousa MD PhD Professor, Director Lab Cell Biology Institute of Biomedical Sciences Department of Genetics Faculty of Medicine University of Porto and Scientific Director Centre for Reproductive Genetics A. Barros Porto Portugal Herman Tournaye MD PhD Professor, Centre for Reproductive Medicine Brussels Free University Brussels Belgium F Haynes van der Merwe MD Department of Obstetrics and Gynaecology Stellenbosch University Tygerberg South Africa André Van Steirteghem PhD Professor and Director, Centre for Reproductive Medicine and Research Centre for Reproduction and Genetics Vrije Universiteit Brussels Belgium

Valérie Vernaeve MD PhD Instituto Valenciano de Infertilidad (IVI) IVI – Barcelona Barcelona Spain Marie-Lena Windt PhD Reproductive Biology Unit Department of Obstetrics and Gynaecology Tygerberg Hospital and Stellenbosch University Tygerberg South Africa Andrei Zalensky PhD Associate Professor, The Jones Institute for Reproductive Medicine Norfolk, VA USA and Institute of Cytology Russian Academy of Sciences St Petersburg Russia

Color section

(f) pk

qk pι qι

Color plate 1 (Figure 5.1) Chromosome organization in human sperm. (a) Chromosome territory: chromosome 6 (CHR6) (green) was localized using a painting probe. Total DNA counterstained with propidium iodide (PI) (red). (b) Centromeres (green) were visualized using immunofluorescence with antibodies against CENP-A (centromere protein A). Total DNA counterstained with PI (red). (c) Fluorescence in situ hybridization (FISH) using TTAGGG probe (yellow/green) shows that the majority of telomeres are joined as dimers and tetramers. Subtelomeric sequences located at the p and q arms of one chromosome are spatially close. Total DNA counterstained with PI (red). (d) Subtelomeric sequences located at the p and q arms of chromosome 3 (subTEL3q, pink; subTEL3p, emerald) are spatially close. Total DNA counterstained with diamidino-2-phenylindole (DAPI) (blue). (e) FISH using arm-specific probes microdissected from CHR1 (1q, green; 1p, red) indicates looping of this chromosome. Total DNA counterstained with DAPI (blue). (f) Schematic model of sperm nuclear architecture. Selected chromosome territories (pink and ocher), telomeres (TEL) (green circles) and centromeres (CEN) (red circles) are shown within a section through the nucleus. Non-homologous CEN are clustered into a chromocenter, while TEL interact at the nuclear periphery. Modified from Ward and Zalensky 1996 (reference 38)

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PY20

anti+AKAP3

PY20+anti+AKAP3

Color plate 2 (Figure 2.3) Immunofluorescence analysis of fixed and permeabilized human spermatozoa. Confocal microscopy of double immunolabeling for tyrosine phosphorylated proteins ((b), PY20 antibody, green) and Akinase anchoring protein 3 (AKAP3) ((d), anti-AKAP3 antibody, red) reveals positivity for both antibodies in sperm tails. Simultaneous analysis of dual fluorescence confirms that tyrosine phosphorylation corresponds to AKAP3 in the tail ((f), double fluorescence, yellow). (a), (c), (e), negative controls without primary antibody. From reference 9, with permission

P44

P09

P12

Color plate 3 (Figure 5.2) Determination of chromosome intranuclear localization using fluorescence in situ hybridization (FISH) with painting probes. (a) Typical patterns of chromosome 1 (CHR1) painting probe hybridization (yellow) in normal sperm. (b) Typical patterns of CHR1 arm-specific probe hybridization (1p, green; 1q, red) in normal sperm. (c) Patterns of CHR1 hybridization in three samples of abnormal sperm.

d a

c a

Color plate 4 (Figure 30.1) Left panel: Mature (a) and diminished-maturity sperm with cytoplasmic retention (b–e) after creatine kinase (CK) immunostaining. Right panel: CK-immunostained sperm–hemizona complex. Observe that only the clear-headed mature spermatozoa without cytoplasmic retention are able to bind

COLOR SECTION

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Color plate 5 (Figure 30.2) Human testicular biopsy tissues immunostained with HspA2 antiserum. Sections represent lower (upper panel) and high (lower panel) magnifications to illustrate the tubular structure, and staining pattern of the adluminal area. HspA2 expression begins in meiotic spermatocytes, but is predominant during terminal spermiogenesis in elongated spermatids and spermatozoa

Color plate 6 (Figure 30.3) A model of normal and diminished maturation of human sperm. In normal sperm, maturation HspA2 is expressed in the synaptonemal complex of spermatocytes, supporting meiosis. HspA2 is likely also involved in the processes of late spermiogenesis, such as cytoplasmic extrusion (represented by loss of the residual body, RB), plasma membrane remodeling and formation of the zona pellucida- and hyaluronic acid-binding sites (change from blue to red membrane and stubs). Diminished-maturity sperm lack HspA2 expression, which causes meiotic defects and a higher rate of retention of creatine kinase (CK) and other cytoplasmic enzymes, increased levels of lipid peroxidation (LP) and consequent DNA fragmentation, abnormal sperm morphology and deficiency in zona and hyaluronic acid binding

Chromosomal aneuploidies Cytoplasmic retention

DNA degeneration

Cytoplasmic extrusion ↑LP

RB Abnormal head shape HspA2

HspA2 expression

Deficient zona binding

Plasma membrane remodeling (Zona-binding site)

Diminished fertility in conventional fertilization

Normal maturation

Diminished maturation

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Color plate 7 (Figure 30.4) Sperm movement patterns on the hyaluronic acid-coated spots used for sperm selection. Mature sperm are bound, and diminished-maturity sperm remain motile. Sperm are stained with cyber green DNA stain (Molecular Probes, Eugene, OR) that permeates viable sperm

Y X 18 18

SGA

ST1-unpaired

ST1-paired

ST1-telophase I

ST2

Sa1

Color plate 8 (Figure 31.4) Cocultures. Fluorescence in situ hybridization (FISH) analysis of spermatogonia A (SGA), primary spermatocytes (ST1), secondary spermatocytes (ST2) and early round spermatids (Sa1). 18 = violet, X = yellow, Y = red

Section 1

Basic concepts: sperm physiology and pathology

1 Anatomy and molecular morphology of the spermatozoon Christaan F Hoogendijk, Thinus F Kruger, Roelof Menkveld

INTRODUCTION Acrosomal cap

This chapter summarizes light and electronmicroscopic features that outline the basic characteristics of the anatomy of the human spermatozoon. Furthermore, sperm chromosomes are discussed in terms of the highly ordered and specific structure and packaging of the chromatin, together with the potential relationship between the increased incidence of numerical chromosomal aberrations and abnormal sperm morphology observed in infertile men.

Head

Equatorial segment Postacrosomal region

Connecting piece

Midpiece

Flagellum Principal piece

LIGHT AND ELECTRON MICROSCOPIC MORPHOLOGICAL CHARACTERISTICS OF SPERMATOZOA

End segment

Figure 1.1 Schematic drawing of light microscopic human spermatozoon

Spermatozoa are highly specialized and condensed cells that do not grow or divide. A spermatozoon consists of a head, containing the paternal heredity material (DNA), and a tail, which provides motility (Figures 1.1 and 1.2). The spermatozoon is endowed with a large nucleus, but lacks the large cytoplasm that is characteristic of most somatic cells. Men are unique among mammals in the degree of morphological heterogeneity of spermatozoa found in the ejaculate1–3.

Sperm head Light microscopy

Human spermatozoa are classified using brightfield microscope optics on fixed, stained specimens2,3. The heads of stained human spermatozoa are slightly smaller than the heads of living

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Mature spermatozoon

Head

Nucleus

Acrosome

Neck

Vacuoles

Midpiece

Postacrosomal envelope

Mitochondria

Proximal part of tail

Cell membrane

9 Peripheral fibrils

Distal and end part of tail

9 Double fibrils Fibril sheath 2 Central fibrils

Figure 1.2

Light and electron microscopic diagrams of human spermatozoon

spermatozoa in the original semen, although the shapes are not appreciably different4. The normal head should be oval in shape. Allowing for the slight shrinkage that fixation and staining induce, the length of the head is about 3–5 µm, and the width 2–3 µm. These values span the 95% confidence limits of comparative data for both Papanicolaou-stained and living sperm heads4. Two slightly different types of normal spermatozoa head forms have been described, based on spermatozoa found in endocervical canal mucus after coitus3. The first and most common form, as identified under the microscope with bright-field illumination, is the perfectly smooth oval head; the second form is oval, still having a smooth or regular contour, but being slightly tapered at the postacrosomal end3. Since diversity is a fact of all biological systems, trivial variations must be regarded as normal3. The following head aberrations can be observed: head shape/size defects, including large,

small, tapering, pyriform, amorphous, vacuolated (> 20% of the head surface occupied by unstained vacuolar areas), and double heads, or any combination of these5. Human spermatozoa have a welldefined acrosomal region constituting about twothirds of the anterior head area2,3,5. They do not exhibit an apical thickening like many other species, but show a uniform thickness/thinning towards the end, forming the equatorial segment. Because of this thinning, the area is visualized as more intensely stained when examined with the light microscope. Depending on this staining intensity, the acrosome will appear to cover 40–70% of the sperm head. Scanning electron microscopy

Scanning electron microscopy (SEM) is useful for demonstration of the surface structures of spermatozoa in great detail. Owing to its threedimensional image, furthermore, it is possible to observe and interpret the complex structure of a

ANATOMY AND MOLECULAR MORPHOLOGY OF THE SPERMATOZOON

human spermatozoon more easily and completely than with either light or transmission electron microscopy. The sperm head is divided into two unequal parts by a furrow that completely encircles the head, i.e. the acrosomal and postacrosomal regions. The acrosomal region can represent up to two-thirds of the head length and, in some cases, a depression is noted in this area, which is regarded as morphologically normal. The equatorial segment is not always clearly visible with SEM. Just after the equatorial segment is the beginning of the postacrosomal region, which is marked by maximal thickness and width of the spermatozoon. The postacrosomal region is divided into two parts by the posterior ring, forming two equal bands. The band closest to the acrosome stands out1. The surface of the human spermatozoon, washed free of seminal plasma, appears smooth, without coarse particles. The only exception is the acrosome, especially the anterior part, that may frequently appear rough1.

devoid of plasma and outer acrosomal membrane and is covered only by the inner acrosomal membrane6. The equatorial segment of the acrosome persists more or less intact, since it does not participate in the acrosome reaction (Figure 1.3). The posterior portion of the sperm head is covered by the postnuclear cap, which is a single membrane. The equatorial segment consists of an overlap of the acrosome and the postnuclear cap (Figure 1.3). The nucleus (Figure 1.3), constituting 65% of the head, is composed of DNA conjugated with protein. The chromatin within the nucleus is very compact, and no distinct chromosomes are visible. Sperm nuclei can have incomplete condensation with apparent vacuoles. The genetic information carried by the spermatozoon is ‘encoded’ and stored in the DNA molecule, which is made up of many nucleotides. The hereditary characteristics transmitted by the sperm nucleus include sex determination1.

Light and electron microscopic and molecular morphological characteristics of spermatozoa

The electron microscopic morphological characteristics of human spermatozoa are presented in Figures 1.2–1.6. The sperm head is a flattened ovoid structure consisting primarily of the nucleus. The acrosome is a cap-like structure covering the anterior two-thirds of the sperm head (Figures 1.2 and 1.3), which arises from the Golgi apparatus of the spermatid as it differentiates into a spermatozoon. Unlike in other mammalian species, the acrosome of the human spermatozoon does not exhibit apical thickening, but has an anterior segment of uniform thickness. The acrosome contains several hydrolytic enzymes, including hyaluronidase and proacrosin, which are necessary for fertilization1. During fertilization of the egg, the enzymerich contents of the acrosome are released at the time of acrosome reaction. During fusion of the outer acrosomal membrane with the plasma membrane at multiple sites, the acrosomal enzymes are released. The anterior half of the head is then

Acrosome cap

Nucleus Vacuoles

Equatorial segment of acrosome

Neck region Proximal centriole Mitochondria

Figure 1.3 Schematic drawing of longitudinal section of sperm head

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Protoplasmic droplet Midpiece Mitochondria

Mitochondria Annulus flagellum Dense fibers

Principal piece Fibrous sheath

Annulus Fibrous sheath

Figure 1.4 Longitudinal section of region between the midpiece and principal piece of human spermatozoon

Molecular morphology

The sperm chromosome structure is very complex. Some of the attributes are similar to somatic cell DNA organization and others are unique to spermatogenic cells. Sperm DNA packaging can be subdivided into four levels. Level I: chromosomal anchoring by the nuclear annulus The two strands of naked DNA which make up each chromosome are attached to a sperm-specific structure, the nuclear annulus. This represents a novel type of DNA organization, termed chromosomal anchoring, that is found only in spermatogenic cells. The nuclear annulus is shaped like a bent ring, and is about 2 µm in length. It is found only in sperm nuclei, although it is currently unknown at what stage of spermiogenesis it is first formed. So far there is no evidence for a nuclear annulus-like structure in any somatic cell type. In contrast, there is evidence of its existence in hamster7, human8, mouse and Xenopus sperm nuclei. Its existence in a wide variety of species suggests a fundamental role in sperm function.

Figure 1.5

Longitudinal section through midpiece

Unique DNA sequences were found to be associated with the nuclear annulus. Ward9 termed these sequences NA-DNA. The existence of these unique sequences suggests that the nuclear annulus anchors chromosomes according to particular sequences and not by random DNA binding. By organizing the chromosomes so that the NADNA sites of each chromosome are aggregated onto one structure, the nuclear annulus may also affect the determination of sperm nuclear shape. For example, in the hamster spermatozoon, the longer chromosomes may extend into the thinner hook of the nucleus, while a portion of every chromosome is located at the nuclear annulus. This is supported by image analysis of the distribution of DNA throughout the hamster sperm nucleus, which demonstrates that the highest concentration of DNA in the packaged sperm nucleus is at the base, where the nuclear annulus is located; in contrast, the lowest concentration of DNA is in the hooked portion10.

ANATOMY AND MOLECULAR MORPHOLOGY OF THE SPERMATOZOON

Cell membrane Midpiece

Dense fibers Flagellum Doublet tubules

Mitochondria

Central pair Fibrous sheath Principal piece

Figure 1.6

Cross-section of human sperm tail

This hypothesis is further supported by electron microscopic evidence that the chromatin near the implantation fossa is one of the first areas to condense during spermiogenesis11. Thus, the nuclear annulus may represent the only known aspect of sperm chromatin condensation that is specific for individual chromosome sites. Level II: sperm DNA loop domain organization Anchored chromosomes are organized into DNA loop domains. Parts of the nuclear matrix, protein structural fibers, attach to the DNA every 30–50 kb by specific sequences termed matrix attachment regions (MARs). This arranges the chromosome strands into a series of loops. This type of organization can be visualized experimentally in preparations known as nuclear halos. Halos consist of loops of naked DNA, 25–100 kb in length, attached at their bases to the matrix. Each loop domain visible in the nuclear halo consists of a structural unit of chromatin that exists in vivo in a condensed form.

The organization of DNA into loop domains is the only type of structural organization resolved thus far that is present in both somatic and sperm cells. In somatic cells, DNA is coiled into nucleosomes, then further coiled into a 30-nm solenoidlike fiber and then organized into DNA loop domains. The corresponding structures in sperm chromatin have a very different appearance. Protamine binding causes a different type of coiling, and DNA is folded into densely packed toroids, but still organized into loop domains. Mammalian sperm nuclei contain a small amount of histones that are presumably organized into nucleosomes12,13, but most of the DNA is reorganized by protamines. This means that with the evolutionary pressure to condense sperm DNA, all aspects of chromatin structure are sacrificed other than organization of the DNA into loop domains. This suggests that DNA loop domains play a crucial role in sperm DNA function. Level III: protamine decondensation The binding of protamines condenses the DNA loops into tightly packaged chromatin. DNA protamine binding forms toroidal or doughnut-shaped structures in which the DNA is very concentrated14. During spermiogenesis, histones, the DNAbinding proteins of somatic spermatogenic precursor cells, are replaced by protamines. Since histone-bound DNA requires much more volume than the same amount of DNA bound to protamines15, this change in chromatin structure probably accounts for some of the nuclear condensation that occurs during spermiogenesis. Protamines bind DNA along the major groove; this completely neutralizes DNA so that neighboring DNA strands bind to each other by van der Waals forces. Protamine binding leads to condensation and preservation of the DNA loop domain organization present in the round spermatid9. Level IV: chromosome organization The results of several studies10,16,17 have led to the proposal of a model18 in which there are limited constraints on the actual position of the chromosomes in the sperm nucleus. The NA-DNA sequences are

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located at the base of the nucleus, centromeres are located centrally and telomeres are located peripherally. Outside these three constraints, the folding of the chromosomal p and q arms is flexible.

narrows towards the posterior end. A longitudinal column and transverse ribs are visible. The short endpiece has a small diameter due to the absence of outer fibers1. Transmission electron microscopy

Sperm tail Light microscopy

Sperm tail formation arises at the spermatid stage. During spermatogenesis the centriole is differentiated into three parts: midpiece, main or principal piece and endpiece (Figures 1.1 and 1.2). The midpiece is of similar length to the head, and is separated from the tailpiece by a ring, the annulus (Figure 1.5). The following tail aberrations can be observed: • Neck and midpiece aberrations include their absence (seen as ‘free’ or ‘loose’ heads), noninserted or ‘bent’ tail (the tail forms an angle of about 90° with the long axis of the head), distended/irregular/bent midpiece, abnormally thin midpiece (i.e. no mitochondrial sheath) or any combination of these5; • Tail aberrations include short, multiple, hairpin, broken (angulation > 90°) tails, irregular width, coiling tails with terminal droplets or any combination of these5; • Cytoplasmic droplets greater than one-third of the area of a normal sperm head are considered abnormal. They are usually located in the neck/midpiece region of the tail, although some immature spermatozoa may have a cytoplasmic droplet at other locations along the tail3,5. The endpiece is not distinctly visualized by light microscopy.

The midpiece possesses a cytoplasmic portion and a lipid-rich mitochondrial sheath that consists of several spiral mitochondria, surrounding the axial filament in a helical fashion (Figures 1.2, 1.5 and 1.6). The midpiece provides the sperm with the energy necessary for motility. The central axial core of eleven fibrils is surrounded by an additional outer ring of nine coarser fibrils (Figures 1.2 and 1.6). Individual mitochondria are wrapped around these outer fibrils in a spiral manner to form the mitochondrial sheath, which contains the enzymes involved in the oxidative metabolism of the sperm (Figures 1.2 and 1.4–1.6). The mitochondrial sheath of the midpiece is relatively short, being slightly longer than the combined length of the head and neck1. The principal piece (main piece), the longest part of the tail, provides most of the propellant machinery. The coarse nine fibrils of the outer ring diminish in thickness and finally disappear, leaving only the inner fibrils in the axial core for much of the length of the principal piece (Figure 1.2)19. The fibrils of the principal piece are surrounded by a fibrous tail sheath, which consists of branching and anastomosing semicircular strands or ‘ribs’ held together by their attachment to two bands that run lengthwise along opposite sides of the tail1. The tail terminates in the endpiece with a length of 4–10 µm and a diameter of < 1 µm. The small diameter is due to the absence of the outer fibers and sheath and distal fading of microtubules.

Scanning electron microscopy

With SEM the tail can be subdivided into three distinct parts, i.e. midpiece, principal piece and endpiece. In the midpiece the mitochondrial spirals can be clearly visualized. This ends abruptly at the beginning of the midpiece. The midpiece

SPERM MORPHOLOGY AND CHROMOSOMAL ANEUPLOIDIES Many authors have studied the association between abnormal sperm shape and increased

ANATOMY AND MOLECULAR MORPHOLOGY OF THE SPERMATOZOON

frequency of aneuploidies. The conclusions of these studies are inconsistent; this is most probably because the sperm attributes were evaluated in the same semen sample, but not in the same sperm. As early as 1991, Martin studied sperm karyotypes20. She demonstrated that all chromosomes undergo nondisjunction during spermiogenesis, but that the G-group chromosomes (21 and 22) and the sex chromosomes have a significantly increased frequency of aneuploidy. Using fluorescence in situ hybridization (FISH), Spriggs and co-workers21 determined that most chromosomes have a disomy frequency of approximately 0.1% (1/1000); in contrast, the sex chromosomes and chromosomes 21 and 22 have a significantly increased frequency of aneuploidy. Thus, the sex chromosome bivalent and the G-group chromosomes are more susceptible to nondisjunction during spermatogenesis. Bernardini et al.22 suggested a relationship between increased frequencies of aneuploidy and diploidy in semen samples containing spermatozoa with enlarged heads. Several other studies have concluded that morphologically abnormal sperm may also have a significantly increased risk for being aneuploid23–27. An interesting report, based on the examination of sperm injected into mouse oocytes, suggested that in semen samples with high incidences of amorphous, round and elongated sperm heads, there was an increased proportion of structural chromosome abnormalities, such as chromosome and chromatid fragments and dicentric and ring chromosomes, but no increase in numerical chromosomal aberrations28. Further, Ryu et al.29 studied 120 normal and abnormal sperm (according to Tygerberg strict criteria) each in eight men, and concluded that normal morphology is not a valid indicator for the selection of sperm with haploid nuclei. Rives et al.30 showed that although the disomy frequencies of infertile males were directly related to the severity of oligozoospermia, there was no relationship between aneuploidy frequency and abnormal

morphology. In men with increased levels of globozoospermia, shortened flagella syndrome or sperm with acrosomal abnormalities, no association was found between sperm shape and numerical chromosomal aberrations31. In another study, De Vos and co-workers32 determined the influence of individual sperm morphology on fertilization, embryo morphology and pregnancy outcome after intracytoplasmic sperm injection (ICSI). With regard to the different morphological defects observed, they found the following fertilization rates: 63.4% (52 of 82) for spermatozoa with elongated heads; 63.3% (124 of 196) for spermatozoa with cytoplasmic droplets; 59.6% (223 of 374) for spermatozoa with amorphous heads; and 34.1% (15 of 44) for spermatozoa with broken necks. One hundred and one injected spermatozoa showed a combination of two morphological defects (overall fertilization rate, 57.4%). No fertilization ensued from six round-headed spermatozoa lacking acrosomes, and 12 spermatozoa showing vacuoles in their acrosomes provided a fertilization rate of 66.6%. These authors concluded that sperm morphology assessed at the moment of ICSI correlated well with fertilization outcome but did not affect embryo development. Furthermore, the implantation rate was lower when only embryos resulting from injection of abnormal spermatozoa were available. Recently, Celik-Ozenci and co-workers33 studied the relationship between sperm shape and numerical chromosomal aberrations in individual spermatozoa, using FISH, objective morphometry and sperm dimension and shape assessment, along with Tygerberg strict criteria. The results indicate that numerical chromosomal aberrations can be present in sperm heads of any size or shape, but the risk is greater with amorphous sperm. Even the most normal-appearing sperm with normal head and tail size could be disomic or diploid, although diploidy is less prevalent with normal sperm dimensions and shape.

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CONCLUSIONS Although many of the structures described here, especially the ultrastructural characteristics based on electron microscopy studies, are not visible by standard light microscopic examination, a basic knowledge of these structures is very important for the correct evaluation and interpretation of sperm morphology. In turn, this information will assist the clinician in the estimation of male fertility potential. From the molecular structure of the sperm, it is evident that the sperm DNA is packaged within the nucleus in an extremely complex and ordered fashion; there is, however, some degree of flexibility to this organization. A detailed model of how chromosomes are packaged in the sperm nucleus is gradually emerging; implications of this knowledge are already having an impact upon the study of fertility, particularly in preparations of nuclei for ICSI, diagnosis of semen samples and understanding the fate of sperm DNA after fertilization. As our knowledge of sperm chromatin increases, it is becoming more evident that visual assessment is an unreliable method for selection of sperm for ICSI. More specific methods for sperm selection, such as hyaluronic acid binding34, may alleviate the problem of fertilization with sperm of diminished maturity and genetic integrity during ICSI.

8. 9.

10.

11. 12.

13.

14.

15.

16.

17.

REFERENCES 1. Hafez ESE. Human Semen and Fertility Regulation in Men. St Louis: CV Mosby, 1976 2. Kruger TF, et al. Sperm morphological features as a prognostic factor in in vitro fertilization. Fertil Steril 1986; 46: 1118 3. Menkveld R, et al. The evaluation of morphological characteristics of human spermatozoa according to stricter criteria. Hum Reprod 1990; 4: 586 4. Katz DF, et al. Morphometric analysis of spermatozoa in the assessment of human male fertility. J Androl 1986; 7: 203 5. World Health Organization. WHO Manual for the Examination of Human Semen and Sperm–Cervical

18.

19.

20.

21.

Mucus Interaction, 2nd edn. London: Cambridge University Press, 1992 Barros C, Franklin LE. Behavior of the gamete membranes during sperm entry into the mammalian egg. J Cell Biol 1986; 37: 13 Ward WS, Coffey DS. Identification of a sperm nuclear annulus: a sperm DNA anchor. Biol Reprod 1989; 41: 361 Barone JG, et al. DNA organization in human spermatozoa. J Androl 1994; 15: 139 Ward WS. DNA loop domain tertiary structure in mammalian spermatozoa. Biol Reprod 1993; 48: 1193 Ward WS, et al. Localization of the three genes in the assymetric hamster sperm nucleus by fluorescent in situ hybridization. Biol Reprod 1996; 54: 1271 Loir M, Courtens JL. Nuclear reorganization in ram spermatids. J Ultrastruct Res 1979; 67: 309 Tanphaichitr N, et al. Basic nuclear proteins in testicular cells and ejaculated spermatozoa in man. Exp Cell Res 1978; 117: 347 Choudhary SK, et al. A haploid expressed gene cluster exists as a single chromatin domain in human sperm. J Biol Chem 1995; 270: 8755 Hud NV, Downing KH, Balhorn R. A constant radius of curvature model for the organization of DNA in toroidal condensates. Proc Natl Acad Sci USA 1995; 92: 3581 Ward WS, Coffey DS. DNA packaging and organization in mammalian spermatozoa: comparison with somatic cells [Review]. Biol Reprod 1991; 44: 569 Zalensky AO, et al. Well-defined genome architecture in the human sperm nucleus. Chromosoma 1995; 103: 577 Haaf T, Ward WS. Higher order nuclear structure in mammalian sperm revealed by in situ hybridization and extended chromatin fibers. Exp Cell Res 1995; 219: 604 Ward WS, Zalensky A. The unique, complex organization of the transcriptionally silent sperm chromatin. Crit Rev Eukaryot Gene Expr 1996; 6: 139 White IG. Mammalian sperm. In Hafez ESE, ed. Reproduction of Farm Animals, 3rd edn. Philidelphia: Lea & Febiger, 1974 Martin RH. Cytogenetic analysis of sperm from a man heterozygous for a pericentric inversion, inv(3)(p25q21). Am J Hum Genet 1991; 48: 856 Spriggs EL, Rademaker AW, Martin RH. Aneuploidy in human sperm: the use of mulicolor FISH to test various theories of nondisjunction. Am J Hum Genet 1996; 58: 356

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22. Bernardini L, et al. Study of aneuploidy in normal and abnormal germ cells from semen of fertile and infertile men. Hum Reprod 1998; 13: 3406 23. Colombero LT, et al. Incidence of sperm aneuploidy in relation to semen characteristics and assisted reproductive outcome. Fertil Steril 1999; 72: 90 24. Calogero AE, et al. Aneuploidy rate in spermatozoa of selected men with abnormal semen parameters. Hum Reprod 2001; 16: 1172 25. Rubio C, et al. Incidence of sperm chromosomal abnormalities in a risk population: relationship with sperm quality and ICSI outcome. Hum Reprod 2001; 16: 2084 26. Yakin K, Kahraman S. Certain forms of morphological anomalies of spermatozoa may reflect chromosomal aneuploidies. Hum Reprod 2001; 16: 1779 27. Templado C, et al. Aneuploid spermatozoa in infertile men: teratozoospermia. Mol Reprod Dev 2002; 61: 200 28. Lee JD, Kamiguchi Y, Yanagimachi R. Analysis of chromosome constitution of human spermatozoa with normal and aberrant head morphologies after injection into mouse oocytes. Hum Reprod 1996; 11: 1942

29. Ryu HM, et al. Increased chromosome X, Y, and 18 nondisjunction in sperm from infertile patients that were identified as normal by strict morphology: implication for intracytoplasmic sperm injection. Fertil Steril 2001; 76: 879 30. Rives N, et al. Relationship between clinical phenotype, semen parameters and aneuploidy frequency in sperm nuclei of 50 infertile males. Hum Genet 1999; 105: 266 31. Viville S, et al. Do morphological anomalies reflect chromosomal aneuploidies? Case report. Hum Reprod 2000; 15: 2563 32. De Vos A, et al. Influence of individual sperm morphology on fertilization, embryo morphology, and pregnancy outcome of intracytoplasmic sperm injection. Fertil Steril 2003; 79: 42 33. Celik-Ozenci C, et al. Sperm selection for ICSI: shape properties do not predict the absence or presence of numerical chromosomal aberrations. Hum Reprod 2004; 19: 2052 34. Huszar G, et al. Hyaluronic acid binding by human sperm indicates cellular maturity, viability, and unreacted acrosomal status. Fertil Steril 2003; 79: 1616

2 Physiology and pathophysiology of sperm motility Michaela Luconi, Elisabetta Baldi, Gustavo F Doncel

INTRODUCTION

the complex organization of the flagellum (Figure 2.1). With the exception of the distal part (endpiece) containing only the central couple of microtubules, the entire flagellum is organized in a cylindrical structure called the axoneme, consisting of nine pairs of tubulin A and B microtubules (doublets) connected to each other by nexin arms and to the central doublet by radial spokes. Each microtubule doublet is externally anchored to nine asymmetric outer dense fibers (ODFs), which are surrounded by the fibrous sheath in the principal piece and packed by mitochondria in the middle piece of the sperm tail (Figure 2.1). The base of the flagellum is thickened by a connecting piece consisting of nine segmented columns which distally fuse with the corresponding ODFs2, and is responsible for the transmission of tail movement to the head. The reciprocal sliding of each pair of microtubules originates from the sequential anchoring of the dynein arms to the neighboring doublet and adenosine triphosphate (ATP)dependent generation of sliding force. This sliding results in bends of alternating direction, which propagate the oscillation along the tail. The asymmetry of the axonemal structure as well as the outer microtubule connections to the central doublet and the ODF–fibrous sheath complexes confer a helical shape to the propagating flagellar beat. ODFs are essential for the development of forward motility in the mature sperm, and their

Mammalian spermatozoa become motile and acquire the ability to swim during their transit from the testis to the oviduct. These changes are initiated and controlled by several extra- and intracellular factors, which also play a pivotal role in regulating the acquisition of hyperactivated motility and chemotaxis. This chapter summarizes the mechanochemical basis of sperm movement, placing special emphasis on the regulatory factors involved in acquisition and maintenance of sperm motility, hyperactivation and chemotaxis. It also covers the molecular basis of asthenozoospermia, a sperm pathology characterized by reduced sperm motility, which represents one of the main causes of male infertility. Finally, it presents systemic and in vitro therapeutic approaches for asthenozoospermia, along with the most recent findings on pharmacological and physiological molecules capable of stimulating sperm motility.

MECHANOCHEMICAL BASIS OF SPERM MOTILITY Sperm swimming is characterized by a rhythmic, three-dimensional, asymmetric movement of the flagellum. This unique movement is assured by 13

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(a)

(b) MP

CP MS

Connecting piece

ODF

Midpiece

Microtubule pairs (MP)

Annulus

PM DA RS

Outer dynein arm Inner dynein arm

Outer dense fibers (ODF) 1 2

ODF

Principal piece

B-subunit

3 8

Nexin link 4

FS Endpiece

(c)

A subunit

Plasma membrane (PM)

Radial spoke (RS)

Central pair (CP)

Figure 2.1 Schematic representation of a human spermatozoon. (a) Longitudinal section showing head, middle piece, principal piece and endpiece. The insets on the right show the cytoskeletal organization of the sperm tail in transverse sections at different levels: middle (top), principal (center) and endpiece (bottom). Electron microscopy of transverse sections of the sperm tail at the levels of the middle (left) and the principal piece (right) are presented in (c). (b) Drawing showing organization of the axoneme. CP, central pair; MS, mitochondria; ODF, outer dense fibers; PM, plasma membrane; DA, dynein arms; RS, radial spoke; MP, microtubule pairs; FS, fibrous sheath. Modified from reference 1, with permission

structure and number are highly conserved throughout evolution. In particular, their crosssectional area correlates positively with the length of the flagellum3. Oscillations can originate in different regions of the flagellum; however, the beat frequency seems to be controlled by the basal region, which acts as a sort of pacemaker. Although different models have been proposed, the mechanism underlying the initiation of a new bend at the flagellar base is still unknown4. A recent paper on a knock-out mouse model for the functional dynein heavy chain has demonstrated the importance of these arms on the development of sperm motility5. In fact, mice in which the dynein inner-arm heavy

chain gene has been deleted show asthenozoospermic characteristics, with the majority of spermatozoa unable to achieve forward progressive motility. In such spermatozoa, the outer dense fibers retain their attachments to the inner surface of the mitochondria. These links are essential in normal spermatozoa for midpiece development, but disappear when spermatozoa acquire the ability to swim upon release from the epididymis. Conversely, disruption of dynein inner-arm heavy chains in knock-out mice results in insufficient force to overcome these bridges, and spermatozoa are unable to undergo normal tail bending. Energy to support the sliding force of the microtubules is provided by ATP, which is

PHYSIOLOGY AND PATHOPHYSIOLOGY OF SPERM MOTILITY

hydrolyzed by the dynein ATPase arms associated with the outer doublets of the microtubules. Although oxidative phosphorylation in midpiece mitochondria has long been considered a major source of ATP, local production of energy in the sperm principal piece through an alternative glycolytic enzyme pathway has recently been proposed as the main source of energy for flagellar movement. In fact, albeit reduced, motility is still present when mitochondrial oxidative phosphorylation is uncoupled in sperm6. Moreover, these two metabolic processes are strictly compartmentalized to the middle and principal pieces of the sperm flagellum, and although oxidative phosphorylation is more efficient than glycolysis in producing ATP, it is unlikely that ATP diffusion from the former to the latter compartment could supply enough energy to support flagellar movement in the distal region of the flagellum. Miki et al.7 elegantly demonstrated that the sperm-specific glycolytic enzyme glyceraldehyde-3-phosphate dehydrogenase-S (GAPDS, and its human ortholog GAPD2) is necessary for sperm motility and fertility, since sperm from Gapds(–/–) knock-out mice, in which oxidative phosphorylation is unaffected, generate only 10.4% of the ATP produced in wild-type controls. Moreover, sperm motility was impaired, with virtual absence of forward movement, and the mice were infertile7. Therefore, glycolysis seems to be the pivotal metabolism producing ATP for sperm motility. This concept is reinforced by the presence of sperm-specific isoforms of other glycolytic enzymes such as hexokinase and lactate dehydrogenase, which are selectively expressed in the sperm principal piece8.

REGULATION OF SPERM MOTILITY Upon release from the testis, human and all mammalian spermatozoa are immotile. In order to reach and fertilize the oocyte, they acquire the ability to swim during their transit through the epididymis and the female genital tract. Several

extra- and intracellular factors are important for the development and maintenance of sperm motility (Figure 2.2). These two processes appear to be regulated in a similar way. However, the majority of in vitro studies have been focused on the maintenance of sperm motility, using ejaculated or caudal epididymal spermatozoa. The following are some of the main factors regulating sperm movement.

Calcium Under physiological conditions, calcium is one of the most important ions regulating human sperm motility10. However, the role of calcium in activating spermatozoa has always been regarded as controversial. Indeed, voltage-gated, cyclic nucleotide-gated and transient receptor potential calcium channels have been described along the plasma membrane of the entire flagellum (for reviews see references 11 and 12), thus suggesting the importance of calcium entry for motility. Transient receptor potential calcium channels have recently been demonstrated in the sperm tail and are involved in stimulation of sperm motility by capacitation-dependent calcium entry13. Knock-out mice for the newly discovered CatSper calcium channel specifically expressed in the tail are infertile due to loss of progressive motility14. An increase in intracellular calcium levels is also indirectly implicated in the activation of intracellular calcium stores via inositol 1,4,5-triphosphate (IP3) signaling15,16. Upon entry, calcium activates phospholipases and modulates several enzyme activities. In particular, the activated calcium/calmodulin (CaM) complex has been shown to stimulate sperm motility through direct interaction with soluble adenylate cyclase (sAC)17,18, protein kinases19,20, phosphatases21 and phosphodiesterases22, finally leading to an increase in cyclic adenosine monophosphate (cAMP) and phosphorylation of sperm proteins. CaM has been characterized in sperm axonema and proposed as the intracellular calcium sensor regulating motility23. CaM levels are reduced in sperm from

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Kinases

Phosphatases 9. Acrosome reaction

10. Fertilization HCO3–

8. Chemotaxis

Ovum pH

ATP ROS

Cyclic nucleotides

PAF 3. Epididymal maturation and storage

7. Hyperactivation Intracellular Ca2+

6. Capacitation 2. Spermiation

Temperature Extracellular Ca2+

1. Spermatogenesis

Osmolarity 5. Forward motility

4. Ejaculation

Figure 2.2 Factors regulating sperm motility during the ‘sperm journey’ from the testis (right) to the ovary (left). External and intracellular factors controlling sperm motility are indicated (oval labels) together with the activation processes (numbered) that spermatozoa undergo during their transit through the male and female reproductive tracts. ATP, adenosine triphosphate; PAF, platelet-activating factors; ROS, reactive oxygen species. Modified from reference 9, with permission

asthenozoospermic patients24, and inhibitors of this enzyme negatively affect sperm motility25. Among CaM target enzymes, Marin-Briggiler et al.19 characterized a CaM-dependent protein kinase. Inhibition of the isoform IV of this kinase results in a specific decrease in motion parameters and ATP levels without affecting sperm viability, protein tyrosine phosphorylation or acrosome reaction19. Incubation of motile sperm in the absence of calcium dramatically reduces motion parameters19, suggesting the importance of calcium in the maintenance of human sperm motility. Extracellular calcium has been demonstrated to be essential for sperm motility. Evidence also suggests that its intracellular concentrations must be strictly regulated to allow for precise timing of sperm activation26,27. Decreasing levels of external calcium between the caput and cauda of the epididymis are associated with progressive development of sperm motility and an increase in protein

tyrosine phosphorylation28,29. Calcium addition to demembranated human sperm suppresses motility30, and increased intracellular calcium levels following cryopreservation negatively correlate with sperm motility and fertilizing ability31. Although many papers have focused on the role of calcium entry channels, very little is known about calcium extrusion from the cell. Recently, + plasma membrane Ca2 /calmodulin-dependent + Ca2 ATPases (PMCA) have been demonstrated to be essential for maintaining intracellular calcium homeostasis32. Indeed, homozygous male mice with a targeted gene deletion of PMCA isoform 4, which is highly enriched in the sperm tail, are infertile due to severely impaired sperm motility. Furthermore, this detrimental effect can be mimicked by inhibition of the enzyme in wild-type animals, thus supporting the hypothesis of a pivotal role of PMCA4 in the regulation of sperm function and intracellular Ca2+ levels32.

PHYSIOLOGY AND PATHOPHYSIOLOGY OF SPERM MOTILITY

The molecular mechanisms underlying such striking stimulatory and detrimental effects of calcium on sperm motility are still unclear; however, they seem to be linked to the activation of concurrent signaling pathways such as those involving protein kinases and phosphatases. Indeed, calcium levels must be kept low in order to prevent activation of phosphatases such as calcineurin27, which dephosphorylates and inactivates tail proteins involved in sperm motility27,33. An alternative hypothesis developed by Aitken’s group suggests that keeping internal calcium homeostasis in the presence of high extracellular calcium decreases ATP availability for tyrosine phosphorylation and sperm movement34.

Bicarbonate and adenylate cyclases Bicarbonate has long been demonstrated to enhance sperm motility in different species both in vitro and in vivo 35–38. The importance of this molecule in regulating sperm activation in vivo is further suggested by the increasing millimolar gradient of HCO3– that spermatozoa encounter during their journey from the testis to the site of fertilization. The increased level of HCO3– in seminal plasma compared with the epididymal fluid may allow motility to develop in the ejaculate. Okamura et al.39 showed a positive correlation between lower levels of HCO3- in the semen of infertile men with poor sperm motility. However, in male reproductive fluids, HCO3- levels must be kept low to prevent spermatozoa from undergoing premature activation and hyperactivated motility, processes that are stimulated by the 3–4-fold higher HCO3– concentrations present in the female reproductive tract40. The molecular mechanism by which HCO3– stimulates sperm motility involves a direct activation of sperm sAC, independent of intracellular pH41. sAC, which is insensitive to forskolin and G-protein regulation, and is selectively activated42–44 by HCO3–, appears to be the main adenylate cyclase present in mature spermatozoa, although different isoforms of the membrane

adenylate cyclase (mAC) have also been described45–47. In somatic cells, the precise compartmentalization of sAC in distinct subcellular microdomains provides the mechanism for localized cAMP rise specifically to activate protein kinase A (PKA) in different cellular compartments48,49. In fact, unlike mAC, sAC could diffuse and generate cAMP at the site where its target enzyme, PKA, is localized50. Sperm sAC activity, however, seems to be predominantly associated with the sperm particulate fraction51. Mice defective for sAC are infertile, apparently due to impairment of sperm motility52. Interestingly, motility can be restored in sAC knock-out mice by cAMP administration52. However, such treatment does not reverse hyperactivation and tyrosine phosphorylation defects or the sperm inability to fertilize, suggesting that sAC is also necessary for appropriate spermatogenesis and/or epididymal maturation53. Treating sperm with an inhibitor of sAC, KH7, the same authors were able to distinguish between sAC-dependent and independent processes during mouse sperm capacitation, showing that tyrosine phosphorylation of protein as well as sperm motility and hyperactivation are regulated by sAC, while the acrosome reaction is not53. A role played by mAC in controlling sperm motility, however, cannot be ruled out. In fact, selective knock-out of membrane olfactory adenylate cyclase 3 is associated with male infertility due to the sperm’s inability to penetrate the zona pellucida. These spermatozoa show a significant reduction in both motility and acrosome reaction54.

Kinases and phosphatases Although abundant evidence indicates the importance of protein phosphorylation as one of the key processes in transducing the stimulatory signals governing motility, little is known about the specific kinases and phosphatases involved. Generally, sperm motility has been demonstrated to be associated with increased tyrosine phosphorylation of specific sperm-tail proteins following

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tyrosine and serine–threonine kinase activation. Furthermore, sperm motility is negatively associated with phosphatase activation26,29,33,55. Tyrosine phosphorylated proteins in response to sperm capacitation are mainly localized in sperm tails56–58. A defect in the tyrosine phosphorylation of specific sperm proteins in response to capacitation has been described in asthenozoospermic patients, associated with reduced motility and hyperactivation capacity59,60. This defect in protein tyrosine phosphorylation seems to be linked to membrane fluidity in spermatozoa from asthenozoospermic patients60 and infertile men with varicocele61. Interestingly, even semen from normozoospermic men present distinct sperm subpopulations that show different plasma membrane fluidity and ability to undergo protein tyrosine phosphorylation and hyperactivation in response to capacitation62. Sperm protein phosphorylation is regulated by a finely tuned balance between kinase and phosphatase activities33,63. In particular, the adenylate cyclase/cAMP/PKA system has been demonstrated to be involved in tyrosine phosphorylation of different sperm proteins associated with motility27,56,63–66. cAMP produced by the activation of adenylate cyclase binds to PKA holoenzyme, inducing the release and activation of the catalytic subunit. Sperm treatments enhancing intracellular cAMP and PKA activity stimulate motility64,67. Since protein kinase A is a serine–threonine kinase, it is assumed that in order to stimulate tyrosine phosphorylation it activates some intermediate tyrosine kinases. An alternative pathway involving tyrosine kinase activation upstream to PKA has recently been reported by our groups55,68,69. In fact, both inhibition of phosphatidylinositol 3-kinase (PI3K) by LY294002 and physiological activation of sAC by bicarbonate stimulate an increase in intracellular cAMP levels in concurrence with enhanced tyrosine phosphorylation of the tail scaffolding protein, A kinase anchoring protein 3 (AKAP3). Confocal microscopy of fixed and permeabilized spermatozoa confirms that capacitation-induced tyrosine phosphorylation of sperm

proteins occurs mainly at the tail and, in particular, on AKAP3 (Figure 2.3). The stimulated phosphorylation of AKAP3 results in an increased binding of PKA regulatory subunit RIIβ, which is thus selectively recruited and activated in the sperm tail, where it interacts with its targets, finally resulting in an increase in sperm motility. Disruption of PKA–AKAP3 interaction results in the inhibition of sperm motility68. Sperm treatment with the PKA inhibitor H89 results in the inhibition of sperm motility, but not of AKAP3 tyrosine phosphorylation55,68, thus suggesting that PKA is involved in the regulation of sperm motility downstream to tyrosine kinases. Inhibition of motility and tyrosine phosphorylation following sperm treatment with H89 has been reported by other authors, conversely suggesting an upstream effect of PKA70–72. Such discrepancy could be explained either by differences in H89 concentrations and timing of H89 addition or by hypothesizing that tyrosine phosphorylation affects different targets upstream and downstream of PKA activation. The importance of AKAP scaffolding proteins in regulating sperm motility has recently been highlighted by targeted disruption of the Akap4 gene, whose product, AKAP4, is closely related to AKAP3. These mutant mice show defects in sperm flagella and motility resulting in infertility73. Contradictory reports exist regarding the alteration of AKAP genes in men affected by dysplasia of the fibrous sheath74,75. However, defects in the ability of such scaffolding proteins to undergo tyrosine phosphorylation, thus affecting PKA recruitment, have not been excluded.

Cell volume and osmolarity During their transit and maturation through the epididymis, spermatozoa acquire the ability to regulate cell volume, a very important process for the adequate development of motility. In fact, the osmolarity of the luminal fluid increases from the testis to the epididymis, and normal spermatozoa counteract shrinkage by increasing the uptake of organic osmolytes such as L-carnitine and amino

PHYSIOLOGY AND PATHOPHYSIOLOGY OF SPERM MOTILITY

PY20

anti+AKAP3

PY20+anti+AKAP3

Figure 2.3 Immunofluorescence analysis of fixed and permeabilized human spermatozoa. Confocal microscopy of double immunolabeling for tyrosine phosphorylated proteins ((b), PY20 antibody, green) and A kinase anchoring protein 3 (AKAP3) ((d), anti-AKAP3 antibody, red) reveals positivity for both antibodies in sperm tails. Simultaneous analysis of dual fluorescence confirms that tyrosine phosphorylation corresponds to AKAP3 in the tail ((f), double fluorescence, yellow). (a), (c), (e), negative controls without primary antibody. From reference 9, with permission. See also Color plate 2 on page xxvi

acids secreted by the epithelium76. Conversely, upon ejaculation, spermatozoa are subjected to the relatively hyposmotic environment of the female genital tract (osmotic pressure falls from 420 to 300 mmol/kg, from the epididymal cauda to the uterus76,77), and in order to prevent swelling, spermatozoa lose water and osmolytes acquired in the epididymis. Defects in such a delicate mechanism of volume regulation can cause an abnormal increase in sperm head volume and angulation of the sperm tail76, resulting in defects of sperm motility and fertility. A similar hairpin shape in the sperm tail and its detrimental consequence on motility has been demonstrated in both c-ros knock-out mice and following sperm treatment with the ion-channel blocker quinine78. Interestingly, seminal plasma osmolarity (intermediate between epididymis and uterus) is significantly higher in asthenozoospermic patients, irrespective of the cause of asthenozoospermia, than in normozoospermic men79. Moreover, seminal osmolarity correlates negatively with sperm progressive motility and kinetic characteristics80, suggesting a potential pathological role for seminal hyperosmolarity in the reduction of sperm motility in asthenozoospermic subjects. Sperm exposure to lowosmolarity media such as oviductal and uterine fluids activates an influx of Ca2+ through osmolarity-sensitive calcium channels79.

The role of fluid resorption in sperm maturation in the apical region of the epididymis has been extensively investigated81. Estrogens control differential expression of Na+/H+ exchangers82 and aquaporin channels83 through estrogen receptor α in the initial segment and caput of the epididymis. Aquaporin channels (e.g. AQ7) are also expressed in sperm tails and seem to be important for the control of cell volume, motility and fertility84. Therefore, it is conceivable that sperm maturation in the epididymis may be modulated by active water transport at two levels: the non-ciliated epidydimal epithelium and the sperm plasma membrane. L-carnitine, which is one of the main osmolytes captured by sperm during their transit through the epididymis, is essential for acyl transport in the mitochondrial β-oxidation of longchain fatty acids, and may also prevent sperm DNA and membrane damage induced by reactive oxygen species. Indeed, a positive effect of oral administration of carnitine in increasing semen quality, in particular sperm forward motility, in oligoasthenoteratozoospermic and asthenozoospermic patients has been demonstrated in clinical trials85,86.

Reactive oxygen species Reactive oxygen species (ROS), in particular hydrogen peroxide, produced either by spermatozoa

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or seminal leukocytes, have been described to affect different sperm functions including motility87. Their effects appear to depend on the concentration of ROS; low levels can induce the cAMP–PKA signaling cascade leading to an increase in sperm motility and tyrosine phosphorylation of proteins associated with capacitation, while high levels exert an inhibitory effect88,89. The detrimental action of ROS on sperm motility has been associated with increased lipid peroxidation of the plasma membrane90. High production of ROS as well as low antioxidant capacity may account for certain types of sperm pathology, in particular asthenozoospermia91. In such cases, the use of antioxidants may be indicated92,93. However, levels of glutathione-dependent seleno-enzymes in human spermatozoa, which are responsible for more general protection against ROS, have been reported to be similar in spermatozoa isolated from both normozoospermic and asthenozoospermic subjects94.

HYPERACTIVATED MOTILITY Hyperactivation is a special type of sperm motility developed in association with the process of capacitation in the female genital tract. It can also be achieved in vitro by seminal plasma removal and incubation of sperm in capacitating media95. It is characterized by a more energetic and less symmetric flagellar beat, which helps sperm to progress through the cervical mucus, the oviduct and, finally, the cumulus oophorus and zona pellucida surrounding the oocyte96–98. Furthermore, in species in which the oviductal isthmus represents a reservoir for spermatozoa, this particular swimming pattern seems to be important for the release of sperm entrapped in the folds and crypts of the oviductal epithelium98. In these cases, ovulation appears to induce a modification in the carbohydrate moieties of the oviductal epithelium, resulting in the release of fully activated sperm which have developed hyperactivation. This

phenomenon ensures appropriate timing for the acquisition of sperm fertilization potential99. The development of hyperactivation, especially at the oviducts, may be orchestrated by ovulation, since follicular fluid has been demonstrated to have a dose-dependent stimulatory effect on sperm hyperactivation100,101. The specific component capable of directly affecting sperm motility, however, has not yet been isolated102,103. The importance of adequate timing for hyperactivation has been demonstrated by the infertile t-haplotype mice, whose spermatozoa undergo premature hyperactivation in the female reproductive tract104. Interestingly, forward progressive motility and hyperactivation appear to be discontinuous and reversible processes, allowing sperm to switch alternately from one pattern to the other105. Capacitation and hyperactivation are two complementary aspects of sperm activation and develop simultaneously under physiological conditions. If capacitation is conceptualized as the complex of physiological changes enabling sperm to fertilize95, hyperactivation should be considered as part of such a process. However, they occur as independent pheomonena. In t-haplotype mice, spermatozoa show premature hyperactivation, but normal timing of capacitation in vitro. Although sharing similar signaling pathways, capacitation and hyperactivation are distinct processes that show different thresholds for activating factors. Indeed, the calcium and bicarbonate concentrations required for hyperactivation are far higher than those needed for capacitation16,97. The molecular bases underlying hyperactivation have been studied by different investigators, especially using a demembranated sperm model in which both plasma and mitochondrial membranes were removed by Triton X 100, leaving the axonemal structure intact and functional97. The development of hyperactivated and activated motility share the same signaling pathways and molecular players; however, different activation thresholds are involved. In particular, although ATP and cAMP are able to stimulate motility of

PHYSIOLOGY AND PATHOPHYSIOLOGY OF SPERM MOTILITY

demembranated spermatozoa, it is only following the addition of calcium that hyperactivation begins106, suggesting that this ion is a key regulator of the process97. Both external sources and intracellular stores are important for the increase in intracellular calcium levels associated with hyperactivation. Intracellular calcium stores showing inositol 1,4,5trisphosphate receptors (IP3R) have been demonstrated not only in the acrosome15, but also in the neck of the sperm16. In the distal region of the sperm neck, the axoneme associates with mitochondria and is surrounded by a redundant nuclear envelope, whose enlarged cisternae represent the flagellum intracellular calcium stores16. The release of calcium from this structure through IP3-gated channels seems to initiate sperm hyperactivation directly97,107, perhaps through the activation of calmodulin-dependent kinases. Calmodulin kinase II is one of the few discovered calcium targets in spermatozoa. Upon its activation by the calcium/calmodulin complex, it specifically stimulates hyperactivation20. Hyperactivation is also modulated by calcium entry through plasma membrane-specific channels such as voltage-gated, receptor-associated, storeoperated and cyclic nucleotide-gated channels (for reviews see references 11 and 12). A recently discovered family of sperm-specific voltage-operated calcium channels, the CatSper family, plays a pivotal role in the development and maintenance of sperm motility. The four members of the family are differentially expressed along the tail. While CatSper1 seems to regulate sperm-activated motility14, CatSper2 is important for hyperactivation. CatSper2 knock-out mice are infertile due to their inability to develop hyperactivation and penetrate the zona pellucida; however, capacitation, motility and the acrosome reaction are normal108. Interestingly, male infertility in a mutant CatSper2 family has recently been described109. Similar to activated motility, hyperactivation is regulated by a complex balance between kinase and phosphatase activity. Increased tyrosine phosphorylation of several sperm proteins in the tail

has been described to be associated with physiological59,66,69,110 and temperature-induced hyperactivation111. Inhibition of tyrosine and cAMPdependent kinases decreases hyperactivated motility55,69,112, whereas an increase in intracellular cAMP enhances this type of motility55,69,113.

CHEMOTAXIS AND SPERM MOTILITY Spermatozoa from invertebrates and mammals demonstrate attraction to chemoattractants secreted by the egg. This mechanism plays a pivotal role in guiding sperm towards the oocyte, which is particularly important for those species characterized by external fertilization. By binding to sperm-specific receptors, these molecules affect sperm motility, inducing a directed movement towards the chemical gradient of the chemoattractant (chemotaxis). In the sea urchin, speract secreted by the eggs induces, in a species-specific manner, a sperm chemotactic response by stimulating a transmembrane guanylate cyclase receptor complex associated with K+ channels preferentially localized along the flagellum, which results in an increase in intracellular cAMP and calcium114,115. In vitro induction of chemotaxis by follicular fluid (FF) has been extensively demonstrated in human sperm116. Progesterone117 and chemokines such as RANTES (T)118 have been suggested to be the active components of FF involved in sperm chemotaxis, even when the major effect of the steroid appears to be on sperm hyperactivation rather than on chemotaxis119. Furthermore, odorant-like molecules, through their specific olfactory receptors expressed on human spermatozoa, induce a membrane adenylate cyclasedependent increase in intracellular calcium, resulting in redirection of sperm along the ascending gradient of the odorant120,121. Sperm chemoattractants are secreted by the preovulatory follicle as well as the mature oocyte and its surrounding cumulus122, contributing to guiding sperm to the site of fertilization. However, the physiological role of chemotaxis in human spermatozoa is still

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controversial. Rather than being important guiding sperm toward the oocyte, chemotaxis humans seems more likely to be involved recruiting a selected, activated subpopulation spermatozoa123,124.

in in in of

COMPUTER-ASSISTED ASSESSMENT OF SPERM MOTILITY Classically, sperm motility has been assessed using phase-contrast microscopy, subjectively classifying sperm trajectories as forward progression (a and b), in situ (c) and immotile (d) according to the World Health Organization (WHO) Manual for the Examination of Human Semen and Sperm– Cervical Mucus Interaction (1999)125. The definition of asthenozoospermia is based on this classification, using 50% of forward-motile sperm as the normal cut-off. Computer-assisted analysis of sperm movement has significantly increased the objectivity of this assessment, providing a series of measurements such as sperm velocity, amplitude of head displacement and flagellar beat frequency, which otherwise could not be obtained with classical subjective microscopic evaluation. Furthermore, computer-assisted sperm analysis (CASA) systems are capable of sorting sperm subpopulations according to established threshold values, allowing for the quick and accurate determination of the percentage of spermatozoa displaying hyperactivated motility (for review see Mortimer 1997)126. The sensitivity and confidence of these instruments have greatly improved in the past few years, and they can now be referred to as potent research and clinical tools to measure both basic and hyperactivation parameters1. Essentially, CASA allows for the simultaneous evaluation of kinematic parameters in a high number of spermatozoa in a short period. All parameters are measured by CASA using the sperm head (centroid-derived movement) instead of the tail, as head movement passively reflects the flagellar beat and can be more easily followed due to its lower frequency of move-

ment. Velocity values are based on curvilinear velocity (VCL), straight-line velocity (VSL) and average path velocity (VAP). The VCL is referred to as the real distance that the sperm head covers during the observation time; the VAP is the distance that the sperm covers in the average direction of movement; and the VSL is the straight-line distance between the starting and the ending points of the sperm trajectory (Figure 2.4). More strictly associated with sperm head characteristics, lateral head displacement (ALH) and beat cross frequency (BCF) measure, respectively, the width of lateral movement and the number of times that the sperm head crosses the direction of movement. As indicated above, CASA systems can also derive from the obtained data in terms of a sort fraction, which represents the percentage of spermatozoa showing hyperactivation. The criteria for sorting hyperactivated sperm at 60 Hz can be manually set, and have been defined as VCL > 150 µm/s, ALHmax > 7.0 µm, linearity LIN < 50%127. Modern CASA instruments capture 60 images per second, which is ideal for properly characterizing sperm hyperactivated motility. To allow for unimpeded tridimensional sperm movement, motility should be analyzed in > 30-µm chambers, prewarmed to 37°C128.

BCF VAP

ALH

VCL B

VSL A

Figure 2.4 Schematic representation of a digitized sperm trajectory analyzed by a computer-assisted sperm analysis (CASA) system. VCL, curvilinear velocity; VSL, straight-line velocity; VAP, average path velocity; BCF, beat cross frequency; ALH, lateral head displacement; LIN, linearity = VSL/VSL; STR, straightness = VSL/VAP. From reference 9, with permission

PHYSIOLOGY AND PATHOPHYSIOLOGY OF SPERM MOTILITY

Besides its undisputed utility for research studies, CASA has also been widely adopted in the clinic. Several studies have correlated CASA parameters with assisted reproductive technologies (ART) outcomes129–131. Although no single parameter has shown good predictive value, some are valuable contributors to a multiparameter equation that predicts fertilization potential.

ETIOLOGY AND PATHOPHYSIOLOGY OF ASTHENOZOOSPERMIA Alterations in the previously described external and internal factors regulating sperm motion and metabolism in the flagellar structure may result in defects in sperm motility and infertility. A recent study reported that out of 1085 sperm samples analyzed from infertile subjects, 81% had defective motility, 20% of which presented pure asthenozoospermia132. Thus, asthenozoospermia is one of the main seminal pathologies underlying male infertility. Severe asthenozoospermia is frequently caused by flagellar alterations133. Ultrastructural studies of men with severe asthenozoospermia revealed two types of tail abnormalities: non-specific flagellar anomalies, which are random secondary alterations that affect variable numbers of spermatozoa in different samples, and dysplasia of the fibrous sheath (DFS), which is a systemic primary anomaly that affects most spermatozoa and is associated with respiratory pathology and familial incidence134,135. Non-specific flagellar anomalies constitute the most frequent flagellar pathology underlying asthenozoospermia. Its structural phenotype of random microtubular alterations is characteristically heterogeneous, and is sometimes associated with other andrological disorders (e.g. varicocele). Some of these patients respond to conservative treatment, while others require ART136. Dysplasia of the fibrous sheath is a different condition associated with extreme asthenozoospermia or total sperm immobility. It has a

homogeneous and distinctive phenotype characterized by distortions of the fibrous sheath and other axonemal and periaxonemal structures135,136. It has been postulated to be a variant of the immotile cilia syndrome, also known as primary ciliary dyskinesia, a congenital anomaly presenting with respiratory disease and male infertility. The axonemes of the respiratory cilia and sperm flagella show missing dynein arms, radial spokes and central microtubules, and general microtubular translocations137. In the Kartagener’s syndrome presentation, the ciliary/flagellar immotility is accompanied by dextrocardia. The familial clustering of these syndromes strongly suggests a genetic origin of the disease134. Intracytoplasmic sperm injection is the treatment of choice, but genetic counseling is required138. Not all cases of asthenozoospermia, especially those that are not severe in nature, are associated with structural anomalies of the flagellum, however. Our studies demonstrate that spermatozoa from less severe asthenozoospermic patients show a clear impairment in motility and their capacity to develop hyperactivation, which is associated with low membrane fluidity and a concomitant inability to undergo protein tyrosine phosphorylation59,60. This is particularly evident when spermatozoa are challenged with a capacitating incubation (e.g. 6 hours at 37°C, 5% CO2, in protein-supplemented medium) (Figure 2.5). Changes in membrane dynamics have been associated with tyrosine phosphorylation, as well as sperm function and fertilizing ability139,140. Spermatozoa from asthenozoospermic patients reveal significantly less fluid membranes before and after capacitation, in comparison with normozoospermic patients and proven-fertile donors60. Such a difference in membrane fluidity could be due to the increased susceptibility of these spermatozoa to suffer peroxidative damage91, as the generation of membrane lipid hydroperoxides has been associated with membrane fluidity reduction141,142. This susceptibility of asthenozoospermic sperm could be explained, in part, by their membrane composition, which is

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(a)

(b) Tyrosine phosphorylated sperm tails (%)

Hyperactivated cells (%)

18 16 14 12 10 8 6 4 2 0 Normozoospermic

Asthenozoospermic

Fertile donors

60 50 40 30 20 10 0 Normozoospermic

(c)

Asthenozoospermic

Fertile d donors

0.15

(d)

c A

0.13 d 0.11

220

0.09 97.4 0.07 66 0.05

T0 T0

Normozoospermic

Asthenozoospermic

Figure 2.5 Tyrosine phosphorylation, hyperactivation and membrane fluidity deficiencies in asthenozoospermic samples in comparison with samples from normozoospermic and proven-fertile men. Spermatozoa were incubated for 0 (T0, baseline) or 6 h under capacitating conditions and the incidence of hyperactivated motility (a), the incidence (immunofluorescence) and intensity (Western blot) of tyrosine phosphorylation (b) and (c) and the sperm membrane fluidity (fluorometry) (d) were determined. Asterisks (*) and letters (a vs. b, c vs. d, a vs. c and b vs. d) above bars indicate statistical significance. In the Western blot image (c), A = normozoospermic, B = asthenozoospermic, C = proven-fertile. In the membrane fluidity plot (d) GP is Laurdan’s general polarization. From reference 60, with permission

responsible for their reported higher oxidation coefficient91. Sperm membranes of asthenozoospermic samples contain high levels of polyunsaturated fatty acids, making them more prone to attack by reactive oxygen species. Since oxidizing conditions are normal during sperm capacitation and have been linked to signal transduction and tyrosine phosphorylation87,143, the predisposition of the asthenozoospermic samples to oxidative damage may be the origin of their membrane dysfunction, resulting in tyrosine

phosphorylation deficiency and alteration of motility.

TREATMENT OF ASTHENOZOOSPERMIA Systemic modalities Before considering any treatment, a correct diagnosis has to be established. Hence, the evaluation of subfertile men begins with a detailed history

PHYSIOLOGY AND PATHOPHYSIOLOGY OF SPERM MOTILITY

and physical examination. The history should identify the duration of attempted conception, intercourse timing and frequency, erectile function, ejaculation, life-style factors (alcohol, smoking, etc.) and any medications144. Other pertinent details include previous mumps orchitis, chemotherapy and/or radiation for cancer, cryptorchidism, previous reproductive tract infections, prior illnesses and any systemic disease. Physical examination should seek any sign of hypogonadism (virilization, body proportions, gynecomastia, etc.); a careful genitourinary examination should be performed to evaluate testicular size and consistency and the presence of masses and eventual penile pathology (hypospadias, etc.), and to identify the presence of the most common condition associated with male infertility – varicocele145,146. Severe ultrastructural sperm anomalies such as dysplasia of the fibrous sheath should also be ruled out. If a treatable condition responsible for male factor infertility, such as hypogonadism, varicocele, infections, immunologic infertility, obstructions and cryptorchidism, is found, then it should be corrected using current medical and/or surgical therapies147. Conversely, if a diagnosis of idiopathic asthenozoospermia is made, there are a few treatment options that have some degree of evidence-based support. Placebo-controlled double-blind randomized trials of men with idiopathic asthenozoospermia have demonstrated that L-carnitine and its analogs, especially L-acetyl-carnitine, after daily oral administration, increase sperm motility and kinematic parameters85,148,149. Although these studies do not have enough statistical power in themselves to draw unequivocal conclusions, they all show clear trends toward improvement of motility. This was especially notable in patients who started with the lower values of motility and motion parameters. Although the etiopathogenic mechanisms being modified by the oral administration of carnitines are not clearly established, increases in mitochondrial energy production and total

antioxidant capacity have been suggested85,86,149,150. Glutathione and coenzyme Q10 administration may also have beneficial effects in the treatment of idiopathic asthenozoospermia151,152. Another systemic treatment that has been tested in ART patients presenting with oligozoospermia or combined oligoasthenoteratozoospermia is pure follicle stimulating hormone (FSH). Although results are still controversial, several studies show an improvement of in vitro fertilization (IVF) outcome153–157.

Assisted reproductive modalities To date, albeit not curative, the most efficient treatment for asthenozoospermia is ART. Improving sperm motility in vitro before insemination is a common practice for moderate asthenozoospermic samples. Certain molecules have been demonstrated to be capable of improving sperm motility in vitro. Among them are inhibitors of phosphodiesterases such as pentoxifylline (PF), analogs of cAMP and a plasma membrane phospholipid, plateletactivating factor (PAF), which is physiologically produced and released by sperm158. PF is often used in ART to improve the fertilization rate and outcome in couples with male factor infertility159,160, since this compound not only stimulates motility in sperm obtained from asthenozoospermic subjects, but also positively affects sperm capacitation, binding to the zona pellucida and the acrosome reaction161. Sperm treatment with PF before IVF has been demonstrated not to be teratogenic for the developing embryo160; however, potential toxic effects cannot be definitively ruled out162. Furthermore, the presence of nonresponder subjects decreases the overall efficacy of the treatment163. The most striking negative sideeffect exerted by the majority of these compounds, including PF, is their ability also to stimulate the acrosome reaction102. Unfortunately, acrosomereacted spermatozoa are unable to bind the oocyte’s zona pellucida, thus decreasing their efficacy in conventional IVF.

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In this regard, during the past few years, our research has been focused on two molecules which seem very good candidates for potential adjuvant in vitro treatment of asthenozoospermia. LY294002 is a pharmacological inhibitor of phosphatidylinositol 3-kinase (PI3K), a kinase which phosphorylates in the 3-OH position, the inositol ring of the plasma membrane phosphoinositides164. This enzyme has been demonstrated to play a negative role in the control of sperm motility68,165–167, and its inhibition by LY294002 stimulates a significant increase in forward and rapid motility in both ejaculated and selected human spermatozoa, independently from the technique used for selection10,68,166.This stimulatory effect was more evident on samples from oligoasthenozoospermic compared with normozoospermic subjects10,165,166. In particular, direct addition of LY294002 to seminal samples of severe asthenozoospermic subjects increases the number of sperm showing forward motility recovered after a swim-up selection for ART166. PI3K inhibition by LY294002 stimulates tyrosine phosphorylation of AKAP3 in the fibrous sheath of sperm tails, allowing local recruitment and activation of PKA by increased binding of PKA regulatory subunit RIIβ to the phosphorylated form of AKAP310,68,167. PKA activation finally results in stimulation of sperm motility and hyperactivation68. Interestingly, in contrast to the above-mentioned molecules, LY294002 effects on sperm motility are not associated with an increase in the acrosome reaction165. Moreover, no toxic effect on embryo development has been demonstrated following sperm, oocyte or embryo treatment with LY294002 in a mouse model168. All these findings support the possible use of this drug as well as other PI3K inhibitors as potential tools to improve sperm motility in ART. In addition to the use of this pharmacological tool, our group (University of Florence) has also focused its attention on a physiological stimulus of sperm motility, the bicarbonate ion (HCO3–). We have recently demonstrated that in swimup-selected human spermatozoa, physiological

concentrations of bicarbonate (15 and 75 mmol/l) rapidly stimulate an increase in intracellular cAMP levels and tyrosine phosphorylation of AKAP3, the latter phenomenon resulting in an increased amount of PKA bound to this scaffolding protein, in a manner resembling LY294002 effects68,69. The stimulatory effects of bicarbonate on both sperm motility and AKAP3 phosphorylation seem to involve entry of the ion into the cell and activation of sAC, since they are inhibited by 4,4′-diisothiocyanostilbene-2,2′-disulfonic acid, a specific blocker of bicarbonate transporter, and by 2OH-estradiol, a selective inhibitor of sAC69. Thus, our findings strongly suggest that both HCO3– and LY294002 increase sperm motility by converging on the same signaling pathway involving stimulation of cAMP production by sAC and tyrosine phosphorylation of AKAP3 in the sperm tail. Redundancy of the signaling pathways leading to AKAP3 phosphorylation further highlights the importance of this process in regulating sperm motility. Molecules acting in promoting phosphorylation could potentially be used for increasing the number of motile spermatozoa selected for ART, offering infertile couples better chances for less invasive and expensive techniques.

CONCLUSIONS AND FUTURE DIRECTIONS Although progress has been significant, much remains to be elucidated concerning the biochemical pathways that regulate and maintain sperm motility. In particular, it is still unclear how spermatozoa begin to move following their release from the testis and their transit through the epididymis, and which signals are necessary for such activation. The identification of molecules involved in controlling sperm motility appears difficult, however. Genetic studies in mice show that many genes are involved in the development and maintenance of sperm motility. Some of them are testis-specific genes belonging to the fibrous sheath of the principal piece. Clarifying the

PHYSIOLOGY AND PATHOPHYSIOLOGY OF SPERM MOTILITY

molecular mechanisms involved in the onset of sperm motility will be of great benefit for the development of possible therapeutic strategies. Indeed, although some systemic therapies (such as oral administration of carnitine and antioxidants) have proved to be relatively efficacious, at present in vitro treatments remain the best option for the treatment of asthenozoospermia.

ACKNOWLEDGMENTS GF Doncel wishes to thank CONRAD and the US Agency for International Development for supporting his work on sperm motility and immobilizing agents. The views expressed in this manuscript do not necessarily represent those of the funding agencies or their programs. The authors also wish to express gratitude to Ms Charlotte Neumann for her editorial assistance.

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3′,5′-monophosphates and calcium. Biol Reprod 1997; 56: 1450 Leclerc P, de Lamirande E, Gagnon C. Cyclic adenosine 3′,5′monophosphate-dependent regulation of protein tyrosine phosphorylation in relation to human sperm capacitation and motility. Biol Reprod 1996; 55: 684 Patil SB, et al. Reactivation of motility of demembranated hamster spermatozoa: role of protein tyrosine kinase and protein phosphatases. Andrologia 2002; 34: 74 Si Y, Okuno M. Role of tyrosine phosphorylation of flagellar proteins in hamster sperm hyperactivation. Biol Reprod 1999; 61: 240 Visconti PE. Regulation, localization, and anchoring of protein kinase A subunits during mouse sperm capacitation. Dev Biol 1997; 192: 351 Luconi M, et al. Increased phosphorylation of AKAP by inhibition of phosphatidylinositol 3kinase enhances human sperm motility through tail recruitment of protein kinase A. J Cell Sci 2004; 117: 1235 Luconi M, et al. Tyrosine phosphorylation of the A kinase anchoring protein 3 (AKAP3) and soluble adenylate cyclase are involved in the increase of human sperm motility by bicarbonate. Biol Reprod 2005; 72: 22 Galantino-Homer HL, Visconti PE, Kopf GS. Regulation of protein tyrosine phosphorylation during bovine sperm capacitation by a cyclic adenosine 3′5′-monophosphate-dependent pathway. Biol Reprod 1997; 56: 707 Visconti PE, et al. Capacitation of mouse spermatozoa. II. Protein tyrosine phosphorylation and capacitation are regulated by a cAMP-dependent pathway. Development 1995; 121: 1139 O’Flaherty C, de Lamirande E, Gagnon C. Phosphorylation of the arginine-X-X-(serine/threonine) motif in human sperm proteins during capacitation modulation and protein kinase A dependency. Mol Hum Reprod 2004; 10: 355 Miki K, et al. Targeted disruption of the Akap4 gene causes defects in sperm flagellum and motility. Dev Biol 2002; 248: 331 Turner RM, et al. Molecular genetic analysis of two human sperm fibrous sheath proteins, AKAP4 and AKAP3, in men with dysplasia of the fibrous sheath. J Androl 2001; 22: 3025 Baccetti B, et al. Gene deletions in an infertile man with sperm fibrous sheath dysplasia. Hum Reprod 2005; 20: 2790

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76. Yeung CH, Sonnenberg-Riethmacher E, Cooper TG. Infertile spermatozoa of c-ros tyrosine kinase receptor knockout mice show flagellar angulation and maturational defects in cell volume regulatory mechanisms. Biol Reprod 1999; 61: 1062 77. Yeung CH, et al. The cause of infertility of male cros tyrosine kinase receptor knockout mice. Biol Reprod 2000; 63: 612 78. Yeung CH, et al. Sperm volume regulation: maturational changes in fertile and infertile transgenic mice and association with kinematics and tail angulation. Biol Reprod 2002; 67: 269 79. Rossato M, et al. Role of seminal osmolarity in the reduction of human sperm motility. Int J Androl 2002; 25: 230 80. Rossato M, Di Virgilio F, Foresta C. Involvement of osmo-sensitive calcium influx in human sperm activation. Mol Hum Reprod 1996; 903 81. O’Donnell L, et al. Estrogen and spermatogenesis. Endocr Rev 2001; 22: 289 82. Zhou Q, et al. Estrogen action and male fertility: roles of the sodium/hydrogen exchanger-3 and fluid reabsorption in reproductive tract function. Proc Natl Acad Sci USA 2001; 98: 14132 83. Oliveira CA, et al. Aquaporin-1 and -9 are differentially regulated by oestrogen in the efferent ductule epithelium and initial segment of the epididymis. Biol Cell 2005; 97: 385 84. Saito K, et al. Localization of aquaporin-7 in human testis and ejaculated sperm: possible involvement in maintenance of sperm quality. J Urol 2004; 172: 2073 85. Lenzi A, et al. Use of carnitine therapy in selected cases of male factor infertility: a double-blind crossover trial. Fertil Steril 2003; 79: 292 86. Garolla A, et al. Oral carnitine supplementation increases sperm motility in asthenozoospermic men with normal sperm phospholipid hydroperoxide, glutathione peroxidase levels. Fertil Steril 2005; 83: 355 87. Aitken RJ, et al. A novel signal transduction cascade in capacitating human spermatozoa characterised by a redox-regulated, cAMP-mediated induction of tyrosine phosphorylation. J Cell Sci 1998; 111: 645 88. Aitken RJ, Sawyer D. The human spermatozoon – not waving but drowning. Adv Exp Med Biol 2003; 518: 85 89. Ford WC. Regulation of sperm function by reactive oxygen species. Hum Reprod Update 2004; 10: 387 90. Williams AC, Ford WC. Relationship between reactive oxygen species production and lipid

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peroxidation in human sperm suspensions and their association with sperm function. Fertil Steril 2005; 83: 929 Calamera J, et al. Superoxide dismutase content and fatty acid composition in subsets of human spermatozoa from normozoospermic, asthenozoospermic, and polyzoospermic semen samples. Mol Reprod Dev 2003; 66: 422 Martin-Du Pan RC, Sakkas D. Is antioxidant therapy a promising strategy to improve human reproduction? Are anti-oxidants useful in the treatment of male infertility? Hum Reprod 1998; 13: 2984 Lenzi A, Gandini L, Picardo M. A rationale for glutathione therapy. Hum Reprod 1998; 13: 1419 Tramer F, et al. Native specific activity of glutathione peroxidase (GPx-1), phospholipid hydroperoxide glutathione peroxidase (PHGPx) and glutathione reductase (GR) does not differ between normo- and hypomotile human sperm samples. Int J Androl 2004; 27: 88 Yanagimachi R. Mammalian fertilization. In Knobil E, Neill JD, eds. The Physiology of Reproduction. New York: Raven Press, 1994: 189 Suarez SS, Dai X. Hyperactivation enhances mouse sperm capacity for penetrating viscoelastic media. Biol Reprod 1992; 46: 686 Ho HC, Suarezs SS. Hyperactivation of mammalian spermatozoa: function and regulation. Reproduction 2001; 122: 519 Demott RP, Suarez SS. Hyperactivated sperm progress in the mouse oviduct. Biol Reprod 1992; 46: 779 Gualtieri R, Talevi R. Selection of highly fertilization-competent bovine spermatozoa through adhesion to the Fallopian tube epithelium in vitro. Reproduction 2003; 125: 251 Fabbri R, et al. Follicular fluid and human granulosa cell cultures: influence on sperm kinetic parameters, hyperactivation, and acrosome reaction. Fertil Steril 1998; 69: 112 Yao Y, Ho P, Yeung WS. Human oviductal cells produce a factor(s) that maintains the motility of human spermatozoa in vitro. Fertil Steril 2000; 73: 479 Kay VJ, Coutts JR, Robertson L. Effects of pentoxifylline and progesterone on human sperm capacitation and acrosome reaction. Hum Reprod 1994; 9: 2318 Sueldo CE et al. Effect of progesterone on human zona pellucida sperm binding and oocyte penetrating capacity. Fertil Steril 1993; 60: 137

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104. Olds-Clarke P, Johnson LR. t haplotypes in the mouse compromise sperm flagellar function. Dev Biol 1993; 155: 14 105. Tessler S, Olds-Clarke P. Linear and nonlinear mouse sperm motility patterns. A quantitative classification. J Androl 1985; 6: 35 106. Lindemann CB, Goltz JS. Calcium regulation of flagellar curvature and swimming pattern in triton X-100-extracted rat sperm. Cell Motil Cytoskeleton 1988; 10: 420 107. Ho HC, Suarez SS. Characterization of the intracellular calcium store at the base of the sperm flagellum that regulates hyperactivated motility. Biol Reprod 2003; 68: 1590 108. Quill TA, et al. A voltage-gated ion channel expressed specifically in spermatozoa. Proc Natl Acad Sci USA 2001; 98: 12527 109. Avidan N, et al. CATSPER2, a human autosomal nonsyndromic male infertility gene. Eur J Hum Genet 2003; 11: 497 110. Nassar A, et al. Modulation of sperm tail protein tyrosine phosphorylation by pentoxifylline and its correlation with hyperactivated motility. Fertil Steril 1999; 71: 919 111. Si Y. Hyperactivation of hamster sperm motility by temperature-dependent tyrosine phosphorylation of an 80-kDa protein. Biol Reprod 1999; 61: 247 112. Bajpai M, Asin S, Doncel GF. Effect of tyrosine kinase inhibitors on tyrosine phosphorylation and motility parameters in human sperm. Arch Androl 2003; 49: 229 113. Yunes R, et al. Cyclic nucleotide phosphodiesterase inhibition increases tyrosine phosphorylation and hypermotility in normal and pathological human spermatozoa. Biocell 2005; 29: 287 114. Cook SP, et al. Sperm chemotaxis: egg peptides control cytosolic calcium to regulate flagellar responses. Dev Biol 1994; 165: 10 115. Wood CD, et al. Real-time analysis of the role of Ca(2+) in flagellar movement and motility in single sea urchin sperm. J Cell Biol 2005; 169: 725 116. Villanueva-Diaz C, et al. Evidence that human follicular fluid contains a chemoattractant for spermatozoa. Fertil Steril 1990; 54: 1180 117. Villanueva-Diaz C, et al. Progesterone induces human sperm chemotaxis. Fertil Steril 1995; 64: 1183 118. Isobe T, et al. The effect of RANTES on human sperm chemotaxis. Hum Reprod 2002; 17: 1441 119. Jaiswal BS, et al. Human sperm chemotaxis: is progesterone a chemoattractant? Biol Reprod 1999; 60: 1314

120. Spehr M, et al. Identification of a testicular odorant receptor mediating human sperm chemotaxis. Science 2003; 299: 2054 121. Spehr M, et al. Particulate adenylate cyclase plays a key role in human sperm olfactory receptor-mediated chemotaxis. J Biol Chem 2004; 279: 40194 122. Sun F, et al. Human sperm chemotaxis: both the oocyte and its surrounding cumulus cells secrete sperm chemoattractants. Hum Reprod 2005; 20: 761 123. Eisenbach M, Ralt D. Precontact mammalian sperm–egg communication and role in fertilization. Am J Physiol 1992; 262: C1095 124. Cohen-Dayag A, et al. Sperm capacitation in humans is transient and correlates with chemotactic responsiveness to follicular factors. Proc Natl Acad Sci USA 1995; 92: 11039 125. World Health Organization. WHO Laboratory Manual for the Examination of Human Semen and Sperm–Cervical Mucus Interaction. Cambridge: Cambridge University Press, 1999 126. Mortimer ST. A critical review of the physiological importance and analysis of sperm movement in mammals. Hum Reprod Update 1997; 3: 403 127. Mortimer ST, Swan MA, Mortimer D. Effect of seminal plasma on capacitation and hyperactivation in human spermatozoa. Hum Reprod 1998; 13: 2139 128. Mortimer ST, Swan MA. Variable kinematics of capacitating human spermatozoa. Hum Reprod 1995; 10: 3178 129. Paston MJ, et al. Computer-aided semen analysis variables as predictors of male fertility potential. Arch Androl 1994; 33: 93 130. Hirano Y, et al. Relationships between sperm motility characteristics assessed by the computer-aided sperm analysis (CASA) and fertilization rates in vitro. J Assist Reprod Genet 2001; 18: 213 131. Shibahara H, et al. Prediction of pregnancy by intrauterine insemination using CASA estimates and strict criteria in patients with male factor infertility. Int J Androl 2004; 27: 63 132. Curi SM, et al. Asthenozoospermia: analysis of a large population. Arch Androl 2003; 49: 343 133. Chemes H. The significance of flagellar pathology in the evaluation of asthenozoospermia. In Baccetti B, ed. Comparative Spermatology 20 years later. Serono Symposia Publications. New York: Raven Press, 1991; 75: 815 134. Chemes HE, et al. Ultrastructural pathology of the sperm flagellum: association between flagellar

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pathology and fertility prognosis in severely asthenozoospermic men. Hum Reprod 1998; 13: 2521 Chemes HE, et al. Dysplasia of the fibrous sheath: an ultrastructural defect of human spermatozoa associated with sperm immotility and primary sterility. Fertil Steril 1987; 48: 664 Chemes HE. Phenotypes of sperm pathology: genetic and acquired forms in infertile men. J Androl 2000; 21: 799 Afzelius BA, et al. Lack of dynein arms in immotile human spermatozoa. J Cell Biol 1975; 66: 2252 Olmedo SB, et al. Pregnancies established through intracytoplasmic sperm injection (ICSI) using spermatozoa with dysplasia of fibrous sheath. Asian J Androl 2000; 2: 1252 Gadella BM, et al. Dynamics in the membrane organization of the mammalian sperm cell and functionality in fertilization. Vet Q 1999; 21: 142 Flesch FM, et al. Bicarbonate stimulated phospholipid scrambling induces cholesterol redistribution and enables cholesterol depletion in the sperm plasma membrane. J Cell Sci 2001; 114: 3543 Aitken RJ, et al. Analysis of the responses of human spermatozoa to A23187 employing a novel technique for assessing the acrosome reaction. J Androl 1993; 14: 132 Windsor DP, White IG. Assessment of ram sperm mitochondrial function by quantitative determination of sperm rhodamine 123 accumulation. Mol Reprod Dev 1993; 36: 354 Aitken J, Fisher H. Reactive oxygen species generation and human spermatozoa: the balance of benefit and risk. Bioessays 1994; 16: 259 Isidori A, Latini M, Romanelli F. Treatment of male infertility. Contraception 2005; 72: 314 Redmon JB, Carey P, Pryor JL. Varicocele – the most common cause of male factor infertility? Hum Reprod Update 2002; 8: 53 Fretz PC, Sandlow JI. Varicocele: current concepts in pathophysiology, diagnosis, and treatment. Urol Clin North Am 2002; 29: 921 Liu PY, Handelsman DJ. The present and future state of hormonal treatment for male infertility. Hum Reprod Update 2003; 9: 9 Lenzi A, Gandini L. Characterization of human sperm. Hum Reprod 2002; 17: 842 Balercia G, et al. Placebo-controlled double-blind randomized trial on the use of L-carnitine, L-acetylcarnitine, or combined L-carnitine and L-acetylcarnitine in men with idiopathic asthenozoospermia. Fertil Steril 2005; 84: 662

150. Costa M, et al. L-carnitine in idiopathic asthenozoospermia: a multicenter study. Italian Study Group on Carnitine and Male Infertility. Andrologia 1994; 26: 155 151. Lenzi A, et al. Placebo-controlled, double-blind, cross-over trial of glutathione therapy in male infertility. Hum Reprod 1993; 8: 1657 152. Balercia G, et al. Coenzyme Q(10) supplementation in infertile men with idiopathic asthenozoospermia: an open, uncontrolled pilot study. Fertil Steril 2004; 81: 93 153. Acosta AA, Khalifa E, Oehninger S. Pure human follicle stimulating hormone has a role in the treatment of severe male infertility by assisted reproduction: Norfolk’s total experience. Hum Reprod 1992; 7: 1067 154. Ashkenazi J, et al. The role of purified follicle stimulating hormone therapy in the male partner before intracytoplasmic sperm injection. Fertil Steril 1999; 72: 670 155. Dirnfeld M, et al. Pure follicle-stimulating hormone as an adjuvant therapy for selected cases in male infertility during in-vitro fertilization is beneficial. Eur J Obstet Gynecol Reprod Biol 2000; 93: 105 156. Caroppo E, et al. Recombinant human folliclestimulating hormone as a pretreatment for idiopathic oligoasthenoteratozoospermic patients undergoing intracytoplasmic sperm injection. Fertil Steril 2003; 80: 1398 157. Foresta C, et al. Treatment of male idiopathic infertility with recombinant human follicle-stimulating hormone: a prospective, controlled, randomized clinical study. Fertil Steril 2005; 84: 654 158. Krausz C, et al. Effect of platelet-activating factor on motility and acrosome reaction of human spermatozoa. Hum Reprod 1994; 9: 471 159. Yovich JL. Pentoxifylline: actions and applications in assisted reproduction. Hum Reprod 1993; 8: 1786 160. Rizk B, et al. Successful use of pentoxifylline in male-factor infertility and previous failure of in vitro fertilization: a prospective randomized study. J Assist Reprod Genet 1995; 12: 710 161. Paul M, Sumpter JP, Lindsay KS. The paradoxical effects of pentoxifylline on the binding of spermatozoa to the human zona pellucida. Hum Reprod 1996; 11: 814 162. Centola GM, Cartie RJ, Cox C. Differential responses of human sperm to varying concentrations of pentoxyfylline with demonstration of toxicity. J Androl 1995; 16: 136

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163. Tournaye H, et al. Use of pentoxifylline in assisted reproductive technology. Hum Reprod 1995; 10 (Suppl 1): 72 164. Wymann MP, Pirola L. Structure and function of phosphoinositide 3-kinases. Biochim Biophys Acta 1998; 1436: 127 165. du Plessis SS, et al. Phosphatidylinositol 3-kinase inhibition enhances human sperm motility and sperm–zona pellucida binding. Int J Androl 2004; 27: 19

166. Luconi M, et al. Phosphatidylinositol 3-kinase inhibition enhances human sperm motility. Hum Reprod 2001; 16: 1931 167. Aparicio IM, et al. Inhibition of phosphatidylinositol 3-kinase modifies boar sperm motion parameters. Reproduction 2005; 129: 283 168. Luconi M, et al. Enhancement of mouse sperm motility by the PI3-kinase inhibitor LY294002 does not result in toxic effects on preimplantation embryo development. Hum Reprod 2005; 20: 3500

3 The pathophysiology and genetics of human male reproduction Christaan F Hoogendijk, Ralf Henkel

INTRODUCTION

result in malformed, dysfunctional male germ cells. Therefore, to understand the physiology of fertilization, the understanding of spermatogenesis and its morphological and genetic processes is of paramount importance.

The male germ cells, the spermatozoa, are produced in a unique process named spermatogenesis. During this process, spermatogenic stem cells undergo reduction of the genome from diploid cells to haploid cells, as well as unequaled morphological and functional changes. In this respect, spermatozoa are not only the smallest (length of sperm head: 4–5 µm) and most polarized cells (sperm head in front, flagellum at rear) in the body, but also the only cells that fulfill their function outside the body, even in a different individual, the female reproductive tract. Therefore, spermatozoa are highly specialized cells, simply a ‘means of transportation’, that transfer the genetic information from the male to the female, the oocyte for which specific physiological functions of these cells are required. For the sperm cells to acquire these functions, morphological and physiological development of the spermatozoa has to take place. In addition, proper chromosomal and genetic constitution is mandatory, i.e. chromosomal and DNA integrity must be given. During spermatogenesis, spermatozoa acquire the morphological and physiological foundations, which eventually have to mature during epididymal maturation, for normal sperm function. This means that if the processes taking place in the course of spermatogenesis are defective, this will

GENETIC CONTROL OF SPERMATOGENESIS The relationship between structurally abnormal and genetically defective spermatozoa poses a crucial unknown. The long sequence of events involved in spermatogenesis, from germ cell differentiation to functionally mature spermatozoa, is fraught with the possibility of both structural and genetic damage. Spermatogenesis consists of three distinct phases: (1) proliferation and differentiation of diploid spermatogonial stem cells, (2) meiosis where chromosome pairing and genetic recombination occurs and (3) spermiogenesis, a unique series of events in which the rather commonplace-appearing, albeit haploid, round spermatids differentiate into species-specific-shaped spermatozoa. Collectively, these intervals consist of many developmental events, which offer numerous opportunities for the introduction of damage into the genome of the male gamete. These concerns are exacerbated by the ability of scientists and embryologists to use differentiating 35

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male germ cells, prior to the completion of spermatogenesis, for fertilization. This raises the question: are we not introducing ‘incomplete’ male gametes into oocytes?

Spermatogonial differentiation The intricate mechanisms whereby stem cells maintain a population of proliferating and differentiating cells are only beginning to be unraveled1. In the mammalian testis, spermatogonial type A stem cells proliferate, producing three classes of spermatogonia: (1) a group of presumably identical spermatogonial stem cells, (2) a population of differentiating spermatogonia and (3) a large number of cells that undergo cell death by apoptosis2. The originators of this developmental cascade, type A spermatogonia, represent a mixed population of cell types designated type A0, A1, A2, A3 or A4 spermatogonia. Among these cells, the identity of ‘true’ stem cells is yet to be definitively established. Although multiple stem-cell renewal models have been put forward, one commonly accepted model proposes that type A0 spermatogonia represents a reserve population of stem cells, which divide slowly and can repopulate the testis after damage3. Thus, types A1–A4 spermatogonia are believed to be the renewing stem-cell spermatogonia, and these cells maintain the fertility of a man. Type A spermatogonia differentiate into intermediate and type B spermatogonia, which in turn divide and enter the differentiating pathway leading to spermatozoa. These cellular programming events appear to be irreversible, because once committed to differentiation, the spermatogonia appear incapable of re-entering the pathway that produces stem cells. The implications of genetically defective spermatogonia are substantial, since it is these cells that will function as the precursors of spermatozoa throughout the life of the individual. The large number of spermatogonial stem cells that undergo apoptosis suggests that a sophisticated monitoring system has evolved in which ‘defective’ stem cells are removed. Currently,

much effort is being directed towards studies defining mechanisms of apoptosis in somatic cells. Research efforts need to be extended to define the mechanisms by which specific populations of stem cells are selected to be targets for cell death. Specifically in the testis, an understanding of how the differentiating germ cells are continually being assessed, presumably by a self-monitoring system, will help greatly to minimize the production of genetically defective germ cells.

Meiosis Meiosis represents a fascinating interval of spermatogenesis in which genetic alterations, including genetic damage, are intentionally introduced into the genome, which in turn contributes to the evolutionary change of species. In addition to its essential role in producing haploid gametes from diploid stem cells, the extended interval of meiotic prophase has evolved to provide the critical cellular milieu for precise genetic recombination4. Meiotic prophase commences with preleptotene primary spermatocytes, the cell type in which the last semiconservative DNA replication of the male germ cell occurs. All subsequent DNA synthesis in differentiating male germ cells represents DNA repair synthesis. Chromosome condensation initiates concomitantly with the movement of leptotene and zygotene spermatocytes to the adluminal compartment of the seminiferous tubule from the basal membrane region. Alignment and complete pairing of the chromosomal homologs are completed in pachytene spermatocytes. As the chromosomes condense, axial elements appear between the two sister chromatids of each chromosomal homolog. The addition of a visible central element to the chromosomes produces the synaptonemal complex, a highly conserved structure in the meiotic cells of organisms ranging from water mold to the human, that is needed for effective synapsis. Because synapsis of chromosomes represents an event unique and critical to genetic recombination, meiotic cells contain many novel structural proteins and enzymes

PATHOPHYSIOLOGY AND GENETICS OF HUMAN MALE REPRODUCTION

needed for chromosome and DNA alignment, DNA breakage, recombination and DNA repair. Among the proteins recently shown to be important in the genetic recombination process are Rad 51, a human homolog of a bacterial recombination protein5; BRCA1, a tumorsuppressor gene implicated in familial breast and ovarian cancers5; ATM-related genes, members of a gene family proposed to prevent DNA damage6–8; a ubiquitin-conjugating repair enzyme believed to be involved in protein turnover9,10; a mammalian homolog to a meiosis-specific DNA double-strand breaking enzyme11; DNA recombination genes12,13; and a meiotic-specific heat shock protein14. Since meiosis is crucial for the survival of a species, an elaborate series of safeguards has evolved to pair, break and repair chromosomal DNA. Despite such regulatory mechanisms, it is well known that translocations and aneuploidy are regularly introduced during the meiotic divisions. Moreover, in a sizeable population of infertile men, germ cell differentiation arrests during meiosis15. Anomalies in pairing and chromosome segregation are likely to contribute to this population of infertile men. Moreover, the many specific molecular processes essential for meiosis provide many targets for both genetic damage and for the introduction of structural defects, leading to the arrest of germ cell development. Our rapidly advancing knowledge of the mechanisms of meiosis in both males and females will provide substantial insights into a significant cause of male infertility.

Spermiogenesis Spermiogenesis represents an interval of spermatogenesis that appears exceptionally susceptible to the introduction of both genetic and structural defects in the maturing male gamete, as the round spermatid is transformed into the highly elongated and polarized (sperm head in front, flagellum at rear) spermatozoon at a time of reduced repair capabilities. Moreover, during spermiogenesis, a major reorganization of the cell occurs. The

nucleus elongates and an acrosome containing a group of proteolytic enzymes develops. At the chromosomal level, the histones, the predominant chromatin proteins of somatic cells, are replaced by the highly basic transition proteins, which in turn are replaced by the protamines, producing a tightly compacted nucleus with extensive disulfide bridge crosslinking. In fact, sperm chromatin condensation during spermiogenesis results in DNA taking up about 90% of the total volume of the sperm nucleus. In contrast, in normal somatic cells, the DNA takes up only 5% of the nucleus volume, while in mitotic chromosomes DNA takes up about 15% of the nuclear volume16. Displacement of the histones from the nucleosomes during spermiogenesis may leave the DNA of the haploid genome especially susceptible to damage at a time of limited repair capabilities. Although unscheduled DNA repair has been demonstrated to occur in early stages of spermatid development17, as spermiogenesis proceeds, unscheduled DNA synthesis diminishes, and it is not known whether any of the sophisticated DNA repair mechanisms that function during meiosis are still operational. In addition to the major nuclear restructuring taking place during spermiogenesis, the axoneme and tail of the developing male germ cell are produced, requiring synthesis of many structural proteins, including those of the fibrous sheath18 and the outer dense fiber proteins19. These cellular changes require extensive gene expression from the actively transcribed haploid genome before it matures into a genetically quiescent nucleus. In fact, transcription of RNA ceases during mid-spermiogenesis20, and translational regulation plays a prominent regulatory role in the extensive protein synthesis throughout the latter half of spermiogenesis that is required to produce spermatozoa21–23. The major reorganizational events of the differentiating spermatid are accompanied by significant alterations in the energy suppliers of the cell, the mitochondria. Mitochondria exhibit several distinct morphologies as germ cell differentiation proceeds24. Spermatogonia and somatic testicular

MALE INFERTILITY

cells contain the ‘cigar-shaped’ mitochondria found in most somatic tissues. During meiosis, mitochondria with diffuse and vacuolated matrices start replacing the ‘somatic’ mitochondria. By the beginning of spermiogenesis, the ‘somatic’ mitochondria have been totally replaced by ‘germ cell’ mitochondria, which in turn are replaced by the crescent-shaped mitochondria of spermatozoa. These structural changes in mitochondria are accompanied by major changes in protein composition25,26. Although spermatocytes and spermatids are estimated to contain over 103 mitochondria, each spermatozoa midpiece contains only approximately 75 uniquely helically shaped mitochondria. This requires the reduction or possibly selection of mitochondria as the germ cells differentiate27. At the conclusion of spermiogenesis, most of the cytoplasm of the elongated spermatid is removed as the residual body is pinched off, leaving spermatozoa with little cytoplasm, and no cytoplasmic ribosomes. Although cytoplasmic protein synthesis does not occur in spermatozoa, cytoplasmic mitochondrial protein synthesis continues28. Considering the massive changes that occur during spermiogenesis, it is not surprising that many cases of germ cell blockage during spermiogenesis lead to infertility in men. Defects in the synthesis of the midpiece, axoneme, mitochondria or tail assembly would result in structurally abnormal spermatozoa often with poor motility, while mutations in proteins needed for the compaction of sperm nuclei or sperm head shaping would lead to spermatozoa with abnormal heads. Despite the presence of aberrant-appearing spermatozoa, it is premature to equate morphological aberrations with genetic aberrations. More disquieting, minor base-pair substitutions in critical genes that would not alter spermatozoon morphology would lead to genetically defective but normal-appearing spermatozoa! Our inability to detect genetically defective male gametes is of great concern when round spermatid nucleus injections (ROSNI) and round spermatid injections (ROSI) are used to overcome

the sterility of men incapable of completing spermatogenesis29,30. The success of ROSNI and ROSI has demonstrated that although spermiogenesis is essential for reorganization of the male germ cell to become a motile cell, it is not needed for fertilization. Thus, the normal physiological selection processes leading to fertilization can be bypassed in mice and men. Unfortunately, morphological examination of the spermatids tells little of any underlying genetic defects in the spermatid chosen for injection. A major research effort must be undertaken with a mammalian model system such as the mouse in which a large population of progeny produced by ROSNI and ROSI are produced and evaluated. Among the concerns raised by these procedures is whether we are circumventing gene imprinting in the male genome. The detection of DNA methylation of spermatozoa in the epididymis also raises questions31. Without a detailed analysis of this approach in an animal system, we could be facing major genetic dangers introduced by the ROSNI and ROSI technologies.

GENETICS OF THE SPERMATOZOON During the past few years, many exciting discoveries, previously unsuspected by scientists, have been made about the structure and function of sperm DNA. For example, the paternal genome has been shown to contain endogenous nicks, probably as a normal part of spermiogenesis32. In patients in whom these nicks are left unrepaired during the final stages of spermiogenesis, fertility is decreased33. Topoisomerases, the enzymes thought to be responsible for these nicks, are present throughout spermatogenesis; nonetheless, they are not present in spermatozoa34,35. Evenson and colleagues36–38 developed the sperm chromatin structure assay (SCSA) that assesses the potential of sperm DNA to denature under certain conditions. This potential also correlates with reduced fertility39. Perhaps most surprising of all, evidence published by Spadafora and

PATHOPHYSIOLOGY AND GENETICS OF HUMAN MALE REPRODUCTION

colleagues40,41 shows that fully mature mouse spermatozoa have the potential to incorporate exogenous DNA sequences into the paternal genome. Finally, since a sheep has been cloned from an adult cell42, and this technique has also been successful in several other mammalian species such as cattle, the mouse, goat, pig, cat and rabbit and even a primate, the rhesus monkey, this has raised the question of the importance of the paternal genome and its unique structure in embryogenesis. The above-mentioned discoveries have forced us to rethink the idea of sperm DNA structure in which we visualize the paternal genome as being so tightly packaged into an almost crystalline state that it is virtually inert until it is unfolded during fertilization. The sperm chromosome structure is, in fact, very complex – some attributes are similar to somatic cell DNA organization, and others are unique to spermatogenic cells. When discussing sperm chromatin packaging, several different aspects of structure need to be addressed. These can be divided into different levels of complexity based on the length of DNA being discussed. Each chromosome consists of one double-stranded DNA molecule, containing telomeric repeats at both ends, and centromeric repeats somewhere along its length. Chromosomes are in the order of several million base pairs in length, and, when fully decondensed, are each many times longer than the sperm nucleus itself. At the other end of the spectrum are spermspecific protamines, each of which bind to only a few base pairs of DNA. Sperm DNA packaging can be subdivided into four levels. In the following paragraphs, we discuss the structural relationship between the different levels of DNA packaging in the mature sperm nucleus.

Level I: chromosomal anchoring by the nuclear annulus In the first step of the assembly of sperm chromatin, the two strands of naked DNA that make

up the chromosomes are attached to a spermspecific structure, the nuclear annulus. This represents a novel type of DNA organization, termed chromosomal anchoring, that is found only in spermatogenic cells. Spermatozoa that are washed with non-ionic detergents such as NP-40, and then treated with high salt and reducing agents to extract the protamines, will decondense completely, leaving no trace of nuclear structure. The DNA, however, remains anchored to the base of the tail, so that the sperm chromatin resembles a broom, with the tail acting as the handle43. Since this chromosomal anchoring is maintained in sperm nuclei from which protamines have been extracted, it is independent of protamine binding. Ward and Coffey43 have isolated a small structure that is located at the implantation fossa in hamster spermatozoa, which they have termed the nuclear annulus, to which the DNA is attached in these decondensed nuclei. The nuclear annulus is shaped like a bent ring, and is about 2 µm in length. It is found only in sperm nuclei, although it is currently unknown at what stage of spermiogenesis it is first formed. Thus, so far, no evidence for a nuclear annulus-like structure in any somatic cell type has been found. In contrast, there is evidence of its existence in hamster43, human44, mouse and Xenopus sperm nuclei. Its existence in a wide variety of species suggests a fundamental role in sperm function. Unique DNA sequences were found to be associated with the nuclear annulus. Ward and Coffey43 termed these sequences NA-DNA. The existence of these unique sequences suggests that the nuclear annulus anchors chromosomes according to particular sequences and not by random DNA binding. They also hypothesized that NA-DNAs on different chromosomes become associated early during spermiogenesis, to initiate chromatin condensation by aggregating specific sites of each chromosome to one point. This hypothesis is supported by the work of Zalensky et al.45, who suggested that sperm chromosomes are packaged as extended fibers along the length of the nucleus. Each chromosome so far

MALE INFERTILITY

examined has only one site at the base of the nucleus, where the nuclear annulus is located. By organizing the chromosomes so that the NADNA sites of each chromosome are aggregated onto one structure, the nuclear annulus may also affect the determination of sperm nuclear shape. For example, in the hamster spermatozoon the longer chromosomes may extend into the thinner hook of the nucleus, while a portion of every chromosome is located at the nuclear annulus. This is supported by image analysis of the distribution of DNA throughout the hamster sperm nucleus, which demonstrates that the highest concentration of DNA in the packaged sperm nucleus is at the base, where the nuclear annulus is located; in contrast, the lowest concentration of DNA is in the hooked portion46. The hypothesis is further supported by electron microscopic evidence that chromatin near the implantation fossa is one of the first areas to condense during spermiogenesis47. Thus, the nuclear annulus may represent the only known aspect of sperm chromatin condensation that is specific for individual chromosome sites.

Level II: sperm DNA loop domain organization DNA loop domain organization

At this level, anchored chromosomes are organized into DNA loop domains. Parts of the nuclear matrix, protein structural fibers, attach to the DNA every 30–50 kb by specific sequences termed matrix attachment regions (MARs). This arranges the chromosome strands into a series of loops. This type of organization can be visualized experimentally in preparations, and is known as a nuclear halo. A nuclear halo comprises the nuclear matrix with a halo of DNA surrounding it. This halo consists of loops of naked DNA, 25–100 kb in length, attached at their bases to the matrix. Each loop domain visible in the nuclear halo consists of a structural unit of chromatin that exists in vivo in a condensed form.

As with chromosomal anchoring, DNA loop domain formation is independent of protamine binding. The organization of DNA into loop domains is the only type of structural organization resolved thus far that is present in both somatic and sperm cells. In somatic cells, DNA is coiled into nucleosomes, then further coiled into a 30-nm solenoid-like fiber and then organized into DNA loop domains. The corresponding structures in sperm chromatin have a very different appearance. Protamine binding causes a different type of coiling, and DNA is folded into densely packed toroids, but still organized into loop domains. Mammalian sperm nuclei contain a small amount of histones, which are presumably organized into nucleosomes48,49, but most of the DNA is reorganized by protamines. This means that with the evolutionary pressure to condense sperm DNA, all aspects of chromatin structure are sacrificed other than the organization of DNA into loop domains. This suggests that DNA loop domains play a crucial role in sperm DNA function. DNA loop domain function

In somatic cells, DNA loop domain organization has been implicated in both the control of gene expression and in DNA replication. Each DNA loop domain replicates at a fixed site on the nuclear matrix, by being reeled through the enzymatic machinery located at the base of the loop50,51. DNA replication origins have been localized to the nuclear matrix in mammals52, and the varying sizes of replicons in different species have been correlated with the sizes of loop domains53. A replicon can be thought of as the distance between two regions of replication. The attachment sites of individual genes to the nuclear matrix vary between cell types, and are also involved in transcription. Active genes are tightly associated with the nuclear matrix, but inactive genes are usually located within the extended part of the DNA loop54–58. In this manner, the threedimensional organization of DNA plays an important role in DNA function.

PATHOPHYSIOLOGY AND GENETICS OF HUMAN MALE REPRODUCTION

Possible function of sperm DNA loop domains

It has been demonstrated that the specific configurations of DNA loop domains are markedly different in sperm and somatic cells44,59. In somatic cells, DNA replication and transcription are the major functions in which DNA loop domain structures are involved48–53,56–58. However, since mature sperm nuclei perform neither process60, it is not clear what is the function of sperm DNA loop domain organization. Two possibilities exist. First, the DNA loop domain structures in spermatozoa may be residual structures that were required for transcription or DNA replication that occurred during spermatogenesis. Second, they may be involved in regulating these functions during embryonic development, if the embryo inherits them. If, for example, paternal genes in the male pronucleus of a newly fertilized egg were organized into the same DNA loop configurations that they have in sperm nuclei, it would suggest that this organization might help to regulate transcription and DNA replication in early embryonic development. This would have the exciting implication that the sperm nucleus provides the embryo with a specific chromosomal architecture that may be functional during embryogenesis.

Level III: protamine decondensation In the third step of assembly of the sperm chromatin structure, the binding of protamines condenses the DNA loops into tightly packaged chromatin. Hud et al.61 have demonstrated that when protamines bind DNA, they form toroidal, or doughnut-shaped, structures in which the DNA is very concentrated. During spermiogenesis, histones, the DNA-binding proteins of somatic spermatogenic precursor cells, are replaced by protamines. Since histone-bound DNA requires much more volume than the same amount of DNA bound to protamines16, this change in chromatin structure probably accounts for some of the

nuclear condensation that occurs during spermiogenesis. Histones package DNA by organizing it into nucleosomes, in which the DNA is wrapped around an octamer of histone proteins. Protamines, on the other hand, bind DNA in a markedly different manner. These positively charged proteins bind DNA along the major groove, completely neutralizing the DNA so that neighboring DNA strands bind to each other by van der Waals forces. Protamines are believed to coil the DNA into doughnut-like structures in which the DNA exists in an almost crystalline-like state61. If each toroid is a single DNA loop domain62, protamine binding will lead to condensation and preservation of the DNA loop domain organization present in the round spermatid.

Level IV: chromosome organization The next level of sperm chromatin packaging is the spatial arrangement of the condensed chromosomes within the mature sperm nucleus. This has been investigated in several different ways. First, Zalensky and co-workers45 demonstrated that, in human sperm nuclei, the centromeres of all chromosomes are aggregated in the center of the nucleus, while the telomeres are located at the periphery. In a second approach, Haaf and Ward63 analyzed whole chromosomes and found similar results. Finally, Ward and co-workers64 mapped the three-dimensional location of three genes in the hamster sperm nucleus and found that while each one tended to be located in the outer third of the nucleus, there was otherwise little specificity to the positioning of the genes. These data led to the proposal of a model65 in which there are limited constraints on the actual position of chromosomes in the sperm nucleus. The NA-DNA sequences are located at the base of the nucleus, centromeres are located centrally and the telomeres are located peripherally. Outside these three constraints, the folding of the chromosomal p and q arms is flexible. Interestingly, this type of organization does

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not seem to be present in montreme mammal spermatozoa. In these species, chromosomes are aligned end-to-end66. In most eutherian mammals examined, however, the centromeres are organized in a central location, making such an end-to-end arrangement impossible.

ROLE OF SPERMATOZOA IN EMBRYOGENESIS For many years, male fertility has been defined in vitro as the possibility of sperm to fertilize the oocyte, and to obtain early cleavage-stage embryos. In human in vitro fertilization (IVF), the gold-standard/test for sperm fertility potential was the ability of a fertilized egg to develop into a 2–4-cell embryo. It was assumed that all embryos obtained had the same developmental potential, independent of the quality of sperm. Thereafter, several authors67–69 observed that poor morphological embryonic quality and poor embryonic developmental ability are associated with severe sperm morphological defects and oligoasthenozoospermia. In addition, Janny and Ménézo70 observed a negative relationship between sperm quality and the ability to reach the blastocyst stage. We now know that differences in sperm fertility are not simply related to sperm penetration failure. The following is an analysis of the chronology of the steps involved in these embryonic failures.

Early defects at the time of fertilization The centrosome

The first epigenetic contribution of the spermatozoon is the centrosome, the microtubuleorganizing center of the cell. Correct assembly and function of microtubules is fundamental for the separation of chromosomes at meiosis and migration of the male and female pronuclei. Maternal inheritance of the centrosome observed

in mice brought about confusion until the work of Schatten71 and Simerly et al.72. Considering the semiconservative form of this organelle and its critical role in mitosis, it seems obvious that a functionally imperfect centrosome borne by a subnormal spermatozoon induces problems in early embryogenesis, i.e. the formation of cytoplasmic fragments and abnormal distribution of chromosomes72,73. Asch et al.74 reported that up to 25% of non-segmented eggs are in fact fertilized but submitted to cell division defects. Centrin and γ-tubulin could be involved in this pathology of the centrosome75. In bovine oocytes, Navara et al.76 observed a positive relationship between size and quality of the sperm aster and reproductive performance in bulls. Oocyte activation factor(s)

The process of meiosis reinitiation is probably completed through an exit from the M phase due to cyclin B degradation and re-phosphorylation of p34cdc2 following a decrease in cytostatic factor (CSF)77. It is generally accepted that intracellular Ca2+ is the universal signal for triggering oocytes into metabolic activity. It is still not clear how the spermatozoon causes this calcium oscillation. A heat-sensitive78 and soluble protein called oscillin, acting through the inositol phosphate pathway, could be at the origin of these calcium oscillations79. Defects in oscillin (or other soluble activating factors) could account for delays in zygote formation, as described by Ron-El et al.68. However, for Eid et al.80, based on their observations in bovine zygotes, this hypothesis might not be the only one. In a group of embryos sired by low-fertility bulls, they did not observe any delay in pronuclear formation, but a delayed initiation and reduced length of zygotic S-phase correlated with reduced embryonic development in vitro. A longer S-phase was correlated with higher fertility in vivo. Poor chromatin packaging and/or anomalies in DNA packaging could contribute to the failure of sperm decondensation, independently of any activation problems81.

PATHOPHYSIOLOGY AND GENETICS OF HUMAN MALE REPRODUCTION

Developmental arrests between fertilization and the beginning of genomic activation It is quite surprising to expect a paternal-derived influence between fertilization and genomic activation, i.e. before the appearance of the first products resulting from the first massive transcriptions involving the paternal genome. It is now well documented that the longest cleavage stage is linked to embryonic genome activation82. There is obviously a race against the clock between, first, the ineluctable turnover of the maternal mRNA and, on the other hand, the first massive synthesis of the embryonic transcripts. The cumulative delays observed, cycle after cycle, due to epigenetic defects brought about by suboptimal spermatozoa lead to developmental arrests, the maternal stores being exhausted before the beginning of transcription. Under in vitro conditions, one-third of human IVF embryos block around the time of genomic activation70. Antisperm antibodies may also have a deleterious effect on early preimplantation development. Naz83 observed that antibodies against very special epitopes might block embryos, especially if cleavage signal proteins (CS-1) or regulatory products of the OCT-3 gene are immunoneutralized.

Developmental arrests between genomic activation and implantation After genomic activation, the very sensitive transition between morula and blastocyst follows. Complex remodeling within the embryo occurs with the first differentiation. Janny and Ménézo70 observed a loss of blastocysts at this point, which was significantly increased in men with poor sperm quality (31% vs. 22% for the control group). They concluded that poor-quality sperm has a negative influence on preimplantation development even after genomic activation. The lesson from ICSI

One of the most exciting breakthroughs in the treatment of male infertility is intracytoplasmic

sperm injection (ICSI). The success that we observe in ICSI, considering the poor quality of the sperm, can partially be ascribed to the following. In human IVF with poor-quality sperm, delay in the fertilization process68 and delays associated with epigenetic problems, the cumulative effect may be prolonged cell cycles and late divisions (2cell embryos on day 2, 4-cell on day 3), leading to developmental arrest around genomic activation, in relation to depletion of the mRNA maternal store. In contrast, the fertilization process in ICSI is shorter, since the sperm is introduced into the cytoplasm. Van Landuyt and co-workers84 showed that the blastocyst formation rate after ICSI compares to the rate after regular IVF. An in-depth analysis, however, demonstrates that more patients have embryos which are unable to reach the blastocyst stage. Interestingly, there seems to be an ‘all or nothing’ trend regarding blastocyst development. If one blastocyst is obtained, then all embryos from the patient in question normally develop into blastocysts, whilst if no blastocysts are seen at day 5, it is highly unlikely that any embryos will go on to develop into blastocysts. ICSI is of no use if performed 24 hours after failed fertilization: the maternal mRNA reserves are already at this point too depleted to allow development from fertilization to genomic activation. It is very likely that major sperm defects cannot be corrected by the application of ICSI. The ICSI process itself carries other geneticrelated problems such as the genetic link between oligoasthenoteratozoospermia and sperm genetic disorders. Some of these features include microdeletions of the Y chromosome85. The negative influence of suboptimal spermatozoa is linked to the integrity and quality of the paternal DNA. In 1985, Bourrouillou and co-workers86 observed an increase in chromosomal abnormalities as a function of sperm count; in 1995, Moosani and co-workers87 clearly demonstrated increased chromosomal disorders in the sperm population of infertile men with idiopathic infertility.

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In this context, it is also important to mention the consequences of fertilization of oocytes with sperm deriving from an ejaculate containing a high incidence of disturbed DNA integrity in IVF and, especially, in ICSI patients. According to present knowledge, sperm DNA fragmentation might cause not only impaired embryonic development and early embryonic death74,88,89, but also an increased risk of childhood cancer in the offspring90,91. The latter is due to the vulnerability of human sperm DNA during late stages of spermatogenesis and epididymal maturation. At this stage, DNA repair mechanisms have been switched off, resulting in a genetic instability of the male germ cells92, especially on the Y chromosome, resulting in male-specific cancers93. However, this DNA damage is not only caused by these intrinsic factors, but can also be triggered by extrinsic factors such as excess amounts of oxidants producing leukocytes in the ejaculate94. The influence of the spermatozoon-carried mitochondria during ICSI, on early or late embryogenesis, is, however, still a matter of debate.

Genomic imprinting Experimental manipulations of mouse zygotes have clearly proved the necessary complementary relationship between the maternal and the paternal genome to ensure normal embryonic development. Even if implantation and late development can be observed in the rabbit and mouse, parthenogenesis never leads to live births. Surani et al.95 observed that hypertrophy of the inner cell mass and hypotrophy of the extraembryonic tissue is related to gynogenesis. In contrast, androgenesis performed by removal of the female pronucleus followed by duplication of the paternal genome leads to hypertrophy of extraembryonic tissues. This is due to genomic imprinting, which occurs as early as the pronuclear stage. Genomic imprinting seems to be directly related to variations in the methylation pattern of some genes. One of the most important systems in genomic imprinting is IGF2/IGF2-R96. The ligand is contributed by the

paternal genome and the receptor by the maternal one. The maternal and paternal X chromosomes are submitted to differential inactivation, related to different methylation patterns of the Xist locus, in the preimplantation period. Xist is the initiator of methylation carried by the X chromosome. The H19 gene, a tumor suppressor, is expressed in the placenta but not in the mole. The potential invasiveness of the placenta and/or placental tumors is directly related to the paternal genome qualitatively and quantitatively97. Disorganized imprinting may have harmful effects on early-preimplantation and late-postimplantation development.

CONCLUSIONS As discussed, it is clear that paternal factors have major effects on early embryogenesis. In the past decade, major advances have been made in assisted reproductive technologies. ICSI has been proposed as a tool for overcoming sperm deficiencies observed at the time of fertilization. This technology can assist in overcoming some of the defects affecting early-preimplantation development. Time gained by direct sperm insertion into the cytoplasm may help in avoiding delays that impair early-preimplantation development. However, it is unlikely that ICSI can universally compensate for male-factor defects. Moreover, it raises questions regarding the genetic basis of some of the defects observed, and on some other hidden genetic links. The growing number of children that have followed the application of ICSI is beginning to provide us with a good base to evaluate the transmission of genetic defects. To date, there is evidence showing that infertility in fathers due to microdeletions in the Y chromosome is transmitted from one male generation to the next98,99. These examples of male infertility are believed to be due to deletion of genes such as the DAZ (deleted in azoospermia) and RBM (RNAbinding motif ) genes. These genes show mapping to Y chromosome-linked microdeletions100–103.

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95. Surani MAH, Barton SC, Norris ML. Experimental reconstruction of mouse eggs and embryos: an analysis of mammalian development. Biol Reprod 1987; 36: 1 96. De Groot N, Hochberg A. Gene imprinting during placental and embryonic development. Mol Reprod Dev 1993; 36: 390 97. Goshen R, et al. The role of genomic imprinting in implantation. Fertil Steril 1994; 62: 903 98. Chandley AC, et al. Deleted Yq in the sterile son of a man with a sattelite Y chromosome (Yqs). J Med Genet 1989; 26: 145 99. Kent-First MG, et al. Infertility in intracytoplasmicsperm-injection-derived sons. Lancet 1996; 348: 332

100. Reijo R, et al. Diverse spermatogenic defects in humans caused by Y chromosome deletions encompassing a novel RNA-binding protein gene. Nat Genet 1995; 10: 383 101. Najmabadi H, et al. Substantial prevalence of microdeletions of the Y chromosome in infertile men with idiopathic azoospermia and oligozoospermia detected using a sequence-tagged site-based mapping strategy. J Clin Endocrinol Metab 1996; 81: 1347 102. Reijo R, et al. Severe oligozoospermia resulting from deletions of azoospermia factor gene on Y chromosome. Lancet 1996; 347: 1290 103. Pryor JL, et al. Microdeletions in the Y chromosome of infertile men. N Engl J Med 1997; 336: 534

4 Contribution of the male gamete to fertilization and embryogenesis Gerardo Barroso, Sergio Oehninger

INTRODUCTION

BIOLOGY OF FERTILIZATION

The normal progression of fertilization of mammalian oocytes followed by cleavage, blastocyst formation and implantation is dependent upon the successful activation of specific genetic and developmental programs. Successful interaction of the paternal and maternal gametes is required for normal embryonic development. The oocyte controls several important aspects of meiosis, fertilization and early cleavage, and modulates the epigenetic development of the embryonic genome that manifests later in embryogenesis1. The contributing role of the spermatozoon has remained largely ignored. However, a large body of evidence is accumulating demonstrating that (1) the fertilizing spermatozoon plays a significant part in bringing about the development of the zygote, with its contributions being well beyond the delivery of the paternal DNA; and (2) infertile men with or without altered ‘classic’ semen parameters may have associated sperm dysfunction(s) at different levels, including nuclear2, organelle-cytoplasmic3 and cytoskeletal systems4, that can result in aberrant embryogenesis. The mechanism(s) underlying these phenomena is/are not completely understood. This review focuses on examination of the paternal effects that become manifest before and after the major activation of embryonic gene expression.

Sperm–oocyte fusion The sperm equatorial region plays a pivotal role in gamete fusion. The inner and outer acrosomal membranes and the plasma membrane of the equatorial region remain intact after completion of the acrosome reaction and zona penetration5. Electron microscopic studies have shown convincingly that sperm–oocyte membrane fusion takes place at the sperm equatorial region, whereas the posterior acrosome itself is engulfed by the oocyte microvilli in a phagocytic manner. Acrosome-reacted sperm bind to and fuse with eggs by using the plasma membrane at the postacrosomal region of the sperm; this region is capable of fusion only after acrosomal exocytosis has taken place6. Binding of the sperm to the egg plasma membrane appears to be mediated by a member of the ADAM (a disintegrin and metalloprotease) family of transmembrane proteins on the sperm and integrin α6β1 receptors on the egg7. Fertilin is a heterodimeric ADAM glycoprotein that was first identified in the guinea-pig using monoclonal antibodies to sperm surface antigens that could inhibit sperm–egg fusion8. The protein is composed of an α and a β subunit with similar domain structures9,10, and is proteolytically processed during sperm development by 49

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removal of the prodomain and metalloprotease domain. Processing of fertilin is crucial for exposing the disintegrin domain that mediates sperm–egg binding, and for allowing proper localization of fertilin in the head of the mature sperm11,12. More recently, a member of the immunoglobulin superfamily (the membrane protein Izumo) has been found to be critically involved in murine sperm–oocyte fusion13. Equatorin is a sperm-head equatorial protein, the antigenic molecule of the monoclonal antibody mMN914. In mice, after sperm–egg fusion, equatorin dissociates from the sperm-head equatorial region and remains at the vicinity of the decondensing male pronucleus. The equatorial segment containing equatorin is maintained away from the nuclei, possibly due to chromatin swelling and nuclear membrane reconstruction. It remains at the vicinity of the sperm head for a considerable length of time during the first cell cycle, and, after that, it is inherited by one of the proembryonic cells. After intracytoplasmic sperm injection (ICSI), the equatorial segment is directly exposed to the oocyte cytoplasm without prior interaction with the cortical membrane system, but displays similar cellular events of equatorin degeneration to the oocyte after in vitro fertilization (IVF). These observations argue in favor of membrane interaction not being a prerequisite for shedding the equatorial posterior acrosome, equatorin, and their subsequent disintegration after ICSI15. The persistence of equatorin through earlyproembryonic cleavage is comparable with that of sperm-tail microtubules and the midpiece mitochondrial sheath. The residual tail microtubules are retained up to the 8-cell or blastocyst stage. However, the residual equatorin seems to degenerate a little early, before the 4-cell stage15.

Oocyte activation The oocyte and spermatozoon are metabolically quiescent; sperm–oocyte binding and fusion initiate a cascade of events that transform the dormant

oocyte into the dynamic, animated zygote. These processes include metabolic oocyte activation and resumption of meiosis. Although there are still diverse opinions as to the precise manner in which the spermatozoon activates this cascade, it is clear in all fertilization systems that an elevation of intracellular calcium ion concentration is the central messenger in communicating the activating signal. The signaling mechanism(s) utilized by the spermatozoon to initiate and perpetuate these responses is unclear. Two theories have been proposed: the fusion and the receptor theories (reviewed in reference 16). The fusion theory suggests the presence of active calcium-releasing components in the sperm head. It has experimental support in that injections of sperm-derived cytosolic fractions elicit calcium oscillations, and also in that ICSI results in activation without sperm interaction with the membrane. It was recently reported that a cytosolic sperm factor containing a 33-kDa protein called oscillin, which is related to a prokaryote glucosamine phosphate deaminase, appeared to be responsible for causing the calcium oscillations that trigger egg activation at fertilization in mammals17. Oscillin is located in the equatorial segment of the spermatozoon, the region where the spermatozoon is fused with the oocyte in mammals. However, multiple pieces of experimental evidence have now shown that oscillin is not the mammalian sperm calcium oscillogen (reviewed in reference 16). In eggs of all animal species, sperm-triggered inositol (1,4,5)-triphosphate (IP3) production regulates the vast array of calcium wave-patterns observed. Present evidence supports the concept that an IP3 receptor system is the main mediator of calcium oscillations in oocytes (reviewed in reference 16). The spatial organization of calcium waves is driven either by intracellular distribution of the calcium-release machinery or by localized and dynamic production of calcium-releasing second messengers. In the highly polarized egg cell, cortical endoplasmic reticulum-rich clusters act as pacemaker

CONTRIBUTION OF MALE GAMETE TO FERTILIZATION AND EMBRYOGENESIS

sites dedicated to the initiation of global calcium waves. The polarized nature of the calcium signals may in itself influence embryonic patterning by regulating early embryonic cleavage. Finding out whether calcium wave-patterns play a role in later development will require studies that interfere with the normal spatial–temporal pattern of calcium waves without perturbing mitosis and cleavage. The rather simple ascidian embryo, which displays two different meiotic calcium-wave pacemakers and develops into a swimming tadpole within a day, is particularly suited to studies of the relationship between meiotic calcium waves and development18. It should be possible in the future to relate patterns of calcium waves and phenotypic differences in embryos. In recent years, mitochondria have been shown to be major regulators of intracellular calcium homeostasis19,20. In cells such as sea urchin21 and ascidian eggs22, mitochondria sequester calcium during the fertilization calcium transients. Calcium sequestration by mitochondria has two main consequences. First, mitochondria act as passive calcium buffers that can regulate intracellular calcium release19,20. The second consequence is that calcium in the mitochondrial matrix is a ‘multisite’ activator of oxidative phosphorylation (or mitochondrial adenosine triphosphate (ATP) synthesis); it activates the dehydrogenases of the Krebs cycle and the electron transport chain23,24 and has a direct action on the F0/F1 ATP synthase25. In somatic cells and in ascidian eggs, mitochondrial calcium uptake has been shown to stimulate mitochondrial respiration by promoting the reduction of mitochondrial nicotinamide–adenine dinucleotide (NAD+) to NADH22,26–28. Furthermore, mitochondrial ATP production may directly regulate intracellular calcium release: ATP sensitizes the IP3 receptor to activation by calcium29,30, while magnesium-complexed ATP is consumed to refill the endoplasmic reticulum calcium stores. The tight coupling of ATP supply and demand therefore provides a major advantage for early mammalian development. The maternal inheritance of mitochondria requires that

mitochondria be protected from potentially damaging reactive oxygen species (ROS). The maintenance of a low level of oxidative phosphorylation that can be stimulated upon increased ATP demand provides a means of lowering the exposure of mitochondria to damaging oxidative stress. Data suggest that calcium is the functional link that provides a mechanism for coupling ATP supply and demand. As maternal aging is associated with increased oxidative stress in human eggs31, it will be interesting to define whether mitochondrial physiology and the coupling of ATP supply and demand are impaired in eggs from aged women. It has recently been shown that the soluble sperm factor that triggers calcium oscillations and egg activation (oocyte activating factor, OAF) in mammals is a novel form of phospholipase C (PLC) referred to as PLCζ32. This has been demonstrated by injection into eggs of both cRNA encoding PLCζ and a recombinant PLCζ32,33. According to a present hypothesis, after fusion of the sperm and egg plasma membrane, the sperm-derived PLCζ protein (possibly a sperm cytosolic factor) diffuses into the egg cytoplasm. This results in hydrolysis of phosphatidylinositol4,5-bisphosphate (PIP2) from an unknown source to generate IP3 (reviewed in reference 34). The earliest indicators of the transition to embryos in mammalian eggs, or egg activation, are cortical granule extrusion by exocytosis (CGE) and resumption of meiosis. Although these events are triggered by calcium oscillations as described above, the pathways within the egg leading to intracellular calcium release and to downstream cellular events are not completely understood. The calcium transients actuate resumption of the cell cycle by decreasing the activity of both the M phase-promoting factor and the cytostatic factor (reviewed in reference 35). The calcium transients and/or activation of PLCζ lead to CGE by an, as yet, undefined mechanism36. Src family kinases (SFKs) have been suggested as possible inducers of some aspects of egg activation (reviewed in reference 37). A present model

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claims that sperm fusion with the egg membrane results in hydrolysis of PIP2 to form IP3 and diacylglycerol (DAG). IP3 triggers calcium release from the endoplasmic reticulum via the IP3 receptor while DAG activates protein kinase C (PKC). Both an intracellular calcium rise and DAG contribute to egg activation, CGE and resumption of meiosis. The existence of SFK activity is associated with the resumption of meiosis in response to the fertilization signal, whereas the occurrence of CGE is independent of SFK activity. Also, a role for SFKs upstream of calcium release remains plausible (reviewed in reference 37).

Sperm mitochondrial DNA and its role during fertilization Mitochondria have a profound role to play in mammalian-tissue bioenergetics during the processes of growth, aging and apoptosis, and yet they descend from an asexually reproducing independent life form. Most cells in the body contain between 103 and 104 copies of mitochondrial (mt)DNA. There are slightly higher copy numbers (about 105) in mature oocytes. This may be in preparation for the energetic demands of embryogenesis38, but an alternative explanation is that replication does not occur during early embryogenesis and that high copy numbers are needed to give a sufficient reservoir. The DNA exists mainly as a circular molecule of approximately 16.6 kb, encoding 13 proteins that are transcribed and translated in the mitochondrion. These are essential subunits of the electron transport complexes on the inner mitochondrial membrane. The mitochondrial genome also encodes the RNA molecules that are necessary for translation of these proteins39,40. Spermatozoa are metabolically flexible and, in some species, can switch between aerobic and anaerobic metabolism. This perhaps reflects the great range of oxygen tensions that they experience, from near anoxia in the testis and epididymis to ambient tensions in the vagina and in vitro3,41,42. Like somatic mtDNA, that of

spermatozoa is highly vulnerable to mutation, and a significant number of mtDNA deletions are found in the semen of at least 50% of normospermic men43. Given the lengthy process of spermiogenesis and epididymal maturation, during which the sperm mitochondria have to survive the likelihood that they will be exposed to mutagenic agents, this is perhaps not surprising. Indeed, the need to exclude defective sperm mtDNA from contributing to the embryo is possibly one of the major selection pressures against survival of paternal mtDNA. Indeed, Short44 has suggested that this asymmetric inheritance of mtDNA, through the oocyte but not the spermatozoon, may be the fundamental driving force behind amphimixis and anisogamy. This is because of the need to conserve a healthy stock of mtDNA for embryo development through a long period of quiescence in meiosis43. It is well established that the mitochondria from spermatozoa are targeted for destruction by endogenous proteolytic activity during early embryogenesis. Uniparental (generally maternal) inheritance of cytoplasmic organelles such as mitochondria is accomplished by a wide variety of strategies, and thus is clearly of profound importance to long-term fitness. Most evidence indicating the possibility of paternal transmission of mtDNA derives from interspecific crosses, which by definition are uncommon in nature45. In a previous study, Kaneda et al.45 proposed that the zygote cytoplasm has a species-specific mechanism that recognizes and eliminates sperm mitochondria, on the basis of nuclear DNA-encoded proteins in the sperm midpiece, and neither on the mtDNA itself, nor on the proteins it encodes.

The ubiquitination–proteasome pathway The fate of various accessory structures of the penetrating spermatozoon came under scrutiny recently, as it became obvious that in addition to the sperm-borne chromosomes, other structures

CONTRIBUTION OF MALE GAMETE TO FERTILIZATION AND EMBRYOGENESIS

of the fertilizing spermatozoon make important contributions to the mammalian zygote. Yet other sperm accessory structures are degraded in an orderly fashion so as to not interfere with normal embryo development. These include the sperm proximal centriole, perinuclear theca, sperm mitochondria and axonemal fibrous sheath and outer dense fibers. In most mammals, except rodents, the spermatozoon contains a reduced, inactive form of the centrosome, within which one of the two centrioles as well as the entourage of pericentriolar material are degraded during the final stages of spermiogenesis. Such an incomplete centrosome, consisting of a proximal centriole embedded in the dense mass of sperm-tail capitulum, must be released into the oocyte cytoplasm at fertilization in order to attract microtubule-nucleating pericentriolar proteins from the surrounding oocyte cytoplasm. Failure to convert the reduced sperm centriole into such an active zygotic centrosome may be a reason for postfertilization developmental arrests affecting couples treated at IVF clinics. The strictly maternal inheritance of mtDNA in mammals is a developmental paradox, because the fertilizing spermatozoon introduces up to 100 functional mitochondria into the oocyte cytoplasm at fertilization. However, the mandatory destruction of sperm mitochondria appears to be an evolutionary and developmental advantage46, because the paternal mitochondria and their DNA may be compromised by the deleterious action of reactive oxygen species encountered by the sperm during spermatogenesis, storage, migration and fertilization47. Although a number of studies have supported the notion that sperm mitochondria are actively destroyed by the egg, the actual mechanism of this process is not known48–50. Earlier claims that the sperm mitochondria disperse evenly throughout embryonic cytoplasm51 and the misconception about sperm mitochondria not entering the egg were overturned by new research. The dilution of paternal mtDNA in the maternal cytoplasm genome52 and the oxidative damage of sperm

mitochondria during fertilization53 were also implicated in this process, but were not adequately supported by experimental data. Ubiquitination of the sperm mitochondria during spermatogenesis has been implicated in the targeted degradation of paternal mitochondria after fertilization, a mechanism proposed to promote the predominantly maternal inheritance of mitochondria DNA in humans. Recent studies54,55 have shown that some unknown proteins in mammalian sperm mitochondria are tagged with a proteolytic peptide, ubiquitin, which may target sperm mitochondria for destruction in the egg cytoplasm after fertilization. Both lysosomal and proteasomal proteolysis have been implicated in such targeted degradation of sperm mitochondria inside the fertilized oocyte55. This mechanism seems to be feasible for the selective degradation of paternal mitochondria at fertilization, sometimes described as the ‘ultimate war of the sexes’, and is consistent with the prevailing view that the inheritance of mtDNA in mammals is predominantly maternal56. Such a scenario is also supported by studies of mitochondrial inheritance in inter- and intraspecies murine crosses as well as in their back-crossed progeny, in which the mitochondrial membrane proteins, rather than mtDNA, seemed to determine whether the sperm mitochondria and mtDNA were passed on or degraded45. Ubiquitination is the major means in eukaryotic cells for targeted protein proteolysis. By the covalent addition of polyubiquitin to specific proteins, the ubiquitination system regulates protein levels and thereby influences diverse cellular processes. There are three well-established types of enzymes involved in ubiquitination, termed E1, E2 and E3. E1 is the ubiquitin-activating enzyme, which forms a thiol-ester linkage with ubiquitin through its active site cysteine. Ubiquitin is subsequently transferred to an E2 ubiquitin-conjugating enzyme; the E3 enzyme is the ubiquitin protein ligase, which transfers ubiquitin from the E2 enzyme to lysines of a specific protein, targeting the protein for degradation by the proteasome.

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More recently, E4 enzymes have been described that appear to function in ubiquitin chain polymerization.

SNDF Head decondensation

Pronuclear formation and nuclear fusion The fertilizing spermatozoon is essential for contributing three critical components: (1) the paternal haploid genome, (2) the signal to initiate the metabolic–maturational activation of the oocyte and (3) the centrosome, which directs microtubule assembly within the penetrated oocyte leading to oocyte–sperm activation as well as formation of the mitotic spindles during initial zygote development (Figure 4.1). Fertilization is completed once the parental genomes unite (syngamy), and requires migration of the egg nucleus to the sperm nucleus (female and male pronuclei) on microtubules within the penetrated oocyte. The male pronucleus is tightly associated with the centrosome, which nucleates microtubules to form the sperm aster. The growth of the sperm aster drives the centrosome and associated male pronucleus from the cell cortex towards the center of the oocyte. In contrast to the male pronucleus, the female pronucleus has neither an associated centrosome nor microtubule-nucleating activity. Nevertheless, the female pronucleus moves along microtubules from the cell cortex towards the centrosome located in the center of the sperm aster. The current model for movement of the female pronucleus involves its translocation along the microtubule lattice using the minus-end-directed motor dynein53,57,58, in a manner analogous to organelle motility. Mammalian fertilization requires dynein and dynactin to mediate genomic union, and that dynein concentrates exclusively around the female pronucleus. Dynactin, by contrast, localizes around both pronuclei and associates with nucleoporins and vimentin in addition to dynein. The findings that a sperm aster is required for dynein to localize to the female pronucleus and the microtubules are necessary to retain dynein,

Centrosome OAF

Pronuclear formation

[Ca2+]

Cleavage Resumption of meiosis

Figure 4.1 Critical sperm components during fertilization. OAF, oocyte activating factor; SNDF, sperm nuclear decondensing factor

but not dynactin, at its surface, suggest that nucleoporins, vimentin and dynactin might associate upon pronuclear formation, and that subsequent sperm aster contact with the female pronuclear surface allows dynein to interact with these proteins59.

EVIDENCE FOR PATERNAL CONTRIBUTIONS TO ABNORMAL EMBRYOGENESIS Clinical evidence: lessons from the IVF/ICSI setting Several lines of clinical evidence resulting from the use of assisted reproductive technologies have provided additional support for the concept of paternal contribution to faulty fertilization and abnormal embryogenesis: • Abnormal sperm parameters, particularly teratozoospermia (‘poor prognosis pattern’ as defined by strict criteria), are associated with fertilization disorders in IVF, including failure (partial or complete) and delayed fertilization60,61.

CONTRIBUTION OF MALE GAMETE TO FERTILIZATION AND EMBRYOGENESIS

• Results of standard (conventional) IVF in men with severe teratozoospermia and other seminal abnormalities showed not only decreased fertilization but also lower implantation rates compared with normozoospermic samples62–65. • The application of corrective measures in conventional IVF (such as increasing sperm insemination concentration) resulted in an enhanced fertilization rate but implantation rates remained lower than anticipated66. • Poor sperm quality was associated with a decreased ability to reach the blastocyst stage in vitro67. • A comparative analysis of embryo implantation potential in patients with severe teratozoospermia undergoing IVF with a high insemination concentration or ICSI revealed that ICSI produced a significant proportion of embryos with superior morphology and implantation competence68. • Although multiple studies have shown that the outcome of clinical pregnancies following ICSI is not affected by semen quality69–72, patients with total teratozoospermia demonstrated a very low implantation rate73. • Spermatozoa of infertile men have also been shown to contain various nuclear alterations. They include an abnormal chromatin structure, aneuploidy, chromosomal microdeletions and DNA strand breaks74–81. Different theories have been proposed to explain the origin of DNA damage in spermatozoa (reviewed in references 2, 80 and 82). Damage could occur at the time of, or be the result of, DNA packing during the transition of histone to the protamine complex during spermiogenesis. DNA fragmentation could also be the consequence of direct oxidative damage (free radicalinduced DNA damage has been associated with antioxidant depletion, smoking, xenobiotics, heat exposure, leukocyte contamination of semen and the presence of ions in sperm culture media).

Alternatively, DNA damage could be the consequence of apoptosis. • Numerous studies have demonstrated associations between poor sperm quality and increased sperm aneuploidy, DNA damage, fragmentation and instability and singlestranded DNA, with poor pregnancy potential documented in such cases undergoing intrauterine insemination (IUI) or ICSI therapies83–89. • Although the major congenital malformation rate and developmental potential of children conceived after IVF or ICSI and naturally are similar, ICSI is associated with a slight increase in de novo chromosomal abnormalities. Moreover, recent publications mention that diseases caused by imprinting disorders affect a few ICSI children, and sperm from men with severely impaired semen quality may carry microdeletions of the Y chromosome and other genetic disorders (reviewed in references 90 and 91). Consequently, spermatozoa from infertile men may carry chromosomal and/or genetic abnormalities that can be potentially transmitted to the offspring92. In addition, findings in animals and in the human have provided evidence of paternal transmission of genetic damage, including data on paternally mediated behavioral effects, male-mediated teratogenicity and tumor induction and susceptibility in the offspring. The available evidence indicates that preconception paternal exposure to certain mutagens can, under certain conditions, have adverse effects on the offspring. Two principal mechanisms proposed are the induction of germ-line genomic instability or the suppression of germ cell apoptosis (reviewed in reference 93). It is well established that the presence of sperm abnormalities can lead to failure of fertilization. A high proportion of infertile men possess sperm functional deficiencies that result in poor interaction with the zona pellucida, including a diminished capacity to achieve tight binding and/or to

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undergo acrosomal exocytosis. Moreover, a deficient interaction with the oolema can lead to binding or fusion abnormalities94–97. Obviously, failure of the spermatozoon to penetrate the oocyte’s investments or to arrive at the cytoplasm negates fertilization and embryogenesis. Other sperm abnormalities have been associated with failed fertilization and aberrant or arrested embryo development. Such instances include delayed fertilization, abnormal oocyte activation, deficient sperm-head decondensation, defective pronuclear formation and poor embryo cleavage (reviewed in references 96, 98 and 99). Once the spermatozoon penetrates the oocyte, several events must take place to ensure fertilization, including incorporation of the entire spermatozoon into the oocyte, completion of oocyte meiotic maturation with extrusion of the second polar body, metabolic activation of the previously quiescent oocyte, decondensation of the sperm nucleus and the maternal chromosomes into the male and female pronuclei, respectively, and cytoplasmic migrations of the pronuclei, which bring them into apposition. Defects in any of these events can be lethal to the zygote and can be causes of infertility. As mentioned earlier, it is generally accepted that the contributions of the fertilizing spermatozoon to the oocyte include delivery of the DNA/chromatin, a putative oocyte-activating factor (OAF) and a centriole. The DNA/chromatin complex is obviously the most significant contribution to originating a new diploid individual. Nevertheless, the OAF and centriole play a critical part in bringing about oocyte activation, cortical granule extrusion and the first mitotic division, and without these contributions embryogenesis would also be neglected or proceed abnormally. The fate of sperm components in primate models (human and subhuman) during fertilization is being unraveled. The centrosome, introduced by the sperm at fertilization, organizes a microtubule array that is responsible for bringing the parental genomes together at first mitosis. Structural abnormalities or incomplete functioning

of the centrosome have been identified as a novel form of infertility100. Moreover, the paternal sperm-borne mitochondria also enter the cytoplasm and are specifically targeted for degradation by the resident oocyte ubiquitin system101. This phenomenon allows for maternal inheritance of mitochondrial DNA. Defects of paternal mitochondrial degradation could result in heteroplasmy. New evidence has challenged the traditional view of the transcriptional dormancy of terminally differentiated spermatozoa. Several reports have indicated the presence of mRNAs in ejaculated human spermatozoa (reviewed in reference 102). It has been hypothesized that these templates could be critically involved in late spermiogenesis, including a function to equilibrate imbalances in spermatozoal phenotypes brought about by meiotic recombination and segregation, and furthermore, that they could also be involved in early postfertilization events such as establishing imprints during the transition from maternal to embryonic genes. Cell divisions in the human embryo can be compromised by deficiencies in the sperm nuclear genome or sperm-derived cytoplasmic factors, including the OAF and centriole. The newly formed zygote undergoes early cleavage divisions depending upon the oocyte’s endogenous machinery, and at the 4–8-cell stage initiates transcription of the embryonic genome103. Consequently, sperm nuclear deficiencies are usually not detected before the 8-cell stage, when a major expression of sperm-derived genes has begun. On the other hand, sperm cytoplasm deficiencies can be detected as early as the 1-cell zygote and then throughout the preimplantation development104,105. The terms ‘late’ and ‘early’ paternal effect have been suggested to denote these two pathological conditions106. The diagnosis of an early paternal effect is based upon poor zygote and early embryo morphology and low cleavage speed, and is not associated with sperm DNA fragmentation. The late paternal effect, on the other hand, is manifested by poor developmental competence leading

CONTRIBUTION OF MALE GAMETE TO FERTILIZATION AND EMBRYOGENESIS

to failure of implantation, and is associated with an increased incidence of sperm DNA fragmentation in the absence of zygote and early cleavagestage morphological abnormalities. It has been suggested that ICSI with testicular sperm can be an efficient treatment for the late paternal effect107. It can be speculated that the early paternal effect probably includes dysfunctions related to oocyte activation and the centrosome and cytoskeletal apparatus, as well as possible abnormal mRNA delivery. Conversely, the late paternal effect is associated with dysfunctions/abnormalities of the DNA/chromatin (including sperm chromosomal–genetic aberrations, retention of histones and/or DNA damage), and perhaps mitochondrial dysfunctions. Alterations due to genomic imprinting anomalies probably result in both early and late paternal effects.

Disorders of oocyte activation, centrosome and cytoskeletal apparatus dysfunction and mitochondria elimination PLCζ offers the molecular basis for an explanation of how calcium release is triggered during mammalian fertilization. There are clinical situations that can be explained by the absence or dysfunction of the OAF. For example, it has been suggested that up to 40% of failed fertilization cases after ICSI could be due to failure of the egg to activate 99. In these cases the sperm is within the cytoplasm, but a stimulus for activation is apparently missing. Certainly, there may be cases where the spermatozoon provides the OAF, but any of the multiple elements of the oocyte-responsive system (SFKs, PIP2, IP3 receptor or PKC) is aberrant, resulting in failure to resume meiosis or to undergo CGE. During fertilization the zygotic centrosome organizes a large sperm aster critical for uniting the pronuclei before the first mitosis. Dysfunctional microtubule organization in failed

fertilization during human IVF suggests that centrosomal dysfunction might be a cause of fertilization arrest. In a study by Asch et al.98 microtubules and DNA were imaged in inseminated human oocytes that had been discarded as unfertilized. The presence and number of incorporated sperm tails were also documented using a monoclonal antibody specific for the post-translationally modified, acetylated α-tubulin found in the tail, but not oocyte, microtubules. Results showed that fertilization arrested at various levels: (1) metaphase II arrest; (2) arrest after successful incorporation of the spermatozoon; (3) arrest after formation of the sperm aster; (4) arrest during mitotic cell cycle progression; and (5) arrest during meiotic cell cycle progression. Rawe et al.99 analyzed the distribution of β-tubulins to detect spindle and cytoplasmic microtubules, α-acetylated tubulins for sperm microtubules and chromatin configuration in oocytes showing fertilization failure after conventional IVF or ICSI. Immunofluorescence analysis showed that the main reason for fertilization failure after IVF was no sperm penetration (55.5%). The remaining oocytes showed different abnormal patterns, e.g. oocyte activation failure (15.1%) and defects in pronuclei apposition (19.2%). On the other hand, fertilization failure after ICSI was mainly associated with incomplete oocyte activation (39.9%), and to a lesser extent with defects in pronuclei apposition (22.6%) and failure of sperm penetration (13.3%). A further 13.3% of the ICSI oocytes arrested their development at the metaphase of the first mitotic division. Fluorescent imaging scanning has shown that centrosomal defects may result in abnormal microtubule nucleation, preventing genomic union. In a primate model, ICSI (using apparently normal gametes) resulted in abnormal nuclear remodeling during sperm decondensation due to the presence of the sperm acrosome and perinuclear theca, structures normally removed at the oolema during IVF; this in turn caused a delay of DNA synthesis108. Such unusual modifications raised concerns about the ‘normalcy’ of the

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fertilization process and cell-cycle checkpoints during ICSI (reviewed in references 108 and 109). During the ICSI procedure, a spermatozoon is deposited into the ooplasm with both the acrosomal and plasma membranes intact, in addition to the other sperm components that are naturally eliminated in fertilized oocytes. The sperm acrosome contains a variety of hydrolytic enzymes, the release of which into the ooplasm might be harmful110. It is unclear how an oocyte that has been injected with an acrosome-intact spermatozoon will cope with the sperm acrosome. It is believed that an acrosome introduced into the ooplasm by ICSI seems physically to disturb sperm chromatin decondensation. Synthesis of DNA is delayed in both pronuclei when the paternal pronucleus is still undergoing decondensation in the apical region under the acrosomal cap, identifying a unique G1/S cell-cycle checkpoint111. Katayama et al.112 showed morphological characteristics in detail of the acrosome of boar sperm through ICSI, showing that the lectin-binding properties of sperm-head components introduced into the cytoplasm were different from those after IVF. Resumption of meiosis and cortical-granules exocytosis were achieved after micromanipulation techniques. Terada et al.113 assessed centrosomal function of human sperm using heterologous ICSI with rabbit eggs. They demonstrated that the spermaster formation rate was lower in infertile men compared with controls. Moreover, the spermaster formation rate correlated with the embryonic cleavage rate following human IVF. The data suggested that reproductive success during the first cell cycle requires a functional sperm centrosome and that dysfunctions of this organelle could be present in cases of unexplained infertility. Kovacic and Vlaisavljevic114 studied the microtubules and chromosomes of human oocytes failing to fertilize after ICSI, to establish how sperm chromatin and sperm-astral microtubule configuration is related to the phases of the oocyte cell cycle, and to find the defects in these structures causing fertilization arrest. A high proportion of

oocytes were arrested at metaphase II. Damage of the second meiotic spindle was noted in some oocytes. Intact sperm were found in some cases, and a swollen sperm head and prematurely condensed sperm chromosomes were apparent in others. Many monopronucleate oocytes contained sperm, with delay in the process of sperm nucleus decondensation. It was concluded that sperm that do not activate the oocyte may continue decondensing the chromatin, but the oocyte prevents male pronucleus formation before the female one, mostly by causing premature chromatin condensation in the sperm and by duplicating the sperm centrosome. The functional role of the sperm tail (either attached or dissected) in early human embryonic growth is not known. In microinjection experiments, it was demonstrated that the injection of isolated sperm segments (heads or flagella) could permit oocyte activation and bipronuclear formation. However, a high rate of mosaicism was observed in the embryos with disrupted sperm, suggesting that the structural integrity of the intact fertilizing spermatozoon appears to contribute to normal human embryogenesis115. In addition, oocytes injected with mechanically dissected spermatozoa, although capable of pronuclear formation, did not undergo normal mitotic division. The lack of a bipolar spindle, in combination with mosaicism, suggested abnormalities of the mitotic apparatus when sperm integrity is impaired following dissection116. Fertilization is completed once the parental genomes unite, and requires migration of the egg nucleus to the sperm nucleus (female and male pronuclei, respectively) on microtubules within the inseminated egg. The failure of zygotic development in some patients suggests that abnormalities of this step may contribute to infertility. Recently, Payne et al.59 showed that preferentially localized dynein and perinuclear dynactin associate with the nuclear pore complex and vimentin, and are required to mediate genomic union. The data suggest a model in which dynein accumulates and binds to the female pronucleus on sperm-aster

CONTRIBUTION OF MALE GAMETE TO FERTILIZATION AND EMBRYOGENESIS

microtubules, where it acts with dynactin, nucleoporins and vimentin. Mutations in the human gene ubiquitin-specific protease-9 Y chromosome (USP9Y), which encodes a protein with a C-terminal ubiquitin hydrolase domain, result in azoospermia and male infertility117. Knock-out mice lacking the E3 ubiquitin protein ligase SIAH1A or the E2 ubiquitin-conjugating enzyme HR6B demonstrated defects in meiosis, postmeiotic germ cell development and male infertility118. Ubiquitin-mediated proteolysis is also critical for other aspects of reproduction, including the elimination of defective sperm in the epididymis, clearance of paternal mitochondria and progression of embryonic development in mammals119. Sutovsky et al.101 showed that increased sperm ubiquitin (measured through a flow cytometric sperm–ubiquitin tag immunoassay) was inversely correlated with sperm quality. Conversely, Muratori et al.120 observed a positive correlation between sperm ubiquitination and sperm quality. More studies are therefore needed to establish whether sperm ubiquitination can be used as a biomarker of sperm functional capacity and whether anomalies of fertilization result from anomalies of ubiquitin sperm marking. Ubiquitin-mediated degradation targets cellcycle regulators for proteolysis. Cullins are core components of E3 ubiquitin ligases, and CUL-4A has a possible role in cell cycle control. In experiments with CUL-4A deletion mutations in mice, it was observed that homozygous mutants generated no viable pups or recovery of homozygous embryos after 7.5 days postcoitum119. Results indicated that appropriate CUL-4A expression appears to be critical for early embryonic development. The true identity of ubiquitinated substrates in the sperm mitochondria is not known. Nevertheless, it was recently shown that prohibitin, a mitochondrial membrane protein, is one of the ubiquitinated substrates that makes the sperm mitochondria responsible for the egg’s ubiquitin–proteasome-dependent proteolytic machinery

after fertilization121. Abnormalities of this recognition system might be involved in the dysregulation of mitochondrial inheritance and sperm quality control. Occasional occurrence of paternal inheritance of mtDNA has been suggested in mammals, including humans. While most such evidence has been widely disputed, of particular concern is the documented heteroplasmic or mixed mtDNA inheritance after ooplasmic transfusion122. Indeed, there is evidence that heteroplasmy is a direct consequence of ooplasm transfer, a technique that was used to ‘rescue’ oocytes from older women by injecting ooplasm from young oocytes. ICSI has an inherent potential for delaying the degradation of sperm mitochondria. However, paternal mtDNA inheritance after ICSI has not been documented (reviewed in reference 101).

Putative dysfunctions resulting from aberrant delivery of mRNA Recently, mRNA has been discovered in human ejaculated sperm. A non-exhaustive list of transcripts, including c-myc, human leukocyte antigen (HLA) class 1, protamines 1 and 2, heat shock proteins 70 and 90, β-integrins, transition protein-1, β-actin, variants of phosphodiesterase, progesterone receptor and aromatase, reveals a wide range of transcripts in mammalian sperm123–126. In mammals, round spermatids contain a number of transcripts that are produced either throughout early spermatogenesis127 or during spermiogenesis from the haploid gene encoding sperm-specific proteins such as transition proteins and protamines128, or sperm-tail cytoskeletal proteins implied in the molecular make-up of the outer dense fibers129 and fibrous sheath130. The arrest of transcription that is concomitant with major changes in chromatin organization occurs during mid-spermiogenesis131. However, the presence of extremely varied transcripts in mature sperm cells has been described in both rodent132,133 and human spermatozoa134–138.

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Most investigations into RNA identification in mature spermatozoa have been performed with techniques based on the detection of specific or particular sets of RNA by means of polymerase chain reaction (PCR) after reverse transcription (RT-PCR). Indeed, nested RT-PCR of RNA from a single spermatozoon has shown apparently aberrant transcripts in human sperm cells, such as those encoding synapsin I, immunoglobulins or Y-cell receptor α139. Such a phenomenon, named illegitimated transcription, has been defined as a very low-level transcription of any gene in any cell type140. Different mRNA species were found in human ejaculated spermatozoa by carrying out a step-bystep analysis with macroarray hybridization, RTPCR and in situ hybridization. An extended pattern of several transcripts encoding factors (NFκB, HOX2A, ICSBP, JNK2, HBEGF, RXRβ and ErbB3) essential for cellular functioning (including signal transduction and cell proliferation) were demonstrated in human sperm nuclei. The presence of residual DNA and RNA polymerase activity within the sperm chromatin was also formerly reported141–143. Complementary investigations have indicated that, in spite of a high degree of DNA packaging within the human sperm head, chromatin retains some features of active chromatin, mainly acetylated histones144 and the arrangement of certain chromatin domains into nucleosomes145,146. The existence of transcriptional and translational activities in human sperm during capacitation and the acrosome reaction has been described, which could also explain the presence of mRNA in mature sperm147. Lambard et al.124 showed a significant decrease of aromatase mRNA level in sperm with low motility, compared with highly motile sperm from the same sample of normospermic patients; these data suggest that the establishment of sperm mRNA profiles could be used as a genetic fingerprint of normal fertile men. The data therefore suggest that spermatozoa are a repository of information regarding meiotic and postmeiotic gene expression in the human,

and are likely to contain transcripts for genes playing an essential role during spermiogenesis (Figure 4.2). Use of the whole ejaculate as a wholly noninvasive biopsy of the spermatid should therefore be evaluated123. Different mRNA-encoding proteins are probably implicated in cell–cell and cell–substratum interactions, enhancement of fertility rate, lipid transportation, membrane recycling and stabilization of stress proteins, and promotion or inhibition of the death cell mechanism148. It is possible that if the mRNA accumulated in the sperm nucleus is not residual non-functional material, it might be viewed as the male gamete’s contribution to early embryogenesis149. Delivering spermatozoon RNA to the oocyte has been demonstrated in mice150 and humans148. Some sperm transcripts encoding proteins known to participate in fertilization and embryonic development have been specifically detected in early embryos after in vitro fertilization failure, while they have not been found in the oocyte138. Thus, human spermatozoa could act not only as genome carriers but also as providers of specific transcripts necessary for zygote viability and development before activation of the embryonic genome.

+Clusterin

HSF2

MID

Calmegin

HSPA1L

–NLVCF

+AKAP4

DNAJB1

CYR61

–Oscillin

+HSBR1(CDH13)

EYA3

+Protamine-2

DUSP5

+FOXG1B

–RPL2

+WNT5A –WHSC1 –SOX13

Figure 4.2 Spermatozoa mRNA transcripts and putative temporal expression during embryo development (+ refers to mRNA transcripts that are possibly involved in development as reported in reference 148)

CONTRIBUTION OF MALE GAMETE TO FERTILIZATION AND EMBRYOGENESIS

Ostermeier et al.102 recently reported a suite of novel human spermatozoal mRNAs. The authors identified a group of RNAs previously defined as micro-RNAs, and others that were antisense mRNAs of in silico predicted transcripts (or silencing mRNAs). The authors speculated that the delivery of these antisense RNAs upon fertilization could enable their participation in early postfertilization processes. They could be involved in regulation of the transition from maternal to embryonic genome, and could even be related to imprinting. Fukagawa et al.151 and Morris et al.152 have shown that this class of mRNA could confer transcriptional silencing by methylation.

Aberrant embryogenesis secondary to nuclear/chromatin anomalies As mentioned above, spermatozoa of infertile men have been shown to contain various nuclear alterations, including an abnormal chromatin structure, aneuploidy, chromosomal microdeletions and DNA strand breaks (reviewed in reference 2). Since meiosis is crucial for the survival of a species, an elaborate series of safeguards have evolved to pair, break and repair the chromosomal DNA. Despite such regulatory mechanisms, it is well known that translocations and aneuploidy are regularly introduced during the meiotic divisions. Esterhuizen et al.153 evaluated the role of chromatin packaging (CMA3 staining), sperm morphology during sperm–zona binding, sperm decondensation and the presence of polar bodies in oocytes that failed IVF. Odds ratio analyses indicated that being in the ≥ 60% CMA3 staining group resulted in a 15.6-fold increase in the risk of decondensation failure, relative to CMA3 staining of < 44%. Using CMA3 fluorescence to discriminate, 51% of oocytes in the group with elevated CMA3 fluorescence had no sperm in the ooplasm, compared with 32% and 16% penetration failure in the CMA3 staining groups ≥ 44–59% and < 44%, respectively. Sperm chromatin packaging quality and sperm morphology assessments were

demonstrated as useful clinical indicators of human fertilization failure. Ectopic expression and inactivation of apoptosis-related genes have been shown to cause abnormalities in spermatogenesis. During spermatogenesis, the process of germ cell proliferation and maturation causes diploid spermatogonia to develop into mature haploid sperm. A number of the developing germ cells die by apoptosis before reaching maturity, even under normal conditions154. In addition to the physiological germ-cell apoptosis that occurs continuously throughout life, increased germ-cell apoptosis results from such external disturbances as irradiation or exposure to toxicants155. Evidence suggests that within the cellular component of the testicular tissue, caspases play a central role in the apoptotic process that leads to DNA fragmentation of Sertoli cells104. The presence of apoptosis in ejaculated spermatozoa could be the result of various types of injuries156,157. In vivo, apoptosis could be triggered at the testicular (hormonal depletion, irradiation, toxic agents, chemicals and heat have been shown to induce apoptosis), epididymal (the result of signals released by abnormal and/or senescent spermatozoa or by leukocytes – such as ROS and other mediators of inflammation/infection) or seminal (ROS, lack of antioxidants or other causes) levels. Also, apoptosis could be triggered by factors present in the female tract. In vitro, apoptosis could be triggered upon incubation with inappropriate culture media or other manipulation procedures. Irrespective of the stimulus, spermatozoa undergoing apoptosis and unrecognized by currently used methodologies may be dysfunctional (resulting in failure of fertilization) or, more dramatically, they may pose the risk of carrying a damaged genome into the egg resulting in poor embryo development, miscarriage or childhood anomalies158,159. We have published compelling evidence indicative of the presence of somatic cell apoptosis markers, including key constituents of the apoptotic machinery and activation upon defined

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stimuli, in human ejaculated spermatozoa. It can be summarized as follows: • Human spermatozoa exhibit somatic cell apoptosis markers. Spermatozoa from fertile and infertile men demonstrated variable levels of phosphatidylserine (PS) externalization (by Annexin V-FITC (fluorescein isothiocyanate) binding using indirect immunofluorescence) and DNA fragmentation (by immunofluorescence using TUNEL (terminal deoxy nucleotide transferase-mediated dUTP nickend labeling) and also the monoclonal antibody (mAb) F7-26) upon ejaculation and incubation under capacitating conditions2,160,161. • The apoptosis markers PS externalization and DNA fragmentation are expressed with a higher frequency in fractions of sperm with low motility (where dysmorphic and dysfunctional sperm are found), when compared with high-motility fractions2,161. • Apoptosis markers are expressed with a significantly higher frequency in sperm of infertile men when compared with fertile controls156,161. • Human sperm contain caspase-3, the major executioner caspase, in both inactive and active forms. We have unequivocally demonstrated the presence of inactive caspase-3 (32 kDa) and also caspase-3 activation (17-kDa proteolytic fragment) in ejaculated sperm by immunoblotting, and have also confirmed caspase activation by immunofluorescent and enzymatic techniques161. Using immunofluorescence with a FITC-labeled antibody that specifically recognizes the active form, active caspase-3 was exclusively detected to the midpiece, where mitochondria and residual cytoplasm are present. • Human sperm exhibit other members of the caspase family, caspase-7 and -9. By immunoblotting, we have demonstrated the presence of inactive caspase-7 (35 kDa) and caspase-9

(45 kDa) in many samples, as well as active caspase-7 (32 kDa) and caspase-9 (37 kDa) in samples of infertile men162. • Human sperm possess apoptosis-inducing factor (AIF). By immunoblotting, we have demonstrated that human sperm express AIF (67 kDa) (although further studies are needed to establish its cellular location) and possibly a unique PARP (poly [ADP-ribose] polymerase), a specific caspase substrate of 66 kDa, with a different molecular weight from that of the 116–85-kDa analog and proteolytic fragment found in somatic cells162,163. • Human sperm appear not to express Bid protein (neither the 24-kDa intact nor the 15-kDa proapoptotic fragment) as measured by immunoblotting (unpublished observations). • Caspase activation can be triggered in ejaculated human sperm by the mitochondrial disrupter staurosporine. Staurosporine at 10 µmol/l (apoptosis-inducing dose in somatic cells) significantly enhanced caspase activation (by DEVD assay (Asp-Glu-Val-Asp) that measures caspase-3, -6 and -7) and DNA fragmentation, suggesting a mitochondriadependent pathway of caspase activation162. We analyzed the dose-dependent effect of staurosporine on sperm viability, and found no deleterious effects in the range 1–15 µmol/l. Preincubation with the pan-caspase inhibitor zVAD (benzoxy-Val-Ala-Asp) (50 µmol/l, 30 min) inhibited staurosporine-induced DNA fragmentation by 50% (unpublished observations). • Human sperm did not exhibit a response to Fas ligand. Fas ligand did not trigger caspase activation, PS translocation or DNA fragmentation. The Fas ligand (anti-Fas monoclonal antibody) was tested at 1 µg/ml (apoptosisinducing dose in somatic cells) with and without G-protein as a linker (at 2 µg/ml), and did not elicit caspase activation, PS translocation or DNA fragmentation162. These data are in

CONTRIBUTION OF MALE GAMETE TO FERTILIZATION AND EMBRYOGENESIS

agreement with recent studies that failed to demonstrate Fas receptors in ejaculated human sperm164. • Hydrogen peroxide, the most damaging ROS in sperm, induces expression of apoptosis markers. We demonstrated that H2O2 increased PS translocation and DNA fragmentation165. H2O2 produced a dose-dependent effect on PS translocation, with a significant increase at 200 µmol/l, a dose that we previously reported initiated impairment of motility and other sperm functions without affecting viability in vitro166. In addition, H2O2 resulted in a moderate increase in caspase activation. • Ejaculated human sperm show a strong correlation between ROS production and DNA fragmentation, linking mitochondrial dysfunction and expression of apoptosis markers. We have shown a positive, significant correlation between the endogenous generation of ROS (measured by chemiluminescence) and DNA strand breaks in ejaculated sperm2. • Ejaculated sperm show a strong correlation between disruption of mitochondrial transmembrane potential and PS translocation, again linking mitochondrial dysfunction and expression of apoptosis markers. We have documented that samples with live cells presenting PS externalization demonstrated changes in mitochondrial transmembrane potential using a mitochondrial membrane sensor kit167. The test uses a cationic dye, which fluoresces differently in apoptotic and healthy cells. Results showed alterations of the mitochondrial membrane potential that were three times higher in sperm fractions with low motility, compared with high-motility fractions. The oocyte has the capability to repair DNA damage, as oocytes fertilized by DNA-damaged spermatozoa did not develop further in vitro when they were cultured in the presence of inhibitors to DNA repair168–171. The capacity of the oocyte to repair is limited, and is related to the degree of

sperm DNA damage. The fertilization capacity of apoptotic sperm has been observed to be at the same rate as that of intact spermatozoa; however, embryo development to the blastocyst stage is closely related to the integrity of the DNA171. During spermatogenesis, a complex and dynamic process of proliferation and differentiation occurs as spermatogonia are transformed into mature spermatozoa. This unique process involves a series of meioses and mitoses, changes in cytoplasmic architecture, replacement of somatic celllike histones with transition proteins and the final addition of protamines, leading to highly packaged chromatin172. The human is of particular interest, as a single ejaculate normally contains a heterogeneous population of spermatozoa. It has been known for many years that the chromatin of the mature sperm nucleus can be abnormally packaged173. In addition, abnormal chromatin packaging and nuclear DNA damage appear to be linked174, and there is a strong association between the presence of nuclear DNA damage in the mature spermatozoa of men and poor semen parameters77,175. It is postulated that an endogenous nuclease, topoisomerase II, creates and ligates nicks to provide relief of torsional stress and to aid chromatin rearrangement during protamination176. The DNA damage in ejaculated human sperm consists of both single- and double-stranded DNA breaks. Endogenous nicks in DNA are normally expressed at specific stages of spermiogenesis in different animal models; these endogenous nicks are evident during spermiogenesis, but are not observed once chromatin packaging is completed. It is possible that endogenous nuclease topoisomerase II may play a role in both creating and ligating nicks during spermiogenesis, that these nicks may provide relief of torsional stress and that they aid chromatin rearrangement during the displacement of histones by protamines177–179. Several studies have shown that sperm DNA quality has robust power to predict fertilization in vitro175,180–182. Tomlinson et al.183 have reported that the only parameter showing a significant

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difference between pregnant and non-pregnant groups in IVF was the percentage of DNA fragmentation assessed by in situ nick translation. Sperm-derived effects, particularly the degree of DNA fragmentation, have been suggested to affect human embryo development104. The sperm chromatin structure assay (SCSA) has been proposed as a diagnostic tool to predict fertilization by evaluating sperm DNA stability79. The SCSA measures susceptibility to DNA denaturation in situ in sperm exposed to acid for 30 s, followed by acridine orange staining. The use of flow cytometry in the SCSA increases its dependability. Duran et al.84 studied a large infertility population undergoing IUI therapy in a prospective cohort fashion. A total of 119 patients underwent 154 cycles of IUI. DNA fragmentation evaluated by TUNEL and acridine orange staining were measured. The authors reported that sperm DNA quality played a major role as a predictor of pregnancy under such in vivo conditions.

Epigenetic factors ‘Epigenetics’ refers to a process that regulates gene activity without affecting the genetic (DNA) code and is heritable through cell division. Germ cell development and early embryogenesis are crucial windows in the erasure, acquisition and maintenance of genomic imprints. Moreover, a number of genes regulated by imprinting have been shown to be essential to fetal growth and placental function. Increasing attention has recently focused on potential epigenetic disturbances resulting from embryo culture, somatic cell nuclear cloning and assisted reproductive technologies184,185, indicating that a better understanding of genomic imprinting or parent-of-origin effects on gene expression is highly significant to the current study of reproduction and development. Imprinting is an epigenetically controlled phenomenon, because something other than DNA sequence must distinguish the parental alleles and determine sex-specific gene expression. The role of

DNA methylation in genomic imprinting has been extensively investigated. It is estimated that the total number of imprinted genes in the mouse and human genomes may range between 100 and 200. Of those that have been identified to date, a significant number appear to have important roles in fetal development. It has been argued that imprinted genes play essential roles in controlling the placental supply of maternal nutrients to the fetus, by regulating the growth of the placenta and/or the activity of transplacental transport systems. Methylation is important for somatic cell maintenance of imprinting after the global wave of demethylation in the blastocyst186. However, the question arises of how maternal and paternal alleles can be distinguished after global demethylation arises187,188. It has been found that different methylation sites within imprinted genes may demonstrate significant temporal differences in methylation pattern, and that establishment of the final methylation pattern is a dynamic process189. Epigenetic modifications serve as an extension of the information content by which the underlying genetic code may be interpreted. These modifications mark genomic regions and act as heritable and stable instructions for the specification of chromatin organization and structure that dictate transcriptional states. In mammals, DNA methylation and the modification of histones account for the major epigenetic alterations. Two cycles of DNA methylation reprogramming have been characterized (reviewed in reference 190). During germ cell development, epigenetic reprogramming of DNA methylation resets parent-of-origin-based genomic imprints and restores totipotency to gametes. During fertilization, the second cycle is triggered, resulting in an asymmetric difference between parental genomes. Further epigenetic asymmetry is evident in the establishment of the first two lineages at the blastocyst stage. This differentiative event sets the epigenetic characteristics of the lineages as derivatives of the inner cell mass (somatic) and trophectoderm (extraembryonic).

CONTRIBUTION OF MALE GAMETE TO FERTILIZATION AND EMBRYOGENESIS

The erasure and subsequent retracing of the epigenetic checkpoints pose the most serious obstacles to somatic nuclear transfer. Elaboration of the mechanisms of these interactions will be invaluable in our fundamental understanding of biological processes and in achieving substantial therapeutic advances190. Recent studies have suggested a possible link between human assisted reproductive technologies and genomic imprinting disorders (reviewed in reference 191). The presence of Angelman syndrome (caused by a loss of function of the maternal allele or duplication of the paternal allele within a region that spans UBE3A) and Beckwith–Wiedemann syndrome (another disease that exhibits parent-of-origin effects in its inheritance) has been observed following the use of ICSI. Assisted reproductive technologies include the isolation, handling and culture of gametes and early embryos at times when imprinted genes are likely to be particularly vulnerable to external influences. Evidence of sex-specific differences in imprint acquisition suggests that male and female germ cells may be susceptible to perturbations in imprinted genes at specific prenatal and postnatal stages. Imprints acquired first during gametogenesis must be maintained during preimplantation development when reprogramming of the overall genome occurs. The identification of the mechanisms and timing of imprint erasure, acquisition and maintenance during germ cell development and early embryogenesis, as well as their implications for future epigenetic studies in assisted reproductive technologies, should constitute research priorities191.

CONCLUSIONS The fertilizing spermatozoon has a very dynamic and critical participation in embryogenesis during and after the fertilization process. A defective spermatozoon that penetrates the oocyte may cause arrest of development at multiple levels during embryo preimplantational development.

Moreover, sublethal and lethal effects can be ‘carried over’ following implantation, resulting in human disease. The contributions of the fertilizing spermatozoon to the oocyte during normal development include delivery of the DNA/chromatin, the oocyte-activating factor (OAF) and a centriole. The DNA/chromatin complex is obviously the most significant contribution to originating a new diploid individual. Nevertheless, the OAF and centriole play a critical part in bringing about oocyte activation and the first mitotic division, and without their contributions embryogenesis would also be neglected. In addition, recent data have indicated that spermatozoa provide the zygote with a unique suite of paternal mRNAs. Such transcripts might be crucial for early and late embryonic development, and deficient delivery or aberrant transcription might contribute to abnormal development and arrest. A large body of evidence is accumulating demonstrating that abnormal oocyte activation and embryonic development might be the consequence of aberrant paternal contribution(s). An early paternal effect results in failure to complete the fertilization process, syngamy or early cleavage. It can be demonstrated by morphological abnormalities observed at the pronuclear and 2–4cell stage. It is speculated that these defects are mediated by sperm deficiencies, including an abnormal release of OAF and by dysfunctions of the centrosome and cytoskeletal apparatus. A late paternal effect is characterized by failure to achieve implantation competence, but could also be associated with pregnancy loss and postnatal developmental abnormalities. It is associated with sperm nuclear/chromatin defects, including the presence of aneuploidy, genetic anomalies, DNA damage and possible other causes. The strictly maternal inheritance of mitochondrial DNA (mtDNA) in mammals is a developmental paradox promoted by an unknown mechanism responsible for the destruction of sperm mitochondria shortly after fertilization. It has been shown that sperm mitochondria are tagged

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and later subjected to directed proteolysis during preimplantation development. Abnormalities of this process could lead to aberrant embryogenesis. In addition, recent data have indicated that spermatozoa provide the zygote with a unique suite of paternal mRNAs. Such transcripts might be crucial for early and late embryonic development, and deficient delivery or aberrant transcription might lead to abnormal embryogenesis. Furthermore, limited RNA synthesis can be detected in human pronuclei and failure of this early transcription is associated with abnormal pronuclear development and arrest. Finally, gene-imprinting abnormalities, either of gamete origin or taking place during early embyogenesis, may be responsible for severe human disease. Such a problem has a potential impact when using certain forms of assisted reproductive technologies.

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41. Ford WCL, Rees JM. The bioenergetics of mammalian sperm motility. In Gagnon C, ed. Controls of Sperm Motility: Biological and Clinical Aspects. Boca Raton, FL: CRC Press, 1990: 175 42. Max B. This and that: hair pigments, the hypoxic basis of life and the Virgilian journey of the spermatozoon. Trends Pharmacol Sci 1992; 13: 272 43. Cummins J. Mitochondrial DNA in mammalian reproduction. Rev Reprod 1998; 3: 172 44. Short RV. The difference between a testis and an ovary. J Exp Zool 1998; 281: 359 45. Kaneda H, et al. Elimination of paternal mitochondrial DNA in intraspecific crosses during early mouse embryogenesis. Proc Natl Acad Sci USA 1995; 92: 4542 46. Ankel-Simons F, Cummins JM. Misconceptions about mitochondria and mammalian fertilization – implications for theories on human evolution. Proc Nat Acad Sci USA 1996; 93: 13859 47. Aitken RJ. Free radicals, lipid peroxidation and sperm function. Reprod Fertil Dev 1995; 7: 659 48. Shalgi R, et al. Fate of sperm organelles during early embryogenesis in the rat. Mol Reprod Dev 1994; 37: 264 49. Sutovsky P, Navara C, Schatten G. Fate of the sperm mitochondria, and the incorporation, conversion, and disassembly of the sperm tail structures during bovine fertilization. Biol Reprod 1996; 55: 1195 50. Smith LC, Alcívar AA. Cytoplasmic inheritance and its effects on development and performance. J Reprod Fertil 1993; 48 (Suppl): 31 51. Gresson RAR. Presence of the sperm middle-piece in the fertilized egg of the mouse (Mus musculus). Nature 1940; 145: 425 52. Smith LC, et al. Sea urchin genes expressed in activated coelomocytes are identified by expressed sequence tags. Complement homologues and other putative immune response genes suggest immune system homology within the deuterostomes. J Immunol 1996; 15: 593 53. Allen V. Role of motor proteins in organizing the endoplasmic reticulum and Golgi apparatus. Semin Cell Dev Biol 1996; 7: 335 54. Sutovsky P, et al. Ubiquitin tag for sperm mitochondria. Nature 1999; 402: 371 55. Sutovsky P, et al. Ubiquitinated sperm mitochondria selective proteolysis and the regulation of mitochondrial inheritance in mammalian embryos. Biol Reprod 2000; 63: 582 56. Ingman M, et al. Mitochondrial genome variation and the origin of modern humans. Nature 2000; 408: 708

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57. Schatten G. The centrosome and its mode of inheritance: the reduction of the centrosome during gametogenesis and its restoration during fertilization. Dev Biol 1994; 165: 299 58. Reinsch S, Karsenti E. Movement of nuclei along microtubules in Xenopus egg extracts. Curr Biol 1997; 7: 211 59. Payne C, et al. Preferentially localized dynein and perinuclear dynactin associate with nuclear pore complex proteins to mediate genomic union during mammalian fertilization. J Cell Sci 2003; 116: 4727 60. Oehninger S, et al. Failure of fertilization in in vitro fertilization: the ‘occult’ male factor. J In Vitro Fert Embryo Transf 1988; 5: 181 61. Oehninger S, et al. Delayed fertilization during in vitro fertilization and embryo transfer cycles: analysis of causes and impact on overall results. Fertil Steril 1989; 52: 991 62. Kruger TF, et al. Predictive value of abnormal sperm morphology in in vitro fertilization. Fertil Steril 1988; 49: 112 63. Ron-el R, et al. Delayed fertilization and poor embryonic development associated with impaired semen quality. Fertil Steril 1991; 55: 338 64. Parinaud J, et al. Influence of sperm parameters on embryo quality. Fertil Steril 1993; 60: 888 65. Grow DR, et al. Sperm morphology as diagnosed by strict criteria: probing the impact of teratozoospermia on fertilization rate and pregnancy outcome in a large in vitro fertilization population. Fertil Steril 1994; 62: 559 66. Oehninger S, et al. Corrective measures and pregnancy outcome in in vitro fertilization in patients with severe sperm morphology abnormalities. Fertil Steril 1988; 50: 283 67. Janny L, Menezo YJ. Evidence for a strong paternal effect on human preimplantation embryo development and blastocyst formation. Mol Reprod Dev 1994; 38: 36 68. Oehninger S, et al. A comparative analysis of embryo implantation potential in patients with severe teratozoospermia undergoing in-vitro fertilization with a high insemination concentration or intracytoplasmic sperm injection. Hum Reprod 1996; 11: 1086 69. Nagy ZP, et al. The result of intracytoplasmic sperm injection is not related to any of the three basic sperm parameters. Hum Reprod 1995; 10: 1123

70. Oehninger S, et al. Intracytoplasmic sperm injection: achievement of high pregnancy rates in couples with severe male factor infertility is dependent primarily upon female and not male factors. Fertil Steril 1995; 64: 977 71. Mansour RT, et al. The effect of sperm parameters on the outcome of intracytoplasmic sperm injection. Fertil Steril 1995; 64: 982 72. Mercan R, et al. The outcome of clinical pregnancies following intracytoplasmic sperm injection is not affected by semen quality. Andrologia 1998; 30: 91 73. Tasdemir I, et al. Effect of abnormal sperm head morphology on the outcome of intracytoplasmic sperm injection in humans. Hum Reprod 1997; 12: 1214 74. Gorczyca W, et al. Presence of strand breaks and increased sensitivity of DNA in situ to denaturation in abnormal human sperm: analogy to apoptosis of somatic cells. Exp Cell Res 1993; 207: 202 75. Manicardi GC, et al. Presence of endogenous nicks in DNA of ejaculated human spermatozoa and its relationship to chromomycin A3 accessibility. Biol Reprod 1995; 52: 864 76. Hughes CM, et al. A comparison of baseline and induced DNA damage in human spermatozoa from fertile and infertile men, using a modified comet assay. Mol Hum Reprod 1996; 2: 613 77. Lopes S, et al. Sperm deoxyribonucleic acid fragmentation is increased in poor-quality semen samples and correlates with failed fertilization in intracytoplasmic sperm injection. Fertil Steril 1998; 69: 528 78. Aitken RJ, et al. Relative impact of oxidative stress on the functional competence and genomic integrity of human spermatozoa. Biol Reprod 1998; 59: 1037 79. Evenson DP, et al. Utility of the sperm chromatin structure assay as a diagnostic and prognostic tool in the human fertility clinic. Hum Reprod 1999; 14: 1039 80. Sakkas D, et al. Origin of DNA damage in ejaculated human spermatozoa. Rev Reprod 1999; 4: 31 81. Pfeffer J, et al. Aneuploidy frequencies in semen fractions from ten oligoasthenoteratozoospermic patients donating sperm for intracytoplasmic sperm injection. Fertil Steril 1999; 72: 472 82. Sakkas D, et al. Sperm nuclear DNA damage and altered chromatin structure: effect on fertilization and embryo development. Hum Reprod 1998; 13 (Suppl 4): 11

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83. Rubio C, et al. Incidence of sperm chromosomal abnormalities in a risk population: relationship with sperm quality and ICSI outcome. Hum Reprod 2001; 16: 2084 84. Duran EH, et al. Sperm DNA quality predicts intrauterine insemination outcome: a prospective cohort study. Hum Reprod 2002; 17: 3122 85. Virant-Klun I, Tomazevic T, Meden-Vrtovec H. Sperm single-stranded DNA, detected by acridine orange staining, reduces fertilization and quality of ICSI-derived embryos. J Assist Reprod Genet 2002; 19: 319 86. Burrello N, et al. Lower sperm aneuploidy frequency is associated with high pregnancy rates in ICSI programmes. Hum Reprod 2002; 18: 1371 87. Liu CH, et al. DNA fragmentation, mitochondrial dysfunction and chromosomal aneuploidy in the spermatozoa of oligoasthenoteratozoospermic males. J Assist Reprod Genet 2004; 21: 119 88. Virro MR, Larson-Cook KL, Evenson DP. Sperm chromatin structure assay (SCSA) parameters are related to fertilization, blastocyst development, and ongoing pregnancy in in vitro fertilization and intracytoplasmic sperm injection cycles. Fertil Steril 2004; 81: 1289 89. Petit FM, et al. Could sperm aneuploidy rate determination be used as a predictive test before intracytoplasmic sperm injection? J Androl 2005; 26: 235 90. Devroey P, Van Steirteghem A. A review of ten years experience of ICSI. Hum Reprod Update 2004; 10: 19 91. Foresta C, et al. Genetic abnormalities among severely oligospermic men who are candidates for intracytoplasmic sperm injection. J Clin Endocrinol Metab 2005; 90: 152 92. St John JC. Incorporating molecular screening techniques into the modern andrology laboratory. J Androl 1999; 20: 692 93. Brinkworth MH. Paternal transmission of genetic damage: findings in animals and humans. Int J Androl 2000; 23: 123 94. Lanzendorf SE, et al. Penetration of human spermatozoa through the zona pellucida of nonviable human oocytes. J Soc Gynecol Invest 1994; 1: 69 95. Oehninger S, et al. Clinical significance of human sperm–zona pellucida binding. Fertil Steril 1997; 67: 1121 96. Oehninger S. Normal fertilization. In El-Shafie M, et al., eds. An Atlas of the Ultrastructure of Human Oocytes – A Guide for Assisted Reproduction. London: Parthenon Publishing, 2000: 23

97. Liu de Y, Garrett C, Baker HW. Clinical application of sperm–oocyte interaction tests in in vitro fertilization–embryo transfer and intracytoplasmic sperm injection programs. Fertil Steril 2004; 82: 1251 98. Asch R, et al. The stages at which human fertilization arrests: microtubule and chromosome configurations in inseminated oocytes which failed to complete fertilization and development in humans. Hum Reprod 1995; 10: 1897 99. Rawe VY, et al. Cytoskeletal organization defects and abortive activation in human oocytes after IVF and ICSI failure. Mol Hum Reprod 2000; 6: 510 100. Hewitson L, Simerly CR, Schatten G. Fate of sperm components during assisted reproduction: implications for infertility. Hum Fertil (Camb) 2002; 5: 110 101. Sutovsky P, Hauser R, Sutovsky M. Increased levels of sperm ubiquitin correlate with semen quality in men from an andrology laboratory clinic population. Hum Reprod 2004; 19: 628 102. Ostermeier GC, et al. A suite of novel human spermatozoal RNAs. J Androl 2005; 26: 70 103. Braude P, Bolton V, Moore S. Human gene expression first occurs between the four- and eight-cell stages of preimplantation development. Nature 1988; 332: 459 104. Tesarik J, et al. In-vitro effects of FSH and testosterone withdrawal on caspase activation and DNA fragmentation in different cell types of human seminiferous epithelium. Hum Reprod 2002; 17: 1811 105. Tesarik J. The paternal effects on cell division in the human preimplantation embryo. Reprod Biomed Online 2005; 10: 370 106. Tesarik J, Greco E, Mendoza C. Late, but not early, paternal effect on human embryo development is related to sperm DNA fragmentation. Hum Reprod 2004; 19: 611 107. Greco E, et al. Efficient treatment of infertility due to sperm DNA damage by ICSI with testicular spermatozoa. Hum Reprod 2005; 20: 226 108. Hewitson L, Simerly CR, Schatten G. Cytoskeletal aspects of assisted fertilization. Semin Reprod Med 2000; 18: 151 109. Hewitson L, Simerly CR, Schatten G. ICSI, male pronuclear remodeling and cell cycle checkpoints. Adv Exp Med Biol 2003; 518: 199 110. Tesarik J, Mendoza C. In vitro fertilization by intracytoplasmic sperm injection. BioEssays 1999; 21: 791

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111. Ramalho-Santos J, et al. SNAREs in mammalian sperm: possible implications for fertilization. Dev Biol 2000; 223: 54 112. Katayama M, Koshida M, Miyake M. Fate of the acrosome in ooplasm in pigs after IVF and ICSI. Hum Reprod 2002; 17: 2657 113. Terada Y, et al. Centrosomal function assessment in human sperm using heterologous ICSI with rabbit eggs: a new male factor infertility assay. Mol Reprod Dev 2004; 67: 360 114. Kovacic B, Vlaisavljevic V. Configuration of maternal and paternal chromatin and pertaining microtubules in human oocytes failing to fertilize after intracytoplasmic sperm injection. Mol Reprod Dev 2000; 55: 197 115. Colombero LT, et al. The role of structural integrity of the fertilising spermatozoon in early human embryogenesis. Zygote 1999; 7: 157 116. Moomjy M, et al. Sperm integrity is critical for normal mitotic division and early embryonic development. Mol Hum Reprod 1999; 5: 836 117. Sun C, et al. An azoospermic man with a de novo point mutation in the Y-chromosomal gene USP9Y. Nat Genet 1999; 23: 429 118. Dickins RA, et al. The ubiquitin ligase component Siah1a is required for completion of meiosis I in male mice. Mol Cell Biol 2002; 22: 2294 119. Li B, Ruiz JC, Chun KT. CUL-4A is critical for early embryonic development. Mol Cell Biol 2002; 22: 4997 120. Muratori M, et al. Sperm ubiquitination positively correlates to normal morphology in human semen. Hum Reprod 2005; 20: 1035–43 121. Thompson WE, Ramalho-Santos J, Sutovsky P. Ubiquitination of prohibitin in mammalian sperm mitochondria: possible roles in the regulation of mitochondrial inheritance and sperm quality control. Biol Reprod 2003; 69: 254 122. Brenner CA, Kubisch HM, Pierce KE. Role of the mitochondrial genome in assisted reproductive technologies and embryonic stem cell-based therapeutic cloning. Reprod Fertil Dev 2004; 16: 743 123. Miller D. RNA in the ejaculate spermatozoon: a window into molecular events in spermatogenesis and a record of the unusual requirements of haploid gene expression and post-meiotic equilibration. Mol Hum Reprod 1997; 3: 669 124. Lambard S, et al. Expression of aromatase in human ejaculated spermatozoa: a putative marker of motility. Mol Hum Reprod 2003; 9: 117

125. Wykes SM, Miller D, Krawetz SA. Mammalian spermatozoal mRNAs: tools for the functional analysis of male gametes. J Submicrosc Cytol Pathol 2000; 32: 77 126. Dadoune JP, et al. Identification of transcripts by macroarrays, RT–PCR and in situ hybridization in human ejaculate spermatozoa. Mol Hum Reprod 2005; 11: 133 127. Eddy EM. Male germ cell gene expression. Recent Prog Horm Res 2002; 57: 103 128. Steger K. Transcriptional and translational regulation of gene expression in haploid spermatids. Anat Embryol (Berl) 1999; 199: 471 129. Petersen C, Fuzesi L, Hoyer-Fender S. Outer dense fibre proteins from human sperm tail: molecular cloning and expression analyses of two cDNA transcripts encoding proteins of approximately 70 kDa. Mol Hum Reprod 1999; 5: 627 130. Eddy EM, Toshimori K, O’Brien DA. Fibrous sheath of mammalian spermatozoa. Microsc Res Tech 2003; 61: 103 131. Dadoune JP. Expression of mammalian spermatozoal nucleoproteins. Microsc Res Tech 2003; 61: 56 132. Pessot CA, et al. Presence of RNA in the sperm nucleus. Biochem Biophys Res Commun 1989; 158: 272 133. Passananti C, et al. The product of Zfp59 (Mfg2), a mouse gene expressed at the spermatid stage of spermatogenesis, accumulates in spermatozoa nuclei. Cell Growth Differ 1995; 6: 1037 134. Kumar G, Patel D, Naz RK. c-myc mRNA is present in human sperm cells. Cell Mol Biol Res 1993; 39: 111 135. Miller D, et al. Differential RNA fingerprinting as a tool in the analysis of spermatozoal gene expression. Hum Reprod 1994; 9: 864 136. Goodwin LO, Karabinus DS, Pergolizzi RG. Presence of N-cadherin transcripts in mature spermatozoa. Mol Hum Reprod 2000; 6: 487 137. Carreau S, et al. Aromatase expression in male germ cells. J Steroid Biochem Mol Biol 2001; 79: 203 138. Ostermeier GC, et al. Spermatozoal RNA profiles of normal fertile men. Lancet 2002; 360: 772 139. Kimoto Y. A single human cell expresses all messenger ribonucleic acids: the arrow of time in a cell. Mol Gen Genet 1998; 258: 233 140. Chelly J et al. Illegitimate transcription: transcription of any gene in any cell type. Proc Natl Acad Sci USA 1989; 86: 2617 141. Hecht NB. A DNA polymerase isolated from bovine spermatozoa. J Reprod Fertil 1974; 41: 345

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142. Witkin SS, Korngold GC, Bendich A. Ribonuclease-sensitive DNA-synthesizing complex in human sperm heads and seminal fluid. Proc Natl Acad Sci USA 1975; 72: 3295 143. Miteva K, Valkov N, Goncharova-Peinoval J. Electron microscopic data for the presence of post-meiotic gene expression in isolated ram sperm chromatin. Cytobios 1995; 83: 85 144. Gatewood JM, et al. Isolation of four core histones from human sperm chromatin representing a minor subset of somatic histones. J Biol Chem 1990; 265: 20662 145. Gineitis AA, et al. Human sperm telomere-binding complex involves histone H2B and secures telomere membrane attachment. J Cell Biol 2000; 151: 1591 146. Zalenskaya IA, Bradbury EM, Zalensky AO. Chromatin structure of telomere domain in human sperm. Biochem Biophys Res Commun 2000; 279: 213 147. Naz RK. Effect of actinomycine D and cycloheximide on human sperm function. Arch Androl 1998; 41: 135 148. Ostermeier GC, et al. Reproductive biology: delivering spermatozoan RNA to the oocyte. Nature 2004; 429: 154 149. Dadoune JP, Siffroi JP, Alfonsi MF. Transcription in haploid male germ cells. Int Rev Cytol 2004; 237: 1 150. Hayashi S, et al. Mouse preimplantation embryos developed from oocytes injected with round spermatids or spermatozoa have similar but distinct patterns of early messenger RNA expression. Biol Reprod 2003; 69: 1170 151. Fukagawa T, et al. Dicer is essential for formation of the heterochromatin structure in vertebrate cells. Nat Cell Biol 2004; 6: 784 152. Morris KV, et al. Small interfering RNA-induced transcriptional gene silencing in human cells. Science 2004; 305: 1289 153. Esterhuizen AD, et al. Defective sperm decondensation: a cause for fertilization failure. Andrologia 2002; 34: 1 154. Billig H, et al. Apoptosis in testis germ cells: developmental changes in gonadotropin dependence and localization to selective tubule stages. Endocrinology 1995; 136: 5 155. Pentikainen V, Erkkila K, Dunkel L. Fas regulates germ cell apoptosis in the human testis in vitro. Am J Physiol 1999; 276: E310 156. Oehninger S, et al. Presence and significance of somatic cell apoptosis markers in human ejaculated spermatozoa. Reprod Biomed Online 2003; 7: 469

157. Oehninger S. Biochemical and functional characterization of the human zona pellucida. Reprod Biomed Online 2003; 7: 641 158. Bowen JR, et al. Medical and developmental outcome at 1 year for children conceived by intracytoplasmic sperm injection. Lancet 1998; 351: 1529 159. Bonduelle M, et al. Seven years of intracytoplasmic sperm injection and follow-up of 1987 subsequent children. Hum Reprod 1999; 14: 243 160. Schuffner A, et al. Effect of different incubation conditions on phosphatidylserine externalization and motion parameters of purified fractions of highly motile human spermatozoa. J Androl 2002; 23: 194 161. Weng S, et al. Caspase activity and apoptotic markers in ejaculated human sperm. Mol Hum Reprod 2002; 8: 984 162. Taylor SL, et al. Somatic cell apoptosis markers and pathways in human ejaculated sperm: potential utility as indicators of sperm quality. Mol Hum Reprod 2004; 10: 825 163. Benchoua A, et al. Active caspase-8 translocates into the nucleus of apoptotic cells to inactivate poly(ADP-ribose) polymerase-2. J Biol Chem 2002; 277: 34217 164. Castro A, et al. Absence of Fas protein detection by flow cytometry in human spermatozoa. Fertil Steril 2004; 81: 1019 165. Duru NK, et al. Cryopreservation–thawing of fractionated human spermatozoa is associated with membrane phosphatidylserine externalization and not DNA fragmentation. J Androl 2001; 22: 646 166. Oehninger S, et al. Effects of hydrogen peroxide on human spermatozoa. J Assist Reprod Genet 1995; 12: 41 167. Barroso G, et al. Mitochondrial membrane potential integrity and plasma membrane translocation of phosphatidylserine: a comparison of subpopulations of sperm with high and low motility from men consulting for infertility. Fertil Steril 2006; 85: 149 168. Ahmadi A, Ng SC. Destruction of protamine in human sperm inhibits sperm binding and penetration in the zona-free hamster penetration test but increases sperm head decondensation and male pronuclear formation in the hamster-ICSI assay. J Assist Reprod Genet 1999; 16: 128 169. Ahmadi A, Ng SC. Developmental capacity of damaged spermatozoa. Hum Reprod 1999; 14: 2279 170. Ahmadi A, Ng SC. Influence of sperm plasma membrane destruction on human sperm head

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decondensation and pronuclear formation. Arch Androl 1999; 42: 1 Ahmadi A, Ng SC. Fertilizing ability of DNA-damaged spermatozoa. J Exp Zool 1999; 284: 696 Poccia D. Remodeling of nucleoproteins during gametogenesis, fertilization, and early development. Int Rev Cytol 1986; 105: 1 Evenson DP, Darzynkiewicz Z, Relamed MR. Relation of mammalian sperm chromatin heterogeneity to fertility. Science 1980; 210: 1131 Bianchi PG, et al. Effect of deoxyribonucleic acid protamination on fluorochrome staining and in situ nick-translation of murine and human mature spermatozoa. Biol Reprod 1993; 49: 1083 Sun JG, Jurisicova A, Casper RF. Detection of deoxyribonucleic acid fragmentation in human sperm: correlation with fertilization in vitro. Biol Reprod 1997; 56: 602 McPherson SM, Longo FJ. Nicking of rat spermatid and spermatozoa DNA: possible involvement of DNA topoisomerase II. Dev Biol 1993; 158: 122 McPherson S, Longo FJ. Chromatin structure– function alterations during mammalian spermatogenesis: DNA nicking and repair in elongating spermatids. Eur J Histochem 1993; 37: 109 Sakkas D, et al. Relationship between the presence of endogenous nicks and sperm chromatin packaging in maturing and fertilizing mouse spermatozoa. Biol Reprod 1995; 52: 1149 Chen JL, Longo FJ. Expression and localization of DNA topoisomerase II during rat spermatogenesis. Mol Reprod Dev 1996; 45: 61 Duran EH, et al. A logistic regression model including DNA status and morphology of spermatozoa for prediction of fertilization in vitro. Hum Reprod 1998; 13: 1235

181. Larson KL, et al. Sperm chromatin structure assay parameters as predictors of failed pregnancy following assisted reproductive techniques. Hum Reprod 2000; 15: 1717 182. Chan PJ, et al. A simple comet assay for archived sperm correlates DNA fragmentation to reduced hyperactivation and penetration of zona-free hamster oocytes. Fertil Steril 2001; 75: 186 183. Tomlinson MJ, et al. Interrelationships between seminal parameters and sperm nuclear DNA damage before and after density gradient centrifugation: implications for assisted conception. Hum Reprod 2001; 16: 2160 184. De Rycke M, Liebaers I, Van Steirteghem A. Epigenetic risks related to assisted reproductive technologies: risk analysis and epigenetic inheritance. Hum Reprod 2002; 17: 2487 185. Gosden R, et al. Rare congenital disorders, imprinted genes, and assisted reproductive technology. Lancet 2003; 361: 1975 186. Bartolomei MS, Tilghman SM. Genomic imprinting in mammals. Annu Rev Genet 1997; 31: 493 187. Barlow DP. Gametics imprinting in mammals. Science 1995; 270: 1610 188. Jaenisch R. DNA methylation and imprinting: why bother? Trends Genet 1997; 13: 323 189. Shemer R, et al. Dynamic methylation adjustment and counting as part of imprinting mechanisms. Proc Natl Acad Sci USA 1996; 93: 6371 190. Santos F, Dean W. Epigenetic reprogramming during early development in mammals. Reproduction 2004; 127: 643 191. Lucifero D, Chaillet JR, Trasler JM. Potential significance of genomic imprinting defects for reproduction and assisted reproductive technology. Hum Reprod Update 2004; 10: 3

5 Genome architecture in human sperm cells: possible implications for male infertility and prediction of pregnancy outcome Olga Mudrak, Andrei Zalensky

INTRODUCTION

unattended class of sperm chromosome abnormalities may have an impact on fertilization and early development. These aberrations are connected with chromosome packaging and the higher-order chromosome architecture in sperm nuclei.

Infertility and birth defects are often the result of chromosomal abnormalities in gametes1–3, with more than 80% of cases being paternally derived4. The development of multicolor fluorescence in situ hybridization (FISH) has allowed detection and analysis of several types of chromosomal defects in sperm, such as aneuploidies, partial chromosomal duplications, deletions and inversions, translocations and chromosomal breaks2,5–7. While there is consensus concerning a strong correlation between sperm chromosomal abnormalities and male infertility, the analysis of such abnormalities does not guarantee the selection of a ‘good spermatozoon’ without chromosomal defects, especially if intracytoplasmic sperm injection (ICSI) is performed for male factor infertility. There is no doubt that ICSI can enable men with severely impaired sperm to overcome naturally existing barriers to fertilization, yet in doing so it increases the possibility of transmitting genetic defects to the offspring. For example, it was demonstrated that oligozoospermic men carry a higher burden of transmissible chromosome damage8. A common attitude is emerging that detailed molecular cytogenetic tests should be performed on sperm samples from men with abnormal fertility before the execution of ICSI9–11. Here, we put forward a hypothesis that yet another previously

GENOME ARCHITECTURE More than a century ago, Rabl and Boveri proposed the existence of spatial order within the cell nucleus, which is manifested in the preservation of distinct individuality chromosomes in interphase12,13. Nevertheless, until recently, the view prevailed that interphase chromosomes were chromatin ‘spaghetti’ floating randomly in the nucleoplasm14. According to the modern assumption, the ordered and dynamic global architecture of interphase chromosomes exists, and is involved in a variety of nuclear functions (for recent reviews see references 15 and 16). This view resulted from a major breakthrough in the elucidation of chromosome organization that became possible because of FISH techniques, the development of instrumentation for microscopy and completion of the Human Genome Project. The central postulate of this concept is chromosome territorial organization. Interphase chromosomes occupy distinct non-overlapping intranuclear volumes called chromosome 73

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territories (CTs)16,17. We refer here to the higherorder spatial arrangement of CTs within the nuclear volume as genome architecture (GA). Two major characteristics of GA may be distinguished18: chromosome positioning (spatial localization of chromosomes relative to each other or to defined nuclear structures), and chromosome path (chromosome trajectory within nuclei). It appears that intranuclear positioning of CTs in interphase is non-random19. The spatial positioning of a chromosome relative to the center of the nucleus is defined as radial positioning20–22. A number of studies indicate that gene-rich chromosomes are located closer to the nuclear center, while gene-poor chromosomes are preferentially found at the nuclear periphery23,24. In addition to the radial positioning, chromosomes may be localized non-randomly with respect to each other21,25. For example, some authors declare fixed, deliberate chromosome positioning in the prometaphase ring26,27, while another study did not establish such an order in relative chromosome position28. Therefore, this issue is controversial. While dynamic changes in the relative spatial grouping of chromosome domains have been observed during cell-cycle progression, differentiation and malignant transformation29–31, the internal organization of CTs is still largely unknown. Recent studies indicate a relationship between the nuclear arrangement of CTs and the G–R-banding patterns of mitotic chromosomes32. In interphase nuclei, the R-band sequences, which are enriched in constitutively expressed housekeeping genes, are directed towards the nuclear interior. Current studies are focused on elucidation of the higher-order chromatin structures/ chromosome paths within CTs33 and relative spatial arrangement of individual CTs34.

Chromosome territories and chromosome architecture in sperm cells The sperm cell is a highly differentiated cell type, which results from the specialized genetic and

morphological process of spermatogenesis. During postmeiotic stages (spermiogenesis), the somatic histones are gradually replaced with protamines35,36. Consequently, the chromatin structure is reorganized, DNA becomes supercondensed and genetic activity is completely shut down37,38. For a long time, biological functions of this remodeling have been considered limited to the creation of a compact hydrodynamically efficient nuclear shape, with inert DNA fairly well protected from the environment. Therefore, the spermatozoon nucleus has been perceived as a ‘sac’ of genes that are to be transferred to an egg. Contrary to this point of view, specific and non-random chromosome architecture has recently been demonstrated for human sperm cells. In these studies, selected DNA sequences and chromosomal proteins were localized by FISH and immunocytochemistry followed by epifluorescent or laser scanning confocal microscopy. Several elements of GA in human sperm have been established: • Similar to somatic cells, individual chromosomes occupy distinct territories39–41 (Figure 5.1a). • Each chromosome has a preferred intranuclear localization (position), and the relative positioning of chromosomes is non-random42–46. • Centromeres (CEN) belonging to non-homologous chromosomes are collected into a compact chromocenter buried within a nuclear volume41,47,48 (Figure 5.1b). • Telomeres (TEL) are localized at the nuclear periphery where they interact in the form of dimers and tetramers44,49,50 (Figure 5.1c). • Telomere dimers correspond to the contacts between two ends of one chromosome rather than random association between chromosomal ends, and therefore chromosomes in sperm are looped51,52 (Figure 5.1d and e). Based on the acquired data, a general model for GA in human sperm has been proposed (Figure 5.1f ).

GENOME ARCHITECTURE IN HUMAN SPERM CELLS

(f) pk

qk pl ql

Figure 5.1 Chromosome organization in human sperm. (a) Chromosome territory: chromosome 6 (CHR6) (green) was localized using a painting probe. Total DNA counterstained with propidium iodide (PI) (red). (b) Centromeres (green) were visualized using immunofluorescence with antibodies against CENP-A (centromere protein A). Total DNA counterstained with PI (red). (c) Fluorescence in situ hybridization (FISH) using TTAGGG probe (yellow/green) shows that the majority of telomeres are joined as dimers and tetramers. Total DNA counterstained with PI (red). (d) Subtelomeric sequences located at the p and q arms of chromosome 3 (subTEL3q, pink; subTEL3p, emerald) are spatially close. Total DNA counterstained with diamidino-2-phenylindole (DAPI) (blue). (e) FISH using arm-specific probes microdissected from CHR1 (1q, green; 1p, red) indicates looping of this chromosome. Total DNA counterstained with DAPI (blue). (f) Schematic model of sperm nuclear architecture. Selected chromosome territories (pink and ocher), telomeres (TEL) (green circles) and centromeres (CEN) (red circles) are shown within a section through the nucleus. Nonhomologous CEN are clustered into a chromocenter, while TEL interact at the nuclear periphery. Modified from Ward and Zalensky 1996 (reference 38). See also Color plate 1 on page xxv

SPERM NUCLEAR STATUS AND MALE INFERTILITY Annually in the USA, more than 2 million conceptions are lost before the 20th week of gestation, and approximately half of these carry chromosomal defects such as numerical abnormalities, breaks/rearrangements and mutations1,53. Biochemical and FISH-based diagnostic procedures for detection of these chromosomal defects in germ-line cells and early embryos are either currently set up or being developed54–58.

Defective fertilization and/or early development may also be a consequence of abnormal DNA packaging in gamete nuclei. While structural organization of DNA in oocytes is poorly studied, it is generally accepted that a significant fraction of infertile males produce sperm with malformations in spermatozoa nuclei or chromatin defects. Among these are deficiencies in basic chromosomal proteins59,60, or broadly instituted chromatin condensation defects61–64. The latter defects have been determined using cytochemical and electron microscopy methods, while

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the molecular basis of flawed nuclear organization has remained unidentified. Male-factor infertility is a heterogeneous disorder, and the abnormalities in sperm chromatin/nuclear organization are most probably complex and diverse. In the following sections, we provide a few examples of nuclear aberrations that are connected with sperm genome architecture. We use for illustration sperm samples obtained from patients undergoing treatment in the fertility clinic. Comprehensive semen analysis indicated normal sperm count and motility but significantly abnormal sperm morphology (e.g. presence of round or torpedoid cells). Physical examination of the patients failed to reveal any abnormalities, including varicocele.

COMPACTNESS OF CHROMOSOME TERRITORY In 95% of sperm cells of fertile donors, FISH signals obtained using whole-chromosome painting probes (Figure 5.2a) or a combination of p and q arm-specific painting probes (Figure 5.2b) were confined to relatively small areas, and had sharp chromosome territory (CT) contours. Thus, FISH detects tightly packed, compact CTs formed by closely located p and q arms. The CT in normal sperm is approximately four times more condensed than the metaphase chromosome, and therefore is much more condensed than the interphase CT. In sperm of some patients with idiopathic infertility (three of the ten studied), abnormal hybridization patterns were observed (Figure 5.2c). In more detail, 42% of cells in sample P44 and 36% in P09 had large and diffuse signals; 27% of cells in sample P12 had multiple signals. The hybridization picture indicates that sperm in samples P44 and P09 may have had lesions in the formation of chromosome higher-order structures. Sperm of patient P12 may have had aneuploidy of chromosome 1 and/or large-scale rearrangement in its DNA (Figure 5.2 right hand panels.

CHROMOSOME POSITIONING Determination of the intranuclear chromosome position in human sperm is possible because these cells have a non-symmetrical elongated shape, and the site of tail attachment may easily be used as a spatial reference point46. Nevertheless, only a few studies in this direction have been performed so far. FISH using painting probes indicated that chromosome X43,44,46 and chromosome 646 were preferentially located in the anterior part of sperm nuclei, chromosome 18, near the sperm tail43, while chromosome 13 seemed to be randomly positioned44. In recent work, we found that in 90% of cells, chromosome 1 was located in the apical half of the nucleus, and 80% of chromosome 2 and 85% of chromosome 5 were preferentially located in the basal half 52. Using another approach, FISH with chromosome-specific centromere probes, preferential intranuclear positioning was shown for chromosomes 2, 6, 7, 16, 17, X and Y46. In the examples provided in Figure 5.2d–f, we traced the positioning of chromosomes by localization of FISH signals resulting from hybridization with DNA chromosomepainting probes. For each nucleus the position of chromosomes was assigned to a particular nuclear sector, I–IV, as illustrated in Figure 5.2d. About 100 nuclei from each sperm sample were analyzed, and the location of CTs is presented using diagrams of spatial distribution (Figure 5.2e and f ). Figure 5.2e demonstrates that in sperm from fertile donors, chromosome 6 had a tendency towards more anterior localization compared with chromosome 1, and both were rarely found in the posterior half of the nucleus. We also compared the position of chromosome 1 within nuclei of normal sperm and sperm of infertile patient P44. Figure 5.2f shows that the nuclear position of this chromosome in the infertile sperm sample was less confined. This might be a result of improper packaging, as noted above, and/or of an aberration in unknown mechanism(s) governing non-random chromosome localization.

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P44

(d)

P09

P12

(e)

(f) I II III IV

III

P12

I II III IV

Donor

P44

0 CHR1

CHR6

CHR1

Figure 5.2 Determination of chromosome intranuclear localization using fluorescence in situ hybridization (FISH) with painting probes. (a) Typical patterns of chromosome 1 (CHR1) painting probe hybridization (yellow) in normal sperm. (b) Typical patterns of CHR1 arm-specific probe hybridization (1p, green; 1q, red) in normal sperm. (c) Patterns of CHR1 hybridization in three samples of abnormal sperm. (d) Schematic example of CHR territory position in a sectioned sperm nucleus. (e) Charts showing distribution of CHR1 and CHR6 localization within sectors I–IV (percentage of hits to a sector from total FISH signals analyzed). (f) Comparison of nuclear positioning of CHR1 in normal and abnormal sperm cells. See also Color plate 3 on page xxvi

TELOMERE LOCALIZATION Localization of telomere repeat sequences (TTAGGG)N in human sperm reveals that most, if not all, telomeres are joined in dimers and tetramers (Figures 5.1c and 5.3a)49. As a result, on a frequency distribution plot (Figure 5.3c), the majority of nuclei fall into two peaks: the first corresponds to 12 hybridization loci (TEL tetramers), and the second to 24 loci (TEL dimers). In the absence of telomere–telomere interactions in human sperm, 46 hybridization signals (2 telomeres × 23 chromosomes) should be observed.

We compared the localization of telomeres in sperm between donors and patients (total 20 patients) (Figure 5.3). Three sperm samples obtained from infertile males showed strikingly different telomere localizations. In the majority of cells, hybridization was in numerous small dots dispersed over the nucleus (Figure 5.3b). As a result, no telomere grouping was seen in the frequency distribution plot (Figure 5.3c). Such localization reflects the absence of telomere–telomere interactions, which are characteristic of normal human sperm. The molecular basis of this phenotype is unknown. Atypical sperm telomere-binding

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Donor Number of cells

50 Patient

Tetramers 40 30 20 10 0 1

Number of TEL signals

Figure 5.3 Comparison of nuclear localization of telomeres in normal and abnormal cells. (a) Telomeres are joined as dimers and tetramers in normal sperm. (b) Telomere hybridization appears as numerous small dots dispersed over the nucleus in abnormal sperm cells. (c) Frequency of telomere (TEL) hybridization signal distribution in sperm cells determined by fluorescence in situ hybridization (FISH). In the majority of normal sperm cells, the number of TEL hybridization signals peaks at 24 (TEL dimers) and 12 (TEL tetramers)

proteins65 or aberrant telomere DNA may be involved.

GENOME ARCHITECTURE AND UNPACKING OF SPERM GENOME DURING FERTILIZATION Data above were obtained using small, random selections of patients with idiopathic male infertility. Nevertheless, they clearly illustrate the existence of three categories of deviations from the standard genome architecture characteristic of sperm cells: (1) atypical packing of chromosome

territories, (2) unstable or aberrant nuclear positioning of chromosomes and (3) disturbed telomere interactions. What are the possible effects of such faults on successful fertilization and early development? Normal mammalian embryogenesis requires the participation of both a maternal and a paternal genome66. Genetically inert chromatin of the spermatozoa is remodeled into the decondensed and transcriptionally competent chromatin of the male pronucleus upon entry into the ooplasm; this remodeling is controlled by an oocyte activity that appears during meiotic maturation67. Reorganization of the sperm genome after fertilization is

GENOME ARCHITECTURE IN HUMAN SPERM CELLS

a complex process that involves chromosome withdrawal from the nucleus, their decoration with histones (decondensation), formation of the male pronucleus and its movement towards the female pronucleus68,69. Exchange of the basic chromosomal proteins involves chaperones of the nucleoplasmin family70. Overall, the molecular characterization of participants responsible for pronucleus development is at an early stage. While the activity of sperm chromosome remodeling is of maternal origin, the structural organization and biochemical composition of sperm nuclei are equally important. Improperly packed and spatially unorganized sperm chromosomes will have a high probability of being inadequately processed by egg cytoplasm. Transcription is influenced by the underlying chromatin structure, including the organization of chromosome territories71, and therefore activation of the male genome will depend on the specific sperm GA. Recent data show that in mammals, transcription begins earlier than in zygotes from other classes of organisms, starting several hours after fertilization in male pronuclei and continuing in embryonic nuclei72–74. Hence, it is highly probable that abnormal genome architecture in sperm (or undeveloped GA in immature gametes) may cause irregularities in early development. In addition, since paternal and maternal genomes are spatially separated up to the 4-cell embryo stage, chromatin remodeling after fertilization occurs in separate nuclear compartments and consequently may be regulated in a parent-specific manner75. The data overviewed above indicate that each chromosome in human sperm has a preferential intranuclear position. Since, during normal fertilization, sperm penetration begins with the acrosome, there is a sequential order of exposure of sperm chromosomes to the egg cytoplasm during sperm entry. Therefore, a predetermined order of chromosome activation induced by chromatin remodeling by egg factors may exist. We propose that deviation from regular sperm chromosome localization may be deleterious for proper fertilization and development.

It is noteworthy that, in all mammals, sex chromosomes are located in the region nearest to the acrosome, and are presumably the first chromosomes to enter the egg on fertilization76. Such a position has been preserved between monotreme and marsupial mammals, which diverged from eutherian mammals 170 and 130 million years ago, respectively77. This strongly supports the hypothesis of a functional significance of the intranuclear localization of sperm chromosomes. While modern clinical assisted reproductive technologies broadly use intracytoplasmic injection using sperm and occasionally even immature gametes, the molecular/cellular mechanisms of fertilization after ISCI have been poorly studied78. Some publications have reported an increased rate of de novo chromosomal anomalies in human babies following ICSI79. Importantly, in several species, delayed decondensation of the apical region of the sperm nucleus and postponed replication of the male genome after ICSI were observed43,80–82. Immunofluorescent analysis showed that the perinuclear theca of sperm persisted around the condensed apical portion following ICSI, whereas it was removed completely from the sperm nucleus after in vitro insemination80. The presence of sex chromosomes in the condensed apical region of the sperm nucleus might lead to sex chromosomal anomalies, introducing the delay of S-phase entry. In particular, this atypical decondensation may unbalance normal remodeling of sex chromosomes (e.g. introducing delay of their entry to the S-phase, or gene activation), which are located in this region of the nucleus. Therefore, an ICSI procedure itself may lead to birth defects because of improper processing of a well-defined GA characteristic of normal sperm. Examples provided above (Figure 5.3) show disturbed localization of telomeres and telomere–telomere interactions in sperm from patients with idiopathic infertility. In human sperm, the telomere chromosomal domain is characterized by elongated DNA (in comparison with somatic cells) and sperm-specific telomeric

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proteins49,83,84. The elongation of telomere DNA during spermatogenesis is characteristic of all mammals85, and is provided by telomerase, a specific reverse transcriptase, which is highly active in germline cells86,87. In the mouse, the fertilization of oocytes with sperm obtained from telomerase knock-out males resulted in aberrant cleavage and development88. These results suggest that the state of telomere DNA in sperm contributes to defective fertilization and cleavage. Currently there are no equivalent data obtained in humans. Nevertheless, we propose a general hypothesis that telomeres in human spermatozoa have unique molecular and structural features critical for function during fertilization and early embryonic development. Experiments to characterize telomeres in infertile patients are under way in our laboratory.

ACKNOWLEDGMENTS This work was supported by a National Institutes of Health (NIH) grant HD-042748 to one of the authors (AZ).

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8. Schmidt E, et al. Detection of structural and numerical chromosomal abnormalities by ACM-FISH analysis in sperm of oligozoospermic infertility patients. Hum Reprod 2004; 19: 1395 9. Petit FM, et al. Could sperm aneuploidy rate determination be used as a predictive test before intracytoplasmic sperm injection? J Androl 2005; 26: 235 10. Foresta C, et al. Genetic abnormalities among severely oligospermic men who are candidates for intracytoplasmic sperm injection. J Clin Endocrinol Metab 2005; 90: 152 11. Pang MG, et al. The high incidence of meiotic errors increases with decreased sperm count in severe male factor infertilities. Mol Vis 2005; 11: 152 12. Rabl C. Ueber Zelltheilung [On cell division]. Morphol Jahrbuch 1895; 10: 214 13. Boveri T. Die blastomerenkerne von Ascaris megalocephala und die Theorie der Chromosomenindividualitat [The blastomere nucleus of Ascaris megalocephala and the theory of chromosome individuality on cell division]. Arch Zellforsch 1909; 3: 181 14. Marshall WF. Order and disorder in the nucleus. Curr Biol 2002; 12: 185 15. Taddei A, et al. The function of nuclear architecture: a genetic approach. Annu Rev Genet 2004; 38: 305 16. Cremer T, et al. Higher order chromatin architecture in the cell nucleus: on the way from structure to function. Biol Cell 2004; 96: 555 17. Cremer T, Cremer C. Chromosome territories, nuclear architecture and gene regulation in mammalian cells. Nat Rev Genet 2001; 2: 292 18. Zalensky AO. Genome architecture. In Verma RS, ed. Advances in Genome Biology. Greenwich, London: JAI Press, 1998: 179 19. Cremer M, et al. Inheritance of gene density-related higher order chromatin arrangements in normal and tumor cell nuclei. J Cell Biol 2003; 162: 809 20. Cremer M, et al. Non-random radial higher-order chromatin arrangements in nuclei of diploid human cells. Chromosome Res 2001; 9: 541 21. Parada L, Misteli T. Chromosome positioning in the interphase nucleus. Trends Cell Biol 2002; 12: 425 22. Kozubek S, et al. 3D structure of the human genome: order in randomness. Chromosoma 2002; 111: 321 23. Boyle S, et al. The spatial organization of human chromosomes within the nuclei of normal and emerin-mutant cells. Hum Mol Genet 2001; 10: 211 24. Croft JA, et al. Differences in the localization and morphology of chromosomes in the human nucleus. J Cell Biol 1999; 145: 1119

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25. Parada LA, et al. Conservation of relative chromosome positioning in normal and cancer cells. Curr Biol 2002; 12: 1692 26. Leitch A, et al. The spatial localization of homologous chromosomes in human fibroblasts at mitosis. Hum Genet 1994; 93: 275 27. Nagele R, et al. Precise spatial positioning of chromosomes during prometaphase: evidence for chromosome order. Science 1995; 270: 1831 28. Allison DC, Nestor AL. Evidence for a relatively random array of human chromosomes on the mitotic ring. J Cell Biol 1999; 145: 1 29. Manuelidis L. Indications of centromere movement during interphase and differentiation. Ann NY Acad Sci 1985; 450: 205 30. Neves H, et al. The nuclear topography of ABL, BCR, PML, and RAR alpha genes: evidence for gene proximity in specific phases of the cell cycle and stages of hematopoietic differentiation. Blood 1999; 93: 1197 31. Bridger JM, et al. Re-modelling of nuclear architecture in quiescent and senescent human fibroblasts. Curr Biol 2000; 10: 149 32. Sadoni N, et al. Nuclear organization of mammalian genomes. Polar chromosome territories build up functionally distinct higher order compartments. J Cell Biol 1999; 146: 1211 33. Stadler S, et al. The architecture of chicken chromosome territories changes during differentiation. BMC Cell Biol 2004; 5: 44 34. Parada LA, et al. Tissue-specific spatial organization of genomes. Genome Biol 2004; 5: R44 35. Meistrich M. Histone and basic nuclear protein transitions in mammalian spermatogenesis. In Hnilica LS, Stein GS, Stein JL, eds. Histones and Other Basic Nuclear Proteins. Boca Raton, FL: CRC Press, 1989: 165 36. Churikov D, et al. Male germline-specific histones in mouse and man. Cytogenet Genome Res 2004; 105: 203 37. Balhorn R. A model for the structure of chromatin in mammalian sperm. J Cell Biol 1982; 93: 298 38. Ward WS, Zalensky AO. The unique, complex organization of the transcriptionally silent sperm chromatin. Crit Rev Eukaryot Gene Expr 1996; 6: 139 39. Brandriff BF, Gordon LA. Spatial distribution of sperm-derived chromatin in zygotes determined by fluorescence in situ hybridization. Mutat Res 1992; 296: 33 40. Haaf T, Ward DC. Higher order nuclear structure in mammalian sperm revealed by in situ hybridization

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71. Gilbert N, et al. Chromatin organization in the mammalian nucleus. Int Rev Cytol 2005; 242: 283 72. Bouniol-Baly C, et al. Dynamic organization of DNA replication in one-cell mouse embryos: relationship to transcriptional activation. Exp Cell Res 1997; 236: 201 73. Aoki F, et al. Regulation of transcriptional activity during the first and second cell cycles in the preimplantation mouse embryo. Dev Biol 1997; 181: 296 74. Capco DG. Molecular and biochemical regulation of early mammalian development. Int Rev Cytol 2001; 207: 195 75. Mayer W, et al. Spatial separation of parental genomes in preimplantation mouse embryos. J Cell Biol 2000; 148: 629 76. Greaves IK, et al. Conservation of chromosome arrangement and position of the X in mammalian sperm suggests functional significance. Chromosome Res 2003; 11: 503 77. Kirsch JAW, et al. DNA-hybridisation studies of marsupials and their implications for metatherian classification. Aust J Zool 1997; 45: 211 78. Schatten G, et al. Cell and molecular biological challenges of ICSI: ART before science? J Law Med Ethics 1998; 26: 29 79. Bonduelle M, et al. Seven years of intracytoplasmic sperm injection and follow-up of 1987 subsequent children. Hum Reprod 1999; 14: 243 80. Sutovsky P, et al. Intracytoplasmic sperm injection for rhesus monkey fertilization results in unusual chromatin, cytoskeletal, and membrane events, but eventually leads to pronuclear development and sperm aster assembly. Hum Reprod 1996; 11: 1703 81. Hewitson L, et al. Unique checkpoints during the first cell cycle of fertilization after intracytoplasmic sperm injection in rhesus monkeys. Nat Med 1999; 5: 431 82. Terada Y, et al. Atypical decondensation of the sperm nucleus, delayed replication of the male genome, and sex chromosome positioning following intracytoplasmic human sperm injection (ICSI) into golden hamster eggs: does ICSI itself introduce chromosomal anomalies? Fertil Steril 2000; 74: 454 83. Zalenskaya IA, Bradbury EM, Zalensky AO. Chromatin structure of telomere domain in human sperm. Biochem Biophys Res Commun 2000; 279: 213 84. Bekaert S, et al. Telomere biology in mammalian germ cells and during development. Dev Biol 2004; 274: 15 85. Kozik A, et al. Identification and characterization of a bovine sperm protein that binds specifically to

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single-stranded telomeric deoxyribonucleic acid. Biol Reprod 2000; 62: 340 86. Kim NW, et al. Specific association of human telomerase activity with immortal cells and cancer. Science 1994; 266: 2011 87. Achi MV, et al. Telomere length in male germ cells is inversely correlated with telomerase activity. Biol Reprod 2000; 63: 591

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6 Sperm pathology: pathogenic mechanisms and fertility potential in assisted reproduction Hector E Chemes, Vanesa Y Rawe

INTRODUCTION

analyzed in detail with the light microscope, which allows detailed observation of the external profile of the spermatozoon but does not give information on its internal structure. The combination of high-resolution light and electron microscopy, immunocytochemistry and molecular studies has provided new insights into the structure of normal and abnormal spermatozoa, and defined the subcellular basis of sperm aberrations. Furthermore, correlation of these data with relevant clinical and fertility information has shed new light on this field. This approach goes beyond descriptive morphology of the appearance of spermatozoa. Several important questions remain. What is it that impairs sperm function in morphologically abnormal sperm? What is wrong with a wrong sperm shape? What hides behind the headshape change in amorphous or tapering spermatozoa? Is it just the abnormal shape, or is there something wrong with specific sperm components? Sperm pathology is the discipline of characterizing structural and functional deficiencies in abnormal spermatozoa. This is significant because it helps to explain the mechanisms of sperm inefficiency, identifies genetic phenotypes, suggests strategies to improve fertilization and opens a door to molecular genetic studies that will probably lead to the design of therapeutic tools of the future. Two main examples of sperm alterations can be distinguished. The most frequent is characterized

Teratozoospermia, asthenozoospermia and necrozoospermia are frequently responsible for infertility in men, and have a negative influence on the fertility prognosis when assisted reproductive technologies (ART), including in vitro fertilization (IVF), are attempted. The introduction of intracytoplasmic sperm injection (ICSI) allowed examination of the motility and morphology of the very same spermatozoon that was to be microinjected. It then became clear that abnormal and immotile spermatozoa could successfully fertilize oocytes, and the issue of the convenience of using them in ART procedures was raised. Some andrologists have stressed the importance of using different tools to characterize sperm pathologies and establish a diagnosis; still others have been more inclined to use spermatozoa in ICSI without paying much attention to the nature of the pathologies involved. Sperm morphology, the subject of numerous studies, has been subjectively assessed or characterized by manual or computer-assisted objective methods1–3. Strict criteria for sperm classification have been introduced, and a correlation between sperm morphology and prognosis in ART has received general acceptance4,5. In all of these methods, the morphometric parameters of the sperm head, middle piece and flagellum have been 85

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by a heterogeneous array of sperm anomalies that do not follow a uniform pattern, and demonstrate different combinations in each individual and among different patients. These are non-specific anomalies that are potentially reversible and usually secondary to diverse conditions affecting the reproductive system. The second type is characterized by a well-defined, uniform pattern of anomalies that affect the vast majority of spermatozoa, and present a similar configuration in different patients suffering from the same condition. These alterations are stable in time, do not respond to therapeutic interventions, may display family clustering and have a recognized or presumed genetic origin. Because of these characteristics, these alterations are known as systematic sperm defects.

PATHOLOGICAL SPERM PHENOTYPES ASSOCIATED WITH MOTILITY DISORDERS To understand fully the physiopathology of asthenozoospermia, it is first necessary to summarize briefly the ultrastructure of the sperm tail. The human sperm flagellum is a long structure, approximately 50 µm in length and 0.4–0.5 µm in diameter. It is composed of a central element, the axoneme, which is a cylinder comprising a circumferential array of nine peripheral microtubular doublets surrounding a central pair of microtubules, the so-called 9 + 2 configuration (Figure 6.1a). Each peripheral doublet is composed of two apposed subunits, microtubules A and B, consisting of protofilaments of tubulin heterodimers. Extending from subunit A, two arms project toward the B subunit of the next doublet. These arms are composed of dynein, a structural protein with adenosine triphosphatase (ATPase) activity that utilizes ATP as an energy source to generate axonemal movement6,7. The axoneme is surrounded by the outer dense fibers (ODFs) and the fibrous sheath. The ODFs are nine slender cylindrical structures associated with the corresponding peripheral doublet. The fibrous sheath is a sort of

flagellar exoskeleton, present only at the main piece, and organized into two longitudinal columns that run along the length of the principal piece and insert into microtubular pairs 3 and 8. These columns are joined regularly by transverse semicircular ribs. Asthenozoospermia is a frequent cause of male infertility. Both non-specific and systematic sperm phenotypes can be responsible for alterations in sperm motility. Non-specific flagellar anomalies (NSFAs) are the underlying cause in most men with severe asthenozoospermia8–12. In NSFAs, the normal 9 + 2 organization of the sperm tail is replaced by a combination of modifications in the number, topography and organization of microtubular pairs and periaxonemal structures of the flagellum (Figure 6.1b). Affected flagella appear normal under light microscopy, and are only identified by ultrastructural examination, because their outer diameter and profile are not modified. NSFAs are either idiopathic or secondary to various andrological conditions such as varicocele, infections, immune factor, orchitis and other endogenous or environmental factors. Since these same kinds of anomalies are found in lower numbers in most fertile men, their incidence should be determined in each particular asthenozoospermic patient by means of careful quantification of no less than 100 transverse sections of the sperm tail. We have set the upper normal limit of NSFAs to 40% of the sperm population; values in the 40–60% range are borderline; and above the 60% threshold they are certainly pathological. There is no genetic background in NSFAs which are potentially responsive to etiological or empirical therapeutic interventions. Their prevalence fluctuates during clinical evolution and among different asthenozoospermic men12–16. Genetically determined sperm phenotypes causing asthenozoospermia have been the subject of numerous studies since the mid 1970s, when the lack of dynein arms was identified as the main underlying cause of ciliar and flagellar paralysis in men suffering from extreme asthenozoospermia

SPERM PATHOLOGY: PROGNOSIS IN ASSISTED REPRODUCTION

Figure 6.1 Abnormalities of the tail and midpiece. (a) Cross-section of a normal sperm flagellum at the principal piece. The nine peripheral doublets of the axoneme, central pair, dynein arms (arrow) and radial spokes are clearly seen. The fibrous sheath is composed of two lateral columns inserted in doublets 3 and 8 (asterisks) and semicircumferential ribs (arrowheads). (b) Sperm tail with non-specific flagellar anomalies. The central pair is displaced (asterisk) and there is microtubular translocation to the center and the periphery of the axoneme or outside the fibrous sheath (arrows). (c, d) Spermatozoa from two patients with primary ciliary dyskinesia. There is a lack of dynein arms (arrow, c) or absence of the central pair (d). Bars (a–d) = 0.1 µm. (e–g) Light and transmission electron microscopy (TEM) of spermatozoa with dysplasia of the fibrous sheath (DFS). (e) Very short, thick and irregular tails are seen (phase-contrast microscopy). (f) Longitudinal section of a DFS sperm. Note absence of the mitochondrial sheath (asterisk) and redundant elements of the fibrous sheath. (g) Cross-section of flagellum with disorganized and hyperplastic fibrous sheath. The axoneme is almost completely obliterated with few remaining microtubular doublets and missing dynein arms (arrow). Bars = 5 µm (e), 1 µm (f), 0.1 µm (g). (h–k) Alterations of the mitochondrial sheath (MS). (h) Under epifluorescence, this spermatozoon displays intense and uniform labeling of the MS that covers a length > 15 µm (normal length 3–5 µm). (i) Abnormally long and distorted MS observed in TEM. (j) Absence of MS (very small labeling in the midpiece corresponding to isolated mitochondrion, arrow). (k) Under TEM, the midpiece is not formed and mitochondria are either absent or abnormal in location and/or arrangement. Bars = 5 µm (h, j), 1 µm (i, k). Panels (f) and (g) were originally published in reference 12. Copyrights European Society of Human Reproduction and Embryology. Reproduced by permission of Oxford University Press/Human Reproduction

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and chronic respiratory disease in the so-called immotile cilia syndrome (ICS)17–19. ICS was more recently renamed as primary ciliary dyskinesia (PCD), because various degrees of reduced or qualitatively abnormal motility were reported in some of these patients20–22. PCD patients are infertile owing to sperm immotility or severe asthenozoospermia, suffer from rhinosinusitis and chronic pneumopathy caused by infections secondary to faulty mucociliary clearance, and have alterations in the visceral situs, with dextrocardia in 50% of patients23. Familial incidence of PCD, most possibly due to autosomal recessive mutation(s), has been reported. There is extensive locus heterogeneity, with a number of mutations in dynein genes found in families with members carrying the PCD phenotype24–29. Spermatozoa in PCD patients have immotile or dyskinetic flagella of normal appearance under the light microscope. The underlying alteration consists of the lack of one or both dynein arms, absence of the central pair, microtubular transposition or a number of less frequent abnormal configurations of the sperm axoneme (Figures 6.1c and d)17,18,20,24,30–35. The possibility also exists of isolated immotility in either cilia or flagella. Another systematic sperm phenotype responsible for severe asthenozoospermia/sperm immotility is dysplasia of the fibrous sheath (DFS). Patients are young males with primary sterility and immotile spermatozoa. Sperm flagella are typically short, thick and of very irregular profile (Figure 6.1e). This appearance prompted the denomination ‘stump tails’ or ‘short tails’, a descriptive name that does not give any clues as to the nature and subcellular basis of this pathology. We have proposed DFS, which recognizes the main alterations in the sperm fibrous sheath and identifies its testicular origin as a consequence of a dysplastic development of the tail during spermiogenesis12,16,36–38. Other authors39,40 have previously indicated that this anomaly involves various components of the tail cytoskeleton, the fibrous sheath being the most visibly affected. DFS sperm should not be confused with other alterations

secondary to necrozoospermia, or sperm aging in men with partial obstruction of the seminal pathway, that lead to flagellar disintegration and thickening. Familial and geographical clustering of DFS has been reported12,39–41. A striking contrast between the high incidence of DFS and low incidence of PCD has been noted in a population of multiethnic origin16, which may indicate the interaction between genetic and environmental influences in the generation of this phenotype. The subcellular basis of DFS is a serious disarray of the sperm-tail cytoskeletal components. The fibrous sheath appears hyperplastic and completely disorganized, and the axoneme may be disrupted. There is also frequent absence of the central pair, missing dynein arms and lack or minimal development of the mitochondrial sheath of the midpiece (Figure 6.1f and g). These abnormalities are very stable during evolution, do not respond to any therapeutic measures, have familial incidence and may be associated with a lack of dynein in the respiratory cilia (see below). These alterations point to a genetic origin of DFS, possibly an autosomic recessive trait12,42–45. About 20% of patients also suffer from chronic respiratory disease due to a lack of dynein in the respiratory cilia. This subgroup of DFS patients constitutes a variety of primary ciliary dyskinesia in which a lack of dynein in the respiratory cilia is associated with the DFS phenotype in spermatozoa35,41. In recent years, extensive work has been carried out on the protein composition of the fibrous sheath. A kinase anchoring protein 3 (AKAP3) and AKAP4 have been recognized as the most abundant structural proteins of the fibrous sheath. They bind to one another and provide the structural framework for docking of protein kinase A to the fibrous sheath46. To analyze the possible role of these proteins in generation of the DFS phenotype, sequence analysis of the AKAP3 and AKAP4 binding sites in DFS patients was carried out, but did not reveal mutations47. However, targeted disruption of the AKAP4 gene in mice resulted in sperm immotility and abnormally short flagella with localized aggregations of fibrous sheath

SPERM PATHOLOGY: PROGNOSIS IN ASSISTED REPRODUCTION

material, somewhat reminiscent of the DFS phenotype48 (Eddy, personal communication). Very recently, Baccetti et al.49 have reported deletion of the AKAP4/AKAP3 binding regions and absence of the AKAP4 protein in spermatozoa of one of five patients with DFS. This report suggests that lack of AKAP4 could be pathogenically responsible for the DFS phenotype. It is possible that DFS is a multigenic disease caused by alterations in several different gene products. Intensive research into this field is currently being carried out. Other more rare forms of axonemal pathologies of genetic origin include deficient respiratory cilia and sperm axonemes in patients with retinitis pigmentosa14,50 or albinism (unpublished personal observation). Mitochondrial anomalies in the sperm midpiece such as an abnormally long extension or absence of the mitochondrial sheath are very infrequent sperm anomalies that are also associated with asthenozoospermia (Figures 6.1h–k)51. Recent investigations have identified various mutations/deletions in mitochondrial genes of immotile spermatozoa whose products are involved in oxidative phosphorylation and generation of ATP necessary for sperm motility52,53. No structural correlates of these anomalies have been described so far.

ABNORMAL HEAD–NECK ATTACHMENT AND ACEPHALIC SPERMATOZOA The region of head–neck attachment or the connecting piece derives from interaction of the centrioles with the spermatid nucleus (Figure 6.2c). Early in spermiogenesis, the sperm flagellum grows from the centriolar complex, while this approaches the nucleus and attaches to its caudal pole, ensuring linear alignment of the tail with the longitudinal axis of the head. Spermatozoa without heads (‘acephalic’, ‘decapitated’, ‘pin heads’; Figure 6.2b) or with an abnormal head–midpiece relationship (‘abaxial implantation’; Figure 6.2a) can be detected in very small numbers in the semen of fertile men, and

can rise up to 10–20% in subfertile patients36,54. Its significance for fertility is not clear in these situations. There are infertile patients in whom 80–100% of the sperm population is composed of acephalic forms and loose heads, or spermatozoa with heads and tails not aligned along the same axis. Each of these two forms can predominate or combine in different proportions. This sperm defect is of rare occurrence albeit underdiagnosed, since these patients are usually considered to suffer from ‘severe teratozoospermia’, without the specificity of this sperm defect being recognized. Several authors55–57 reported individual patients with headless flagella in the semen, and more recently, other authors36,58–60 reported 15 more cases, including familial incidence. The term ‘pin heads’ has been used in reference to this peculiar appearance, but this denomination adds confusion, since there is no nuclear material in these minute ‘heads’. Acephalic forms appear as headless flagella ending cranially in a small cytoplasmic droplet that, when bigger, simulates a head, but has no DNA content (Figure 6.2b and e)36. When a head is present, it attaches either to the tip or to the sides of the midpiece, without linear alignment with the sperm axis (Figure 6.2a and d). This misalignment ranges from complete lack of connection to lateral positioning of the nucleus at a 90–180° angle. All forms of this defect result from failure of the sperm centriole to attach normally to the caudal pole of the maturing spermatid nucleus, reported on the few occasions on which testicular biopsies from these patients have been studied (Figure 6.2f )61,62. These variants express different degrees of abnormality of the head–neck junction, with acephalic forms representing the most extreme situation, and hence the more inclusive denomination of alterations of the head–neck attachment59,62,63. When the relationship between the head and midpiece is looser, increased fragility of this junction determines the generation of acephalic forms and loose heads59,64. The latter are frequently phagocytosed within the testis, and their frequency in semen is lower than that of headless flagella.

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Figure 6.2 Abnormalities of the connecting piece (head–tail junction). In (a) the head and the tail are not aligned along the same axis (abaxial implantation of the tail). (b) Acephalic spermatozoon with minute thickening (arrow). (c) Normal configuration of the connecting piece. The tail is lodged in the concave implantation fossa (arrow). Note the triplets of the proximal centriole (asterisk) and beginning of the axoneme. (d) The head and midpiece are not properly attached and a vesicular structure (V) separates them. (e) Acephalic spermatozoon. The plasma membrane (arrow) covers the connecting piece (asterisk). The midpiece is well formed. (f) Elongating spermatid in testicular biopsy. Note lack of attachment of the tail anlagen to the caudal pole of the nucleus (arrows). Bars = 5 µm (a, b), 0.5 µm (c–f). Panels (a) and (b) were originally published in reference 62 and panels (c–f) in reference 59. Copyrights European Society of Human Reproduction and Embryology. Reproduced by permission of Oxford University Press/Human Reproduction

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The uniform pathological phenotype, its origin as a consequence of a systematic alteration during spermiogenesis, the fact that seminal characteristics remain constant along clinical evolution even when a pharmacological germ cell depletion–repopulation has been induced, and the familial incidence indicate that this condition is very likely of genetic origin59,60. The need for normal migration of the spermatid centriole to generate a normal head–midpiece attachment, and the abnormalities that have been observed in sperm aster formation, syngamy and embryo cleavage when these spermatozoa have been microinjected in bovine and human oocytes, point to a sperm centriolar dysfunction, the nature of which remains to be elucidated. Proteins such as centrin, pericentrin, γ-tubulin and MPM-2 have been localized to the sperm connecting piece and zygote centrosome, but no studies are available that show their (possible) significance in the pathogenesis of this syndrome63,65. Sequencing across the exons of the gene for speriolin (another protein localized to the sperm neck region) has failed to demonstrate any abnormality in two patients with the syndrome (Eddy, personal communication). The release of the sperm centriole after fertilization probably involves the action of sperm proteasomes recently localized to the neck region of human spermatozoa66,67. Experimental neutralization of proteasomes in the zygote has also resulted in defective sperm-aster and pronuclear formation67. Defective enzymatic activity of sperm proteasomes in patients with defects of the head–midpiece attachment has recently been reported68. We are currently conducting research in this exciting area of sperm pathology.

PATHOLOGY OF THE SPERM HEAD: ACROSOME AND CHROMATIN ANOMALIES The sperm acrosome is an organelle derived from the Golgi complex of spermatids. It consists of a

flattened sac covering the anterior two-thirds of the sperm head and is formed by two membranes (the internal and external acrosomal membranes), delimiting a space with a dense content rich in hydrolytic enzymes. The lack or insufficient development of the acrosome are specific sperm defects causing infertility, characterizing two well-defined syndromes: acrosomeless spermatozoa and acrosomal hypoplasia. Spermatozoa lacking acrosomes usually display spherical heads, which has prompted the denominations ‘globozoospermia’ or ‘round-headed acrosomeless spermatozoa’. They can be found in small numbers (approximately 0.5%) in the semen of fertile individuals, and may increase up to 2–3% in cases of infertility69. The denomination globozoospermia applies when they predominate in the vast majority of spermatozoa (up to 100% of ejaculated spermatozoa). Affected spermatozoa have an absence of or detached acrosomes, or very small perinuclear densities that may be abortive attempts at acrosome formation (Figure 6.3a and c). The generation of spermatozoa with absence of an acrosome corresponds to more than one mechanism. Most reports indicate that the Golgi complex fails to join the nucleus and develops a detached acrosome with irregular secretory activity. This structure remains free in the cytoplasm of maturing spermatids, to be eliminated with the residual cytoplasm at spermiation. In this situation, acrosomes are formed but do not attach to the nucleus70–73. In some other patients there is a real lack of or serious deficiency in acrosome formation. In these cases, a rudimentary acrosome may be found on the anterior pole of the spermatozoon71. One characteristic finding is delayed maturation of the chromatin; it appears granular, with incomplete compaction frequently in the form of hypodense areas. These changes are due to failure of the histone–protamine transition and increased rates of DNA fragmentation. Lack of the acrosome is associated with absence of the perinuclear theca, a subacrosomal structure of the sperm nuclear–perinuclear skeletal

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Figure 6.3 Acrosome and chromatin anomalies. (a) Light microscopy of spermatozoa from a patient with globozoospermia. Heads are characteristically spherical. (b) Detailed visualization of sperm heads with pathological acrosomes. Immunolabeling using antiacrosin antibody shows fluorescence on the acrosome. Lack (left) or variable hypoplasia (two right spermatozoa) are clearly observed. (c) A round-headed spermatozoon lacks the acrosome (arrows). There is also a marked lacunar defect of the chromatin. (d) Acrosomal hypoplasia: small and detached acrosome (asterisks). (e) Severe lacunar defect of the chromatin in a grossly distorted amorphous head. Bars = 5 µm (a, b), 0.5 µm (c–e)

complex involved in modeling the shape of sperm heads, attachment of the acrosome to the nucleus and also oocyte activation after sperm penetration74–77. These abnormalities of the perinuclear theca are probably the molecular basis responsible for spherical sperm heads, detached acrosomes and insufficient oocyte activation in acrosomeless spermatozoa. Familial incidence has been reported in men suffering from globozoospermia, and a mono- or polygenic origin has been suggested but not proven43,72,78. Various animal models with similar characteristics have recently been described. Acrosomal hypoplasia is a poorly understood and frequently underdiagnosed sperm pathology

that, according to Zamboni79, is frequent in severe teratozoospermia. Acrosomes are very small, and often lack contact with the amorphous nucleus (Figure 6.3b and d). Chemes16 reported a series of 35 patients with acrosomal anomalies in whom lack of the acrosome or acrosomal hypoplasia was present as a predominant form or in combination. Sperm heads are mostly round, but may also be amorphous or oval. Acrosomal hypoplasia should be investigated in cases of severe teratozoospermia, and can be readily recognized under the electron microscope80, with the use of various antibodies that react against the acrosome or by lectin binding to intact spermatozoa. In the classification of spermatozoa by strict criteria, these abnormalities

SPERM PATHOLOGY: PROGNOSIS IN ASSISTED REPRODUCTION

are included among the severe amorphous varieties that have poor fertility prognosis5. Other forms of acrosomal defects have been reported in infertile males. Premature occurrence of and/or failure to undergo the acrosome reaction have been recognized81. More rare and not wellcharacterized defects of the acrosome include the ‘crater defect’82 and acrosomal inclusions80. In both cases, fertility is compromised by the inability of these spermatozoa to penetrate oocytes normally. The chromatin of maturing spermatids suffers a complex series of chemical and macromolecular changes that are reflected in the structure of the nucleus. Early round spermatids have euchromatic nuclei with dispersed chromatin. During maturation, the chromatin condenses progressively in the form of discrete granules that enlarge as they approach each other and condense to acquire finally a dense, homogeneous structure in which only small (0.1–0.2 µm) hypodense, clear areas can be discerned. This process of progressive maturation and compaction is due to the replacement of nuclear histones that associate with the DNA in a supercoiled structure, similar to that found in somatic cells. Histones are interchanged first by transition proteins and later by protamines that organize in a side-to-side configuration along the groove of the DNA helix, so that chromatin fibers can compact tightly to determine the typical condensed structure of mature spermatids and spermatozoa83–85. In this compacted state, individual chromatin granules cannot be discerned. When the process of chromatin maturation and compaction is altered, the heads of the spermatozoa display large lacunar defects (2–3 µm in diameter), where the compact arrangement of the chromatin is replaced by granulofibrillar or ‘empty’ areas that occupy as much as 20–50% of the nucleus (Figure 6.3c–e)79,86. They originate in the testis as a consequence of abnormal spermiogenesis, as confirmed by their presence in immature spermatids found in testicular biopsies and semen. Spermatozoa with chromatin abnormalities frequently demonstrate abnormal head shapes, have diminished fertility potential or are

associated with first-trimester abortions16. Singlestranded DNA, DNA breaks, abnormal histone–protamine transition or apoptotic changes have been reported, as well as insufficient chromatin condensation, immaturity and intranuclear lacunae that are their ultrastructural correlates. There is not much information about the genetic constitution of morphologically abnormal spermatozoa. A positive correlation between sperm aneuploidy and teratozoospermia has been reported, but in other studies no increased numerical chromosomal aberrations have been found in abnormal spermatozoa87–89. Recent fluorescence in situ hybridization (FISH) studies of infertile men with poor semen quality have shown increased aneuploidy in spermatozoa, despite a normal blood karyotype90,91, which suggests that the same factor(s) causing aneuploidy may also induce teratozoospermia. The question of the acquired versus the genetic etiology of chromatin anomalies has received attention, but is not solved to date. Men who suffer from infectious bowel disease and are treated with sulfasalazine may present with this type of abnormality in the spermatozoa. The question remains whether they are caused by the pathological process itself or the treatment instituted. The same alterations can also be found in men with varicocele, fever, seminal infections and even testicular tumors92–95. In these last cases they are found mixed with other types of non-specific sperm anomalies. Accounts of genetic etiology in patients with chromatin anomalies are not frequent. There are reports of abnormal removal of histones and transition proteins from sperm nuclei, selective absence of protamine P2 or altered ratios of nucleoproteins in spermatozoa from infertile individuals, but no or only occasional mutations in protamine genes have been documented93,95–99. Other nuclear abnormalities include macronuclear/multinuclear polyploid spermatozoa derived from meiotic alterations in nuclear cleavage. A familial pedigree with this anomaly has been reported100–102.

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ACQUIRED SPERM ABNORMALITIES SECONDARY TO ANDROLOGICAL CONDITIONS AND ENDOGENOUS OR ENVIRONMENTAL FACTORS

SPERM PATHOLOGY AND FERTILITY PROGNOSIS: THE SIGNIFICANCE OF SPERM PATHOLOGY IN THE STUDY OF INFERTILE MALES

Non-specific anomalies are the most frequent finding in astheno- and teratozoospermic patients. Non-specific flagellar anomalies are dealt with in the section on pathology of asthenozoospermia (see above). With regard to non-specific head anomalies, these constitute a heterogeneous condition in which various anomalies in the acrosome, chromatin, head cytoskeleton and the neck region coexist in different proportions. Their individualization in clinical andrology is based on the abnormal appearance of the spermatozoa. They constitute the foundation of all current classifications of sperm morphology, including those based on strict criteria. These classifications undoubtedly have an important application in predicting the fertility potential of a given semen sample. However, with the exception of acrosome anomalies that are taken into consideration in the classification presented by Kruger4,5, most head alterations are classified according to their external appearance, without any indication of the nature of the pathologies involved or the morphogenetic mechanisms that originate them. Alterations in chromatin maturation and compaction and insufficient development or vacuolization of the acrosome are frequent findings in amorphous sperm heads. They have been noted to be associated with inflammatory bowel disease80, varicocele103, contact with alkylated imino sugars or pesticides104,105, exposure to fuels, oils, organic solvents, exhaust fumes and hydrocarbons106, cigarette-smoking107, ionizing radiation108,109 or temperatures higher than physiological110. Even though there have been attempts to associate certain types of alterations with specific etiologies (e.g. tapered forms with varicocele1), this has not been confirmed and their non-specific nature is currently accepted.

It has been asserted that the results of ICSI are independent of most sperm parameters, but recent evidence indicates otherwise. Teratozoospermia should be understood not solely as a morphological abnormality but also as the corresponding impairment in sperm function. A higher pregnancy rate has been reported in coincidence with morphology values above the 4% threshold5, and various reports have stressed the importance of normal acrosome and chromatin structure, head–neck junction and centrosomes for adequate fertilization and pregnancy16,62,111–114. It has been claimed115,116 that abnormal morphology does not influence ICSI results, but in 10 of their 15 patients with total fertilization failure, strict morphology was ≤ 2%, and also failed fertilization was documented by these authors in six patients with acrosomeless spermatozoa115,116. In conclusion, many studies have shown that, depending on the nature of the pathologies involved, the outcome of ART can change dramatically. The recent introduction of ICSI provides access to the structural and functional features of spermatozoa that are being used for fertilization. This information can be applied to evaluate the relationship between sperm quality and fertility outcome, and hence a more objective picture is emerging of the differential roles played by specific sperm components in fertilization, early embryonic development and implantation.

Asthenozoospermia: flagellar pathologies and fertility prognosis As previously noted when discussing the subcellular basis of asthenozoospermia, increased rates of non-specific flagellar anomalies (NSFAs) were the underlying cause in 70% of 201 men with severe motility disorders (mean fast forward progression

SPERM PATHOLOGY: PROGNOSIS IN ASSISTED REPRODUCTION

3.6%)10. In patients with asthenozoospermia of genetic origin (fast forward progression 0.2%), specific sperm phenotypes such as primary ciliary dyskinesia and dysplasia of the fibrous sheath (PCD and DFS) were present in all spermatozoa12. Longitudinal studies in these men have shown that 33% of patients with NSFAs, but 0% of those with DFS, obtained fertilizations/pregnancies within 2–6 years of diagnosis, either spontaneously or with the use of ART, including IVF (but not ICSI). These findings indicate that onethird of cases of NSFA are reversible and can obtain fair fertility results, while DFS does not respond to conventional fertility treatments or IVF, as confirmed by the lack of other positive results in the literature. One publication by Kay and Irvine117 has documented a live birth after IVF using sperm with no progressive motility from a patient with primary ciliary dyskinesia. When there are 100% immotile sperm, a misleading tendency exists to equate complete asthenozoospermia with total necrozoospermia. This creates unnecessary confusion in view of the very different natures and fertility potentials of immotile (but live) and dead spermatozoa. Others have reported poor ICSI results with the use of ‘immotile spermatozoa’, but careful examination of the data indicates that, in their ‘immotile’ population, viability was always lower than 10%, which makes it very likely, as also noted by the authors, that their poor results were due to injection of dead spermatozoa (rather than live, immotile)115,116. ICSI has been of great help in cases of men with genetic asthenozoospermia. Indeed, there are now several publications reporting fertilizations/pregnancies with the use of immotile but live spermatozoa118–121. The difficulty in distinguishing between dead and completely immotile but live spermatozoa has been circumvented by various methods, including the hypo-osmotic swelling test, stimulation of motility with pentoxifylline or retrieving testicular spermatozoa122–125. We have recently reviewed numerous reports of ICSI results in 11 patients with PCD and 12

with DFS126. Fertilization was in the 55–70% range, and there were numerous pregnancies and 21 live births. The abortion rate was 20% (three of 15 pregnancies). The encouraging results indicate that this subpopulation of severe male-factor patients can expect good outcomes with microinjection of in situ motile or live, immotile spermatozoa. Therefore, flagellar pathologies causing sperm immotility do not compromise ICSI outcome if sperm viability is not affected. As stated before, DFS and PCD are genetic conditions, and there are concerns about the (possible) transmission of these anomalies to the next generation. Even though the number of cases is limited, there have been no reports of respiratory disease (a common finding in PCD and some DFS) in newborns. The question of fertility potential will have to remain unresolved for some years until the offspring attain reproductive age. Prospective parents should be made aware of the risks involved, but comprehensive genetic counseling will not be possible until the genes involved and the mechanism of inheritance are identified. Informed consent should always be obtained. Affected men tend to accept the risks if transmission of reproductive failure is the only concern, as is the case for individuals carrying Y-chromosome microdeletions that surely will pass to their male descendants.

Fertility potential in abnormalities of the connecting piece We have previously stated that, depending on the sperm anomalies involved, fertility outcomes change dramatically. This is illustrated by anomalies of the connecting piece, which, in contrast to the relatively good results attained in cases of flagellar pathology, are associated with a poor fertility prognosis in ICSI. Anomalies of the connecting piece have a heterogeneous phenotypic manifestation. In some of these patients, acephalic spermatozoa are the only form observed in semen, which makes impossible any attempt at fertilization. Other patients have

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acephalic forms in lower numbers, and spermatozoa with abnormal head–midpiece alignment predominate. Various recent ICSI procedures have been reported in these last patients. Chemes et al.59 documented the first ICSI failure using spermatozoa with a faulty alignment of the head–midpiece junction. Four metaphase II oocytes were fertilized by ICSI but remained at the pronuclear stage, and degenerated after failure to undergo syngamy and cleavage. Shortly after this there were two other failed attempts, with similar characteristics (Saias-Magnan et al.127, one patient, 1 cycle; Rawe et al.62, one patient, 5 cycles) and a further two with pregnancies and live deliveries (Porcu et al.63, two patients, five cycles, two pregnancies; Kamal et al.64, 16 patients, three pregnancies) as well as another successful attempt in one of our patients (personal unreported communication). In summary, from five reports available, four live births resulted from 26 cycles with numerous arrested or degenerated embryos. The question can be asked whether these different evolutions were connected with selection of the ‘right’ spermatozoon for injection. This seemed to be the case in one of our patients (five failed ICSI attempts), since two chemical pregnancies were obtained when the sperm selection criteria were very strict and the ‘best’ spermatozoa were microinjected. However, the two pregnancies reported by Porcu et al.63 seem to indicate otherwise, because the published morphology of the spermatozoa used for ICSI indicated a serious abnormality with severe misalignment at the head–midpiece junction.

Fertility outcome in men with acrosome and chromatin abnormalities Patients with acrosomeless spermatozoa are infertile because their spermatozoa are unable to penetrate oocytes due to the lack of acrosomes, physiologically involved in penetration of the cumulus oophorus that surrounds the oocyte and also in binding and penetration of the zona pellucida128. When ICSI was introduced, it was soon

hypothesized that, since microinjection bypasses all the penetration steps previous to fertilization, it may be an ideal solution for globozoospermia. The practice of ICSI with acrosomeless spermatozoa indicated that this was not exactly the case. While fertilization took place in a good number of instances, it failed in others, suggesting that besides penetration problems these spermatozoa may carry other deficiencies. Unsuccessful ICSI attempts in nine cases of acrosomeless spermatozoa were reported by Bourne et al.129, Liu et al.116, Battaglia et al.130 and Edirishinge et al.131. It was soon realized that the abnormality in cases of failure was probably due to insufficient activation of the oocyte, a function recently attributed to the perinuclear theca of spermatozoa. Indeed, acrosomeless spermatozoa have alterations of the perinuclear theca, and also lack various proteins associated with this structure (see above). Rybouchkin et al.132 and Kim et al.133 obtained successful pregnancies with acrosomeless spermatozoa by means of Ca2+ ionophore activation of the oocytes. However, artificially induced oocyte activation is not always followed by pregnancy130. Since chromatin anomalies are frequently associated with a lack of acrosome, their negative influence on fertilization should be taken into consideration. Besides these failures, there are also various reports of ICSI successes after microinjection of acrosomeless spermatozoa, but fertilization rates were low (10–50%)134–139. These results indicate that even though human acrosomeless spermatozoa are able to fertilize human or hamster oocytes and achieve pregnancies in numerous couples, they bear abnormalities responsible for unsuccessful or low fertilization rates or the need for artificial activation. The maturational changes that chromatin undergoes during spermiogenesis are an essential component of its fertilizing capacity. Spermatozoa with amorphous, elongated or round heads have been shown to have a four-fold increase in chromosomal abnormalities87. Large, intranuclear, hypodense regions (incorrectly called ‘nuclear vacuoles’) represent areas in which the DNA itself or

SPERM PATHOLOGY: PROGNOSIS IN ASSISTED REPRODUCTION

the associated proteins have structural abnormalities. DNA breaks, single-stranded DNA, deletions of variable magnitude and other alterations significantly affect sperm quality, fertilization, embryo development and implantation. Infertility or abortions during the first trimester have been reported in these patients16,79,80,140. Similar results were reported by Francavilla et al.141 when comparing the results of 21 testicular sperm extraction (TESE)–ICSI cycles in azoospermic men with or without chromatin abnormalities. While the fertilization rate was similar in both groups, the delivery rate per cycle was significantly diminished in men with chromatin abnormalities. Others have also reported normal fertilization rates and low pregnancy rates in a study of 17 males with megalohead multitailed spermatozoa that have been shown to be polyploid142. Careful selection of motile spermatozoa for ICSI by means of very high-resolution light microscopy yields dramatic differences in implantation and pregnancy rates between normal spermatozoa and those with ‘nuclear vacuoles’ (indicative of abnormal chromatin constitution)143. The negative influence of DNA fragmentation on ICSI outcome was reported by Greco et al.144 in men with high rates of DNA fragmentation, by comparing ICSI with testicular spermatozoa (low DNA damage) versus ejaculated spermatozoa (found to have high DNA damage).

CONCLUDING REMARKS Sperm pathology is the discipline that characterizes structural and functional deficiencies in spermatozoa. It is not just another denomination for abnormal sperm morphology; it is rather a new concept in which a multidisciplinary approach is applied to the precise description of sperm abnormalities and the understanding of the pathogenic mechanisms that underlie abnormal sperm appearance. Used jointly with classical sperm morphology (in particular the strict criteria), it allows a clear appreciation of what is wrong with

abnormal sperm shapes and facilitates a rational approach to the use of abnormal spermatozoa in assisted reproduction. The distinction between non-specific anomalies and systematic defects of genetic origin is an important one, and couples undergoing ICSI have the right to be informed not only of their diminished chances when this is the case, but also of the possible risk of transmission to their offspring. Whenever possible, genetic counseling is important and follow-up of newborns desirable. However, in view of our present uncertainties, care should be taken to protect patients from excessive information, particularly when no unambiguous conclusions are available. Another important issue refers to the use of appropriate nomenclature, previously addressed by Chemes and Rawe126. We have attempted to highlight each pathological phenotype with a denomination that identifies the organelles involved and the pathogenic mechanisms. The problem of nomenclature is not a trivial one: the way we speak and write conditions the way we think. If descriptive terms are used, thoughts will not go beyond appearances. It is essential to distinguish a dead (immotile) from an immotile (live) spermatozoon, and to use denominations that give us the basic understanding of each pathology. A ‘stump tail’ can either belong to a DFS spermatozoon or be the result of tail disintegration in aging spermatozoa; an ‘amorphous’ head can correspond to a lack of acrosome or to abnormal chromatin maturation and compaction. The introduction of innovative therapeutic approaches such as ICSI has revolutionized the field of reproductive medicine. Besides its obvious advantages for men with severe male factor infertility, it has created new concerns about the ethical and social role of therapeutic interventions. The possibility of inherited sterility is certainly one of the most perplexing paradoxes of our times.

ACKNOWLEDGMENTS The present chapter including most of its information is based on our previous publication:

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Chemes HE, Rawe VY. Sperm pathology: a step beyond descriptive morphology. Origin, characterization and fertility potential of abnormal sperm phenotypes in infertile men. Hum Reprod Update 2003; 9: 405. Figures 6.1a–d and 6.3c–e in the present chapter are taken from the same paper. Copyrights European Society of Human Reproduction and Embryology. Reproduced by permission of Oxford University Press/Human Reproduction. This work was supported by grants from the National Research Council (PIP 0900 and 4584), ANPCyT (PICT 9591) and CEGyR Foundation.

11.

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associated with sperm immotility and primary sterility. Fertil Steril 1987; 48: 664 Baccetti B, et al. The short-tailed human spermatozoa, ultrastructural alterations and dynein absence. J Submicrosc Cytol 1975; 7: 349 Baccetti B, et al. Genetic sperm defects and consanguinity. Hum Reprod 2001; 16: 1365 Alexandre C, Bisson JP, David G. Asthenospermie totale avec anomalie ultrastructurale du flagelle chez deux frères stériles. J Gynecol Obstet Biol Reprod 1978; 7: 31 Bisson JP, Leonard C, David G. Caractère familial de certaines perturbations morphologiques des spermatozoides. Arch Anat Cytol Pathol 1979; 27: 230 Carrera A, Gerton GL, Moss SB. The major fibrous sheath polypeptide of mouse sperm: structural and functional similarities to the A-kinase anchoring proteins. Dev Biol 1994; 165: 272 Turner RMO, et al. Molecular genetic analysis of two human sperm fibrous sheath proteins, AKAP4 and AKAP3, in men with dysplasia of the fibrous sheath. J Androl 2001; 22: 302 Miki K, et al. Targeted disruption of the Akap4 gene causes defects in sperm flagellum and motility. Dev Biol 2002; 248: 331 Baccetti B, et al. Gene deletions in an infertile man with sperm fibrous sheath dysplasia. Hum Reprod 2005; 20: 2790 Bonneau D, et al. Usher syndrome type I associated with bronchiectasis and immotile nasal cilia in two brothers. J Med Genet 1993; 30: 253 Rawe VY, et al. Abnormal organization of mitochondrial sheaths in two cases of severe asthenozoospermia. Int J Androl 2005; 28 (Suppl 1): 88 Holyoake AJ, et al. High incidence of single nucleotide substitutions in the mitochondrial genome is associated with poor semen parameters in men. Int J Androl 2001; 243: 175 Thangaraj K, et al. Sperm mitochondrial mutations as a cause of low sperm motility. J Androl 2003; 24: 388 Panidis D, et al. Headless spermatozoa in semen specimens from fertile and subfertile men. J Reprod Med 2001; 46: 947 Le Lannou D. Teratozoospermie consistant en l’absence de tete spermatique par defaut de connexion. J Gynecol Obstet Biol Reprod (Paris) 1979; 8: 43 Perotti ME, Gioria M. Fine structure and morphogenesis of ‘headless’ human spermatozoa associated with infertility. Cell Biol Int Rep 1981; 5: 113

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57. Baccetti B, Selmi MG, Soldani P. Morphogenesis of ‘decapitated spermatozoa’ in a man. J Reprod Fertil 1984; 70: 395 58. Holstein AF, Schill WB, Breucker H. Dissociated centriole development as a cause of spermatid malformation in a man. J Reprod Fertil 1986; 78: 719 59. Chemes HE, et al. Acephalic spermatozoa and abnormal development of the head–neck attachment. A human syndrome of genetic origin. Hum Reprod 1999; 14: 1811 60. Baccetti B, et al. Morphogenesis of the decapitated and decaudated sperm defect in two brothers. Gamete Res 1989; 23: 181 61. Perotti ME, Giarola A, Gioria M. Ultrastructural study of the decapitated sperm defect in an infertile man. J Reprod Fertil 1981; 63: 543 62. Rawe VY, et al. A pathology of the sperm centriole responsible for defective sperm aster formation, syngamy and cleavage. Hum Reprod 2002; 17: 2344 63. Porcu G, et al. Pregnancies after ICSI using sperm with abnormal head–tail junction from two brothers: case report. Hum Reprod 2003; 18: 562 64. Kamal A, et al. Easily decapitated spermatozoa defect: a possible cause of unexplained infertility. Hum Reprod 1999; 14: 2791 65. Manandhar G, Schatten G. Centrosome reduction during rhesus spermiogenesis: gamma-tubulin, centrin, and centriole degeneration. Mol Reprod Dev 2000; 56: 502 66. Wojcik C, et al. Proteasomes in human spermatozoa. Int J Androl 2000; 23: 169 67. Rawe VY. Proteasome function in mammalian fertilization: implications ferilization faileur in humans. Int J Androl 2005; 28 (Suppl 1): 13 68. Morales P, et al. Decreased proteasome enzymatic activity in sperm from patients with genetic abnormalities of the head–tail junction and acephalic spermatozoa. J Androl 2004; 25 (March–April Suppl): 41 69. Kalahanis J, et al. Round-headed spermatozoa in semen specimens from fertile and subfertile men. J Reprod Med 2002; 47: 489 70. Baccetti B, et al. Further observations on the morphogenesis of the round-headed human spermatozoa. Andrologia 1977; 9: 255 71. Holstein AF, Schirren C. Classification of abnormalities in human spermatids based on recent advances in ultrastructure research on spermatid differentiation. In Fawcett DW, Bedford JM, eds. The Spermatozoon: Maturation, Motility, Surface Properties and Comparative Aspects. Baltimore: Urban and Schwarzenberg, 1979: 341

72. Florke-Gerloff S, et al. Biochemical and genetic investigation of round-headed spermatozoa in infertile men including two brothers and their father. Andrologia 1984; 16: 187 73. Florke-Gerloff S, et al. On the teratogenesis of round-headed spermatozoa: investigations with antibodies against acrosin, an intraacrosomally located acrosin-inhibitor, and the outer acrosomal membrane. Andrologia 1985; 17: 126 74. Longo FJ, Krohne G, Franke WW. Basic proteins of the perinuclear theca of mammalian spermatozoa and spermatids: a novel class of cytoskeletal elements. J Cell Biol 1987; 105: 1105 75. Sutovsky P, et al. The removal of the sperm perinuclear theca and its association with the bovine oocyte surface during fertilization. Dev Biol 1997; 188: 75 76. Sutovsky P, et al. Interactions of sperm perinuclear theca with the oocyte: implications for oocyte activation, anti-polyspermy defense, and assisted reproduction Microsc Res Tech 2003; 61: 362 77. Oko R, et al. The sperm head cytoskeleton. In Robaire B, Chemes H, Morales C, eds. Andrology in the 21st Century. Englewood: Medimond Publishing Company, 2001: 37 78. Nistal M, Herruzo A, Sanchez Corral F. Teratozoospermia absoluta de presentación familiar: espermatozoides microcéfalos irregulares sin acrosoma. Andrologia 1978; 10: 234 79. Zamboni L. The ultrastructural pathology of the spermatozoon as a cause of infertility: the role of electron microscopy in the evaluation of semen quality. Fertil Steril 1987; 48: 711 80. Zamboni L. Sperm structure and its relevance to infertility. An electron microscopic study. Arch Pathol Lab Med 1992; 116: 325 81. Liu DY, Baker HW. Disordered acrosome reaction of spermatozoa bound to the zona pellucida: a newly discovered sperm defect causing infertility with reduced sperm-zona pellucida penetration and reduced fertilization in vitro. Hum Reprod 1994; 9: 1694 82. Baccetti B. et al. Crater defect in human spermatozoa. Gamete Res 1989; 22: 249 83. Balhorn R. A model for the structure of chromatin in mammalian sperm. J Cell. Biol 1982; 93: 298 84. Ward WS, Coffey DS. DNA packaging and organization in mammalian spermatozoa: comparison with somatic cells. Biol Reprod 1991; 44: 569 85. Brewer L, Corzett M, Balhorn R. Condensation of DNA by spermatid basic nuclear proteins. J Biol Chem 2002; 277: 38895

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86. Holstein AF. Morphologische Studien an abnormen Spermatiden und Spermatozoen des Menschen. Virchows Arch 1975; 367: 93 87. Lee JD, Kamiguchi Y, Yanagimachi R. Analysis of chromosome constitution of human spermatozoa with normal and aberrant head morphologies after injection into mouse oocytes. Hum Reprod 1996; 11: 1942 88. Calogero AE, et al. Aneuploidy rate in spermatozoa of selected men with abnormal semen parameters. Hum Reprod 2001; 16: 1172 89. Kovanci E, et al. FISH assessment of aneuploidy frequencies in mature and immature human spermatozoa classified by the absence or presence of cytoplasmic retention. Hum Reprod 2001; 16: 1209 90. Lewis-Jones I, et al. Sperm chromosomal abnormalities are linked to sperm morphologic deformities. Fertil Steril 2003; 1: 212 91. Vicari E, et al. Absolute polymorphic teratozoospermia in patients with oligo-asthenozoospermia is associated with an elevated sperm aneuploidy rate. J Androl 2003; 4: 598 92. Hrudka F, Singh A. Sperm nucleomalacia in men with inflammatory bowel disease. Arch Androl 1984; 13: 37 93. Schlicker M, et al. Disturbances of nuclear condensation in human spermatozoa: search for mutations in the genes for protamine 1, protamine 2 and transition protein 1. Hum Reprod 1994; 9: 2313 94. Evenson DP, et al. Characteristics of human sperm chromatin structure following an episode of influenza and high fever: a case study. J Androl 2000; 21: 739 95. Tanaka H, et al. Single-nucleotide polymorphisms in the protamine-1 and -2 genes of fertile and infertile human male populations. Mol Hum Reprod 2003; 9: 69 96. Balhorn R, Reed S, Tanphaichitr N. Aberrant protamine 1/protamine 2 ratios in sperm of infertile human males. Experientia 1988; 44: 52 97. Blanchard Y, Lescoat D, Le Lannou D. Anomalous distribution of nuclear basic proteins in roundheaded human spermatozoa. Andrologia 1990; 22: 549 98. de Yebra L, et al. Complete selective absence of protamine P2 in humans. J Biol Chem 1993; 268: 10553 99. de Yebra L, et al. Detection of P2 precursors in the sperm cells of infertile patients who have reduced protamine P2 levels. Fertil Steril 1998; 69: 755

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100. Escalier D. Human spermatozoa with large heads and multiple flagella: a quantitative ultrastructural study of 6 cases. Biol Cell 1983; 48: 65 101. Benzacken B, et al. Familial sperm polyploidy induced by genetic spermatogenesis failure. Hum Reprod 2001; 16: 2646 102. Devillard F, et al. Polyploidy in large-headed sperm: FISH study of three cases. Hum Reprod 2002; 17: 1292 103. Muratori M, et al. Functional and structural features of DNA-fragmented human sperm. J Androl 2000; 21: 903 104. ElJack AH, Hrudka F. Patterns and dynamics of teratospermia induced in rams by parenteral treatment with ethylene dibromide. J Ultrastruct Res 1979; 67: 124 105. Bustos-Obregon E, Diaz O, Sobarzo C. Parathion induces mouse germ cells apoptosis. Ital J Embryol 2001; 2: 199 106. Harrison KL. Semen parameter defects and toxin contact related occupation in infertility patients. Middle East Fertil Soc J 1998; 3: 3 107. Banerjee A, et al. Semen characteristics of tobacco users in India. Arch Androl 1993; 30: 35 108. Sailer BL, et al. Effects of X-irradiation on mouse testicular cells and sperm chromatin structure. Environ Mol Mutagen 1995; 25: 23 109. Schevchenko VA, et al. Genetic effects of 131I in reproductive cells of male mice. Mutat Res 1989; 226: 87 110. Mieusset R. Influence of lifestyle on male infertility: potential testicular heating factors. Middle East Fertil Soc J 1998; 3 (Suppl 1): 40 111. Tasdemir I, et al. Effect of abnormal sperm head morphology on the outcome of intracytoplasmic sperm injection in humans. Hum Reprod 1997; 12: 1214 112. Oehninger S, et al. Failure of fertilization in in vitro fertilization: the ‘occult’ male factor. J In Vitro Fert Embryo Transf 1988; 5: 181 113. Nikolettos N, et al. Fertilization potential of spermatozoa with abnormal morphology. Hum Reprod 1999; 14: 47 114. Bartoov B, et al. Real-time fine morphology of motile human sperm cells is associated with IVF–ICSI outcome. J Androl 2002; 23: 1 115. Nagy ZP, et al. The result of intracytoplasmic sperm injection is not related to any of the three basic sperm parameters. Hum Reprod 1995; 10: 1123 116. Liu J, et al. Analysis of 76 total fertilization failure cycles out of 2732 intracytoplasmic sperm injection cycles. Hum Reprod 1995; 10: 2630

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117. Kay VJ, Irvine DS. Successful in-vitro fertilization pregnancy with spermatozoa from a patient with Kartagener’s syndrome: case report. Hum Reprod 2000; 15: 135 118. Nijs M, et al. Fertilizing ability of immotile spermatozoa after intracytoplasmic sperm injection. Hum Reprod 1996; 11: 2180 119. Barros A, et al. Pregnancy and birth after intracytoplasmic sperm injection with totally immotile sperm recovered from the ejaculate. Fertil Steril 1997; 67: 1091 120. Kahraman S, et al. A healthy birth after intracytoplasmic sperm injection by using immotile testicular spermatozoa in a case with totally immotile ejaculated spermatozoa before and after Percoll gradients. Hum Reprod 1997; 12: 292 121. von Zumbusch A, et al. Birth of healthy children after intracytoplasmic sperm injection in two couples with male Kartagener’s syndrome. Fertil Steril 1998; 70: 643 122. Kahraman S, et al. Pregnancies achieved with testicular and ejaculated spermatozoa in combination with intracytoplasmic sperm injection in men with totally or initially immotile spermatozoa in the ejaculate. Hum Reprod 1996; 11: 1343 123. Ved S, et al. Pregnancy following intracytoplasmic sperm injection of immotile spermatozoa selected by the hypo-osmotic swelling-test: a case report. Andrologia 1997; 29: 241 124. Wang CW, et al. Pregnancy after intracytoplasmic injection of immotile sperm. A case report. J Reprod Med 1997; 42: 448 125. Terriou P, et al. Pentoxifylline initiates motility in spontaneously immotile epididymal and testicular spermatozoa and allows normal fertilization, pregnancy, and birth after intracytoplasmic sperm injection. J Assist Reprod Genet 2000; 17: 194 126. Chemes HE, Rawe VY. Sperm pathology: a step beyond descriptive morphology. Origin, characterization and fertility potential of abnormal sperm phenotypes in infertile men. Hum Reprod Update 2003; 9: 405 127. Saias-Magnan J, et al. Failure of pregnancy after intracytoplasmic sperm injection with decapitated spermatozoa: case report. Hum Reprod 1999; 14: 1989 128. Schmiady H, Radke E, Kentenich H. Roundheaded spermatozoa – contraindication for IVF. Geburtshilfe Frauenheilkd 1992; 52: 301

129. Bourne H, et al. Normal fertilization and embryo development by intracytoplasmic sperm injection of round-headed acrosomeless sperm. Fertil Steril 1995; 63: 1329 130. Battaglia DE, et al. Failure of oocyte activation after intracytoplasmic sperm injection using roundheaded sperm. Fertil Steril 1997; 68: 118 131. Edirisinghe WR, et al. Cytogenetic analysis of unfertilized oocytes following intracytoplasmic sperm injection using spermatozoa from globozoospermic man. Hum Reprod 1998; 13: 3094 132. Rybouchkin A, et al. Disintegration of chromosomes in dead sperm cells as revealed by injection into mouse oocytes. Hum Reprod 1997; 12: 1693 133. Kim ST, et al. Successful pregnancy and delivery from frozen–thawed embryos after intracytoplasmic sperm injection using round-headed spermatozoa and assisted oocyte activation in a globozoosperic patient with mosaic Down syndrome. Fertil Steril 2001; 75: 445 134. Lunding K, et al. Fertilization and pregnancy after intracytoplasmic microinjection of acrosomeless spermatozoa. Fertil Steril 1994; 62: 1266 135. Liu J, et al. Successful fertilization and establishment of pregnancies after intracytoplasmic sperm injection in patients with globozoospermia. Hum Reprod 1995; 10: 626 136. Trokoudes KM, et al. Pregnancy with spermatozoa from a globozoospermic man after intracytoplasmic sperm injection. Hum Reprod 1995; 10: 880 137. Stone S, et al. A normal livebirth after intracytoplasmic sperm injection for globozoospermia without assisted oocyte activation: case report. Hum Reprod 2000; 15: 139 138. Nardo LG, et al. Ultrastructural features and ICSI treatment of severe teratozoospermia: report of two human cases of globozoospermia. Eur J Obstet Gynecol Reprod Biol 2002; 104: 40 139. Zeyneloglu HB, et al. Achievement of pregnancy in globozoospermia with Y chromosome microdeletion after ICSI. Hum Reprod 2002; 17: 1833 140. Francavilla S, et al. Chromatin defects in normal and malformed human ejaculated and epididymal spermatozoa: a cytochemical ultrastructural study. J Reprod Fertil 1996; 106: 259 141. Francavilla S, et al. Ultrastructural analysis of chromatin defects in testicular spermatids in azoospermic men submitted to TESE–ICSI. Hum Reprod 2001; 16: 1440

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142. Kahraman S, et al. Fertility of ejaculated and testicular megalohead spermatozoa with intracytoplasmic sperm injection. Hum Reprod 1999; 14: 726 143. Bartoov B, et al. High power light microscope morphological examination as a tool for single sperm

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selection pryor IVF–ICSI. Int J Androl 2005; 28 (Suppl 1): 8 144. Greco E, et al. Efficient treatment of infertility due to sperm DNA damage by ICSI with testicular spermatozoa. Hum Reprod 2005; 20: 226

7 Testicular dysgenesis syndrome: biological and clinical significance Niels Jørgensen, Camilla Asklund, Katrine Bay, Niels E Skakkebæk

INTRODUCTION

affected may show only reduced spermatogenesis which is fully compatible with fertility1. Consequently, a person diagnosed with one of the TDS symptoms must be considered at increased risk of harboring one or more of the other symptoms as well.

A few years ago it was suggested that testicular cancer, hypospadias, cryptorchidism and low sperm counts were all symptoms of a disease complex, the testicular dysgenesis syndrome (TDS), with a common origin in fetal life1 (Figure 7.1). Knowledge of the etiology of TDS is still rather limited, but environmental and life-style factors are suggested as contributing agents. However, genetic polymorphisms or aberrations may render some individuals particularly susceptible to these exogenous factors. The most severe cases of TDS may include all four symptoms, whereas the least

Environmental factors including endocrine disruptors

PRENATAL ORIGIN OF TESTICULAR DYSGENESIS SYNDROME The prenatal origin of hypospadias and cryptorchidism is evident, owing to their congenital nature. However, testicular cancers that do not

Disturbed Sertoli cell function

Impaired germ cell differentiation

Decrease Leydig cell function

Androgen insufficiency

Reduced semen quality

CIS

Testicular cancer

Testicular dysgenesis Hypospadias

Genetic defects including 45,X/46,XY and point mutations

Testicular maldescent

Figure 7.1 Schematic presentation of the components and clinical manifestations of testicular dysgenesis syndrome. CIS, carcinoma in situ. Adapted with permission from reference 1

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manifest until later in life are most probably also of fetal origin. Likewise, the potential of a man’s semen quality may also be determined prenatally.

Spermatogenesis At the beginning of the fourth week of fetal development, germ cells begin to migrate via the yolk sac through the gut and into the mesentery, ending in the celomic epithelium of the gonadal ridges2. The indifferent gonad is composed of three cell types: germ cells, supporting cells, which in the male fetus give rise to Sertoli cells, and stromal (interstitial) cells. The first sign of gonadal differentiation is development of the Sertoli cells and their aggregation into primitive seminiferous cords during the eighth week of development3. Differentiation of the gonad into a testis rather than an ovary is genetically dependent on the SRY gene (the gene of the sex-determining region of the Y chromosome), which is expressed by testicular (Sertoli) cells4. The majority of the Sertoli cell multiplication occurs during fetal life, and only to a lesser extent later5. The final number of Sertoli cells reached during development has consequences in adult life, as these cells can only support a limited number of germ cells6,7. Thus,

factors affecting Sertoli cell development and function during fetal life will have important consequences for a man’s future spermatogenic capacity, as the number of Sertoli cells essentially determines the maximal achievable sperm output. The final sperm output may, however, be adversely influenced by postnatal factors such as irradiation, medical treatment, pesticides, organic solvents, metals and physical agents.

Testicular cancer Testicular germ-cell cancers occurring from puberty and onwards originate from preinvasive carcinoma in situ of the testis (CIS) cells, which are considered to be gonocyte-like transformed germ cells that failed to differentiate during the fetal period8,9. CIS cells have stem-cell properties, as evident from the expression of a number of genes also expressed by gonocytes and embryonic stem cells, for example alkaline phosphatase, c-kit, Oct-4, SSEA-3 (stage-specific embryonic antigen 3) and others8,10–13 (Figure 7.2). Furthermore, CIS cells and gonocytes lack expression of other genes that are specific for postmeiotic germ cells14. Clinical data also indicate that CIS cells arise before adult life15, and CIS cells have been detected even

Figure 7.2 Immunohistochemical staining with placental-like alkaline phosphatase (PLAP). (a) Expression in normal, immature germ cells of 9-week-old fetal testis, and (b) expression in an adult testis with carcinoma in situ cells. Note in both images that the immunohistochemical reaction is not seen in Sertoli cells or interstitial cells

TESTICULAR DYSGENESIS SYNDROME

in the neonatal period16. Epidemiologically, it has been shown that Danish and other Scandinavian men born during the Second World War have a lower risk in all age groups of developing germ-cell tumors than expected from the overall trend in incidences, indicating that important etiological events take place during prenatal life17,18.

Hypospadias and cryptorchidism The secondary sex characteristics are dependent on hormones produced by the newly formed testicles. Testosterone is secreted by the fetal Leydig cells, and is responsible for differentiation of the Wolffian duct into the epididymis, vas deferens and seminal vesicle19. Testosterone is converted to 5α-dihydrotestosterone at the bipotential external genitalia, and stimulates formation of the penile urethra, the penis and the scrotum. Decreased testosterone secretion may lead to formation disturbances, resulting in hypospadias20, for example. Testicular descent appears in two phases. The intra-abdominal descent is quite complex, and its regulation is not fully understood; however, it occurs in the second trimester and is largely dependent on the Leydig cell hormone insulinlike factor 3 (INSL3)21. The following descent through the inguinal canal and into the scrotum is dependent on adequate testosterone secretion22. Thus, impaired INSL3 and/or testosterone may lead to cryptorchidism.

RISK FACTORS FOR TESTICULAR DYSGENESIS SYNDROME Many investigators have found that the TDS symptoms are to be regarded as risk factors for each other, and frequently, patients present with more than one of the symptoms. The association between testicular cancer and low semen quality is firmly established. CIS cells were first detected in infertile men23, and later Berthelsen showed reduced spermatogenesis in testicles contralateral to testicular cancer already before treatment of the

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cancer24,25. More recently, men with unilateral testicular cancer have been shown to have poorer semen quality than expected from a man with only one functioning testicle26. Epidemiologically, the link between testicular cancer and low semen quality has indirectly been confirmed by the detection of reduced fertility in men who later developed testicular cancer27. At the histological level, the non-tumor-bearing testicles in men with testicular cancer often show carcinoma in situ (5–8%), Sertoli-cell-only tubules (13.8%), microcalcifications (6.0%) and undifferentiated Sertoli cells (4.6%). All in all, signs of histological testicular dysgenesis were detected in 25.2% of the examined contralateral testes28. Cryptorchidism is a well-known risk factor for both testicular cancer and poor semen quality29–31, and the association between cryptorchidism and hypospadias is well documented31,32. The associations between the four TDS symptoms point to abnormal germ-cell and/or Sertolicell development during fetal life, and are all coupled to the intrauterine milieu, such as low birth weight, premature birth and low parity33–35. Genetic factors seem to contribute, as indicated by the fact that African-Americans have significantly lower incidence than Caucasians living in the same areas of the USA36,37. Additionally, patients with genetic disorders such as 45,X/46,XY mosaicism or androgen insensitivity syndrome often show testicular dysgenesis due to impaired androgen production or function already in fetal life, increased risk of cryptorchidism, testicular cancer and impaired spermatogenesis. The genetic mechanism(s) behind this is still unresolved; however, genes on the Y chromosome seem to be important for proper testicular function38,39.

REGIONAL AND TEMPORAL TRENDS IN TESTICULAR DYSGENESIS SYNDROME SYMPTOMS For many years, the incidence of testicular germcell cancer has increased in numerous European

MALE INFERTILITY

(a) Median sperm concentration (× 106/ml)

countries40. In particular, the situation in the two Nordic countries Denmark and Finland is remarkable. Danish men have one of the highest incidences of testicular cancer, and Finnish men one of the lowest incidences (11.1 per 105 and 2.8 per 105, respectively)41. The sharp increase in incidence among Danish men shows a birth cohortdependency, as men born recently have a higher lifetime risk than men born in previous decades17,42. In line with the geographical trends observed for testicular cancer, the prevalence of cryptorchidism and hypospadias in Finnish newborn boys is considerably lower than in Danish boys (2.4% vs. 9.0% and 0.27% vs. 1.03%, respectively)43,44. Semen quality also shows a regional difference, with a better situation among Finnish than among Danish men. Young, normal men from the Danish general population have a median sperm concentration of 41 × 106/ml in contrast to Finnish men having 54 × 106/ml45, and overall an East–West gradient in sperm concentration exists in the Nordic–Baltic area45–48, with a better situation in the eastern than in the western part (Figure 7.3). There are, however, indications that the otherwise good reproductive health of Finnish men is also following a worsening tendency. Despite being low, the testicular cancer incidence is increasing40, while the sperm count may be decreasing45. In 1992, Carlsen and co-workers49 reported the results of a meta-analysis of previously published semen quality data, and indicated that sperm concentration among men in Europe and North America had decreased. Following this, reports from several other research groups were published. Some did not find any change over time50–53, whereas others suggested that sperm counts had declined significantly54–57, and thereby also indicated the presence of geographical differences in the adverse male reproductive-health trends. Associations between the individual TDS symptoms are seen not only in Danish and Finnish populations. Norwegian men have a high frequency of testicular cancer and low sperm

80 70 60 50 40 30 20 10 0 Finland

Estonia

Norway

Denmark

Finland

Estonia

Norway

Denmark

(b) Median normal spermatozoa (%)

108

12 10 8 6 4 2 0

Figure 7.3 Illustration of the regional difference in (a) sperm concentration (adjusted for period of abstinence and interlaboratory variation) and (b) frequency of morphologically normal spermatozoa of young men from the Nordic–Baltic area. Bars indicate median values and 95% confidence level of the estimates (from linear regression models taking confounders into account). See text for further explanation. Adapted from reference 45. Results from references 47 and 48 are not included as the presentations in these publications do not provide sufficient information to draw similar bars

counts, whereas the opposite is true for Estonian and Lithuanian men45,46. Unfortunately, very limited information exists from countries outside the Nordic–Baltic area to elucidate the occurrence of TDS. Japanese fertile men seem to have semen quality at the same level as that of comparable Danish fertile men, but at the same time Japanese men have a risk of testicular cancer at or below the level in Finnish men40,58. This finding is compatible with Japanese (or Asian) men having a lower sperm quality, without being at increased risk for the other symptoms of TDS. However, a more thorough analysis is needed before any firm conclusions can be reached.

TESTICULAR DYSGENESIS SYNDROME

Studies have revealed the existence of regional differences in semen quality among fertile US men59, whereas other studies have shown AfricanAmericans having significantly lower incidences of testicular cancer than Caucasians living in the same areas37. These results are compatible with both an environmental and a genetic influence on male reproductive health, but the studies cannot provide any firm information about associations between the different TDS symptoms among US men. Likewise, data from other countries outside Northern Europe are lacking.

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that men with fewer than 9% morphologically normal spermatozoa belong to a group of subfertile men, and that men ought to have more than 12% normal forms to be regarded as fertile63. The Danish and Norwegian young, normal men had only a few more than 6% (median) normal forms45. The majority of these young men were 19 years of age; however, a follow-up study of these men indicated that their low semen quality is unlikely to be a result of immaturity64.

TESTICULAR DYSGENESIS SYNDROME AND FECUNDITY

POSSIBLE LIFE-STYLE OR ENVIRONMENTAL FACTORS CAUSING IMPAIRED MALE REPRODUCTIVE HEALTH

It is of concern that the birth rate in many industrialized countries has declined to below replacement level of the populations. The social structure in these countries acts against a high birth rate, but it is becoming clearer that reduced biological fecundity may also be considered an important contributing factor. The World Health Organization (WHO) states that the reference value for sperm concentration is 20 × 106 spermatozoa/ml60. Whether this is a relevant ‘threshold’ can be questioned) owing to the findings of a prospective study of fecundity. Decreasing waiting time to pregnancy (TTP) with increasing sperm concentrations up to approximately 40 × 106 spermatozoa/ml was shown61. Additionally, a recent crosssectional study of European fertile men demonstrated a reduced TTP with increasing sperm concentration up to 55 × 106 spermatozoa/ml62. Thus, a large fraction of normal young Danish and Norwegian men may already have a semen quality with sperm concentrations below these levels; 20% of the investigated Danish and Norwegian men had a sperm concentration below the WHO reference level, and approximately 40% of the men had fewer than 40 × 106 spermatozoa/ml. Sperm concentration is only one of the parameters having an impact on fecundity. A recent publication by Guzick et al. has indicated

It is possible that genetic predisposition may play a partial role in the observed trends in male reproductive health, at least for some populations. For example, impaired spermatogenesis has in some studies been associated with polymorphisms in the androgen receptor gene or in the Y chromosome39,65. The speed of the observed increase in testicular cancer indicates that life-style or environmental factors may also be contributing agents. Furthermore, poor semen quality, cryptorchidism or hypospadias – at least in some areas – have become more frequent40,43,45, and thus exogenous etiological factors are likely. Three recent studies detected that men exposed to smoking in utero (via maternal smoking during pregnancy) had decreased sperm concentrations: a 20% reduction compared with men not exposed at all66, a 48% reduction among sons exposed to maternal smoking of more than ten cigarettes per day67 and a dose-dependent association between fetal tobacco exposure, lower semen quality and higher risk of oligozoospermia68. The men’s own history of tobacco-smoking was shown to be only of minor importance when taking into account mothers smoking while pregnant. Obesity has increased in the Western world, and a body mass index (BMI) above 25 kg/m2 has been associated with reductions in sperm

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concentration, total sperm count and morphologically normal spermatozoa69. Any causal relationship between semen quality and BMI has yet to be resolved. Generally, TDS is suggested to result from disruption of fetal gonadal development caused by endocrine-disrupting compounds. During recent years, focus has been on the possible disruption of the androgen–estrogen balance and impaired androgen action33,70. Recent experimental evidence comes from a possible animal model for TDS, in which rats exposed in utero to the antiandrogen dibutyl phthalate developed cryptorchidism, hypospadias, infertility and testis abnormalities71,72. The finding that phthalates can induce TDS-like symptoms is of concern, as neonates can be exposed to considerable daily doses of phthalates via breastmilk73,74. Moreover, many of the so-called ‘environmental estrogens’, including a number of pesticides, have also appeared to possess antiandrogenic properties75. In the absence of possibilities to provide evidence of a causal relationship between human exposure to harmful chemicals and male reproductive health, rising concern has led to a number of epidemiological studies dealing with associations between parental exposure to substances with endocrine-disrupting properties and congenital abnormalities in the reproductive organs of their sons34,76–80. Interestingly, an American group has recently published a report using shortening of the anogenital distance (AGD) as a new and more sensitive marker for demasculinization in humans. In 134 boys aged 2–30 months they found a significant inverse correlation between AGD and urinary concentrations of a number of phthalate metabolites81.

CONCLUSIONS Testicular dysgenesis syndrome (TDS) encompasses the disease entities cryptorchidism, hypospadias, testicular cancer and poor semen

quality. Exogenous factors exhibiting antiandrogenic properties or reducing androgen/estrogen functions are suspected to affect the developing fetal gonad, leading to the TDS symptoms. However, a genetic susceptibility to these exposures may contribute to the development. In its most severe form, a man may suffer from all the TDS symptoms, whereas the least affected may only have a slightly reduced semen quality, compatible with fertility. Most likely, all cases of testicular germ-cell cancers are due to TDS. The three other symptoms may also be due to TDS; however, alternative contributing factors may be relevant. A man’s potential semen quality may already be determined prenatally, but may be adversely affected by factors acting postnatally. Nevertheless, diagnosis of one of the TDS symptoms should alert physicians to look for manifestations of the other symptoms, especially the occurrence of preinvasive carcinoma in situ germ cells. Eradication of these cells will prevent the development of overt testicular cancer82.

REFERENCES 1. Skakkebaek NE, Rajpert-De ME, Main KM. Testicular dysgenesis syndrome: an increasingly common developmental disorder with environmental aspects. Hum Reprod 2001; 16: 972 2. Witschi E. Migration of the germ cells of human embryos from the yolk sac to the primitive gonadal folds. Contributions to Embryology, No. 209. Washington, DC: Carnegie Institute of Washington, 1948: 69 3. Jirasek JE. Principles of reproductive embryology. In Simpson JL, ed. Disorders of Sexual Differentiation. New York: Academic Press, 1976: 51 4. Gubbay J, et al. A gene mapping to the sex-determining region of the mouse Y chromosome is a member of a novel family of embryonically expressed genes. Nature 1990; 346: 245 5. Cortes D, Muller J, Skakkebaek NE. Proliferation of Sertoli cells during development of the human testis assessed by stereological methods. Int J Androl 1987; 10: 589 6. Russell LD, Peterson RN. Determination of the elongate spermatid–Sertoli cell ratio in various mammals. J Reprod Fertil 1984; 70: 635

TESTICULAR DYSGENESIS SYNDROME

7. Orth JM, Gunsalus GL, Lamperti AA. Evidence from Sertoli cell-depleted rats indicates that spermatid number in adults depends on numbers of Sertoli cells produced during perinatal development. Endocrinology 1988; 122: 787 8. Skakkebaek NE, et al. Carcinoma-in-situ of the testis: possible origin from gonocytes and precursor of all types of germ cell tumours except spermatocytoma. Int J Androl 1987; 10: 19 9. Rajpert-De Meyts E, et al. Developmental arrest of germ cells in the pathogenesis of germ cell neoplasia. APMIS 1998; 106: 198 10. Jacobsen GK, Nørgaard-Pedersen B. Placental alkaline phosphatase in testicular germ cell tumours and in carcinoma-in-situ of the testis. APMIS 1984; 92: 323 11. Jørgensen N, et al. Expression of immunohistochemical markers for testicular carcinoma in situ by normal human fetal germ cells. Lab Invest 1995; 72: 223 12. Looijenga LH, et al. POU5F1 (OCT3/4) identifies cells with pluripotent potential in human germ cell tumors. Cancer Res 2003; 63: 2244 13. Damjanov I, et al. Immunohistochemical localization of murine stage-specific embryonic antigens in human testicular germ cell tumors. Am J Pathol 1982; 108: 225 14. Rapley EA, et al. Localisation of susceptibility genes for familial testicular germ cell tumour. APMIS 2003; 111: 128 15. Giwercman A, von der Maase H, Skakkebaek NE. Epidemiological and clinical aspects of carcinoma in situ of the testis. Eur Urol 1993; 23: 104 16. Müller J, Skakkebaek NE. Testicular carcinoma in situ in children with androgen insensitivity (testicular feminisation) syndrome. Br Med J 1984; 288: 1419 17. Møller H. Clues to the aetiology of testicular germ cell tumours from descriptive epidemiology. Eur Urol 1993; 23: 8 18. Adami HO, et al. Testicular cancer in nine northern European countries. Int J Cancer 1994; 59: 33 19. Wilson JD, Lasnitzki I. Dihydrotestosterone formation in fetal tissues of the rabbit and rat. Endocrinology 1971; 89: 659 20. Baskin LS. Hypospadias and urethral development. J Urol 2000; 163: 951 21. Toppari J, Kaleva M. Maldescendus testis. Horm Res 1999; 51: 261 22. Wilson JD, George FW, Renfree MB. The endocrine role in mammalian sexual differentiation. Recent Prog Horm Res 1995; 50: 349 23. Skakkebaek NE. Possible carcinoma-in-situ of the testis. Lancet 1972; 2: 516

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24. Berthelsen JG, Skakkebaek NE. Gonadal function in men with testis cancer. Fertil Steril 1983; 39: 68 25. Berthelsen JG. Andrological aspects of testicular cancer. Int J Androl 1984; 7: 451 26. Petersen PM, et al. Gonadal function in men with testicular cancer. Semin Oncol 1998; 25: 224 27. Møller H, Skakkebaek NE. Risk of testicular cancer in subfertile men: case–control study. Br Med J 1999; 318: 559 28. Hoei-Hansen CE, et al. Histological evidence of testicular dysgenesis in contralateral biopsies from 218 patients with testicular germ cell cancer. J Pathol 2003; 200: 370 29. Sohval AR. Testicular dysgenesis as an etiologic factor in cryptorchidism. J Urol 1954; 72: 693 30. Huff DS, et al. Histologic maldevelopment of unilaterally cryptorchid testes and their descended partners. Eur J Pediatr 1993; 152 (Suppl 2): S11 31. Khuri FJ, Hardy BE, Churchill BM. Urologic anomalies associated with hypospadias. Urol Clin North Am 1981; 8: 565 32. Weidner IS, et al. Risk factors for cryptorchidism and hypospadias. J Urol 1999; 161: 1606 33. Sharpe RM. The ‘oestrogen hypothesis’ – where do we stand now? Int J Androl 2003; 26: 2 34. Pierik FH, et al. Maternal and paternal risk factors for cryptorchidism and hypospadias: a case–control study in newborn boys. Environ Health Perspect 2004; 112: 1570 35. Berkowitz GS, et al. Maternal and neonatal risk factors for cryptorchidism. Epidemiology 1995; 6: 127 36. Spitz MR, et al. Incidence and descriptive features of testicular cancer among United States whites, blacks, and Hispanics, 1973–1982. Cancer 1986; 58: 1785 37. McGlynn KA, et al. Increasing incidence of testicular germ cell tumors among black men in the United States. J Clin Oncol 2005; 23: 5757 38. Krausz C, Forti G, McElreavey K. The Y chromosome and male fertility and infertility. Int J Androl 2003; 26: 70 39. McElreavey K, Quintana-Murci L. Y chromosome haplogroups: a correlation with testicular dysgenesis syndrome? APMIS 2003; 111: 106 40. Richiardi L, et al. Testicular cancer incidence in eight northern European countries: secular and recent trends. Cancer Epidemiol Biomarkers Prev 2004; 13: 2157 41. Bray F, et al. Estimates of cancer incidence and mortality in Europe in 1995. Eur J Cancer 2002; 38: 99

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42. Møller H. Trends in incidence of testicular cancer and prostate cancer in Denmark. Hum Reprod 2001; 16: 1007 43. Boisen KA, et al. Difference in prevalence of congenital cryptorchidism in infants between two Nordic countries. Lancet 2004; 363: 1264 44. Boisen KA, et al. Hypospadias in a cohort of 1072 Danish newborn boys: prevalence and relationship to placental weight, anthropometrical measurements at birth, and reproductive hormone levels at three months of age. J Clin Endocrinol Metab 2005; 90: 4041 45. Jørgensen N, et al. East–West gradient in semen quality in the Nordic–Baltic area: a study of men from the general population in Denmark, Norway, Estonia and Finland. Hum Reprod 2002; 17: 2199 46. Punab M, et al. Regional differences in semen qualities in the Baltic region. Int J Androl 2002; 25: 243 47. Richthoff J, et al. Higher sperm counts in Southern Sweden compared with Denmark. Hum Reprod 2002; 17: 2468 48. Tsarev I, et al. Sperm concentration in Latvian military conscripts as compared with other countries in the Nordic–Baltic area. Int J Androl 2005; 28: 208 49. Carlsen E, et al. Evidence for decreasing quality of semen during past 50 years. Br Med J 1992; 305: 609 50. Vierula M, et al. High and unchanged sperm counts of Finnish men. Int J Androl 1996; 19: 11 51. Fisch H, et al. Semen analyses in 1,283 men from the United States over a 25-year period: no decline in quality. Fertil Steril 1996; 65: 1009 52. Paulsen CA, Berman NG, Wang C. Data from men in greater Seattle area reveals no downward trend in semen quality: further evidence that deterioration of semen quality is not geographically uniform. Fertil Steril 1996; 65: 1015 53. Handelsman DJ. Sperm output of healthy men in Australia: magnitude of bias due to self-selected volunteers. Hum Reprod 1997; 12: 2701 54. Swan SH, Elkin EP, Fenster L. The question of declining sperm density revisited: an analysis of 101 studies published 1934–1996. Environ Health Perspect 2000; 108: 961 55. Auger J, et al. Decline in semen quality among fertile men in Paris during the past 20 years. N Engl J Med 1995; 332: 281 56. Irvine S, et al. Evidence of deteriorating semen quality in the United Kingdom: birth cohort study in 577 men in Scotland over 11 years. Br Med J 1996; 312: 467

57. Van Waeleghem K, et al. Deterioration of sperm quality in young healthy Belgian men. Hum Reprod 1996; 11: 325 58. Iwamoto T, et al. Semen quality of 324 fertile Japanese men. Hum Reprod 2006; 21: 760 59. Swan SH, et al. Semen quality in relation to biomarkers of pesticide exposure. Environ Health Perspect 2003; 111: 1478 60. World Health Organization. WHO Laboratory Manual for the Examination of Human Semen and Sperm–Cervical Mucus Interaction, 4th edn. Cambridge: Cambridge University Press, 1999 61. Bonde JP, et al. Relation between semen quality and fertility: a population-based study of 430 first-pregnancy planners. Lancet 1998; 352: 1172 62. Slama R, et al. Time to pregnancy and semen parameters: a cross-sectional study among fertile couples from four European cities. Hum Reprod 2002; 17: 503 63. Guzick DS, et al. Sperm morphology, motility, and concentration in fertile and infertile men. N Engl J Med 2001; 345: 1388 64. Carlsen E, et al. Longitudinal changes in semen parameters in young Danish men from the Copenhagen area. Hum Reprod 2005; 20: 942 65. Ochsenkuhn R, De Kretser DM. The contributions of deficient androgen action in spermatogenic disorders. Int J Androl 2003; 26: 195 66. Jensen TK, et al. Association of in utero exposure to maternal smoking with reduced semen quality and testis size in adulthood: a cross-sectional study of 1,770 young men from the general population in five European countries. Am J Epidemiol 2004; 159: 49 67. Storgaard L, et al. Does smoking during pregnancy affect sons’ sperm counts? Epidemiology 2003; 14: 278 68. Jensen MS, et al. Lower sperm counts following prenatal tobacco exposure. Hum Reprod 2005; 20: 2559 69. Jensen TK, et al. Body mass index in relation to semen quality and reproductive hormones among 1,558 Danish men. Fertil Steril 2004; 82: 863 70. Rivas A, et al. Induction of reproductive tract developmental abnormalities in the male rat by lowering androgen production or action in combination with a low dose of diethylstilbestrol: evidence for importance of the androgen–estrogen balance. Endocrinology 2002; 143: 4797 71. Fisher JS, et al. Human ‘testicular dysgenesis syndrome’: a possible model using in-utero exposure of the rat to dibutyl phthalate. Hum Reprod 2003; 18: 1383

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72. Mahood I K, et al. Abnormal Leydig cell aggregation in the fetal testis of rats exposed to di (n-butyl) phthalate and its possible role in testicular dysgenesis. Endocrinology 2005; 146: 613 73. Calafat AM, et al. Automated solid phase extraction and quantitative analysis of human milk for 13 phthalate metabolites. J Chromatogr B Analyt Technol Biomed Life Sci 2004; 805: 49 74. Mortensen GK, et al. Determination of phthalate monoesters in human milk, consumer milk, and infant formula by tandem mass spectrometry (LCMS-MS). Anal Bioanal Chem 2005; 382: 1084 75. Sohoni P, Sumpter JP. Several environmental oestrogens are also anti-androgens. J Endocrinol 1998; 158: 327 76. Dolk H, et al. Risk of congenital anomalies near hazardous-waste landfill sites in Europe: the EUROHAZCON study. Lancet 1998; 352: 423 77. Garcia-Rodriguez J, et al. Exposure to pesticides and cryptorchidism: geographical evidence of a possible

78.

79.

80.

81.

82.

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association. Environ Health Perspect 1996; 104: 1090 Weidner IS, et al. Cryptorchidism and hypospadias in sons of gardeners and farmers. Environ Health Perspect 2004; 106: 793 Hardell L, et al. Concentrations of polychlorinated biphenyls in blood and the risk for testicular cancer. Int J Androl 2004; 27: 282 Hauser R, et al. The relationship between human semen parameters and environmental exposure to polychlorinated biphenyls and p,p′-DDE. Environ Health Perspect 2003; 111: 1505 Swan SH, et al. Decrease in anogenital distance among male infants with prenatal phthalate exposure. Environ Health Perspect 2005; 113: 1056 Rorth M, et al. Carcinoma in situ in the testis. Scand J Urol Nephrol Suppl 2000: 166

Section 2

Diagnosis of male infertility

8 Evaluation of the subfertile male Agnaldo P Cedenho

INTRODUCTION

needs to be studied with extreme care in search of the true cause of infertility, for once it is found, the physician will be able to decide what is the best treatment plan (with the best possible cost/benefit ratio) for the couple. Evidence-based andrology will thus ultimately allow childless couples to be spared the enormous stress associated with infertility. For many decades it has been conventional to define infertility as 1 year of failed attempts to conceive, and a couple should be investigated for infertility only after 1 year of regular sexual activity without the use of any contraceptives. This period of time was selected from epidemiological studies suggesting that around 85% of couples are able to achieve pregnancy within 1 year3. Therefore, after 1 year only 15% of couples will need infertility work-up. Even though the logic behind this rationale is evident, concessions need to be made considering the current situation and history of each partner in the infertile couple. If, for example, a woman is over 35 years old, or one of the partners has a clinical history that could lower his/her ability to conceive, this period of time may be shortened. On the other hand, since evaluation of the male partner in an infertile couple is simple, fast, inexpensive and usually non-invasive, it may be performed as soon as the infertile couple seeks medical assistance, or whenever the male partner decides to evaluate his

Based on the literature, it can be expected that 15–20% of couples within reproductive age will encounter difficulties in achieving a pregnancy, and medical attention will be required in order to start a family. Around 30% of these couples are infertile due to a significant isolated male factor, and associated male and female factors are present in an additional 20% of cases1. Therefore, an abnormal male factor is involved in about half of the couples seeking infertility treatment. Although male factor infertility plays such a dramatic role in a couple’s infertility, it has been left aside for decades. In fact, for a long time the man was examined solely using conventional semen analysis, without even an interview or a physical examination. Since the advent of intracytoplasmic sperm injection (ICSI) in 19922, this situation has become even worse. ICSI is without doubt a breakthrough in male infertility treatment, but since this technique overcomes virtually all natural barriers to fertilization, research on male factor infertility has lost its momentum, and both physicians and patients have shifted their focus from seeking and treating the cause of male infertility to achieving pregnancy only. Fortunately, as usually occurs in medicine, time and evidence puts everything back in its place. Perhaps more now than ever before, the subfertile man 117

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fertility status4. Evaluation of the male partner should be carried out following basic medical guidelines, which are: the patient’s history, physical examination, as well as all the laboratory and imaging resources available at the time. While the patient’s history and physical examination are of fundamental importance to all patients, imaging and laboratory techniques should be used as required. Many patients will require only two separate standard semen analyses, while others will need to go through many tests in order to find the cause of infertility. Each patient must be evaluated according to the individual situation.

PATIENT HISTORY Patient history and a careful physical examination may be of great help in evaluating the male partner of an infertile couple (Table 8.1) Although this chapter focuses on the infertile man, information regarding the female partner is not only useful, but also extremely relevant in deciding a treatment plan. We should not forget that as far as reproduction is concerned the couple must be seen as one functional unit, not as two separate individuals. It is therefore important to know the female history concerning menstrual cycle regularity, previous infections, pregnancies, abortions, abdominal surgery and possible risks related to sexually transmitted diseases (STDs). In many cases, coupling seminal analysis with the female partner’s examinations, such as pelvic ultrasound and hysterosalpingography, will allow the examiner to assess whether the couple can still achieve pregnancy through natural conception. On the other hand, the reproductive history is one of the most important areas to investigate in the infertile couple. It is very important to know how long the couple has been trying to achieve pregnancy without success, mainly because the longer is this period (> 7 years), the lower are the chances of natural conception and the graver are the factors involved. This is especially true when the female partner has normal menstrual cycles,

Table 8.1 Work-up sheet addressing anamnesis in a chronological fashion Conception

Natural conception or ART was needed

Prenatal

Drugs, pharmaceutical, environmental agents and endocrine disruptors

Childhood

Cryptorchidism, inguinal herniorrhaphy, bladder neck, pelvic or retroperitoneal surgery, testicular torsion

Puberty onset

Precocious or late, testicular trauma or torsion

Adolescence or young adult

Sexual behavior and STDs, viral or bacterial orchitis, recreational drugs, anabolic steroids, inguinal herniorrhaphy

Adult

Tricyclic antidepressives, antihypertensives, sulfasalazine, nitrofurantoin, cimetidine, chemotherapy, radiotherapy, retroperitoneal lymphadenectomy, inguinal herniorrhaphy, diabetes, multiple sclerosis, chronic respiratory diseases

Reproductive issues (female)

Menstrual cycle, infections, pregnancies, abortions, STDs, previous investigation and treatments

Reproductive issues (male)

Previous paternity, investigation, treatments, potency

Reproductive issues (couple)

Infertility duration, intercourse frequency and regularity, coital technique, knowledge about fertile period

ART, assisted reproductive technologies; STD, sexually transmitted disease

regular in frequency and with frequent sexual intercourse throughout the cycle. It is also relevant to ask how much the couple knows about the fertile period during the menstrual cycle. Recent data have shown that the best period for the sperm to penetrate the female reproductive tract is prior to ovulation. This period may last for up to 6 days, and immediately

EVALUATION OF THE SUBFERTILE MALE

after ovulation the cervical mucus becomes hostile to sperm, mostly due to a progestational effect5. Information regarding the sexual act itself is paramount to understanding the mechanisms underlying infertility. Lubricants used during sexual intercourse are usually spermicidal or determine lower sperm motility, and therefore it is necessary to know whether they are used. The most common lubricants used are K-YJelly, Lubrifax, Keri“ lotion or even saliva6–8. Previous fatherhood, albeit not a guarantee of current fertility, may reveal the reproductive potential of the male partner. Varicocele has been pointed out as the leading cause of secondary infertility9. Any previous investigation and treatment will help the evaluation to progress and spare the couple repeating examinations, thus saving time and money. A very productive and meaningful manner of evaluating patient history is through a work-up sheet that addresses anamnesis in a chronological fashion. This will give information regarding the male partner throughout different stages of his development. Prenatal exposure to drugs, pharmaceuticals or environmental agents should be assessed. Fetal exposure to diethylstilbestrol (DES) may lead to epididymal cysts, an increased incidence of cryptorchidism and altered semen variables in adult life10. Patients with hypospadias may present endogenous endocrine abnormalities, including altered testosterone biosynthesis11. On the other hand, it is important to emphasize the role that endocrine disruptors play in male infertility. These substances and their by-products, used in the phytopharmaceutical industry, may affect serum endocrine levels, or alter hormone action, production, release and/or elimination. These deleterious effects have been demonstrated in animal models, and an increased concern about their effects in humans has arisen due to a greater incidence of reproductive-tract abnormalities and decreased sperm concentration in many areas worldwide12. In the near future, even information regarding how the patient was conceived will be necessary. Since ICSI may allow children to carry the same

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genetic defects as their fathers, such as Y-chromosome microdeletions, infertility has ironically become an inheritable clinical condition. Cryptorchidism may affect 2–5% of born male children, and as such is an important cause of infertility13. Studies have demonstrated that 30% of children with unilateral cryptorchidism and 50% with bilateral cryptorchidism will present important semen alterations when adults. It is also noteworthy that, contrary to early indications, orchiopexy, even if performed while still very young, will not prevent future infertility14. Still during childhood, inguinal hernias, and their surgical correction, may play a role in infertility. Inguinal herniorrhaphy is the leading cause of iatrogenic obstruction of the vas deferens and testicular atrophy due to impaired blood supply to the testis15,16. An estimated 0.8–2% of inguinal herniorrhaphies performed in children lead to iatrogenic lesions of the deferent ducts, while in adults that risk decreases to about 0.3%17,18. Furthermore, there is no doubt that the actual number of iatrogenic lesions to the vas deferens is larger, but since the surgical procedure is usually unilateral, fertility is not always affected. Ejaculatory disturbances may be caused by surgery performed during childhood on the bladder neck, in the pelvis or in the retroperitoneum. In the early 1960s, many children presenting with urethral defects were submitted to a surgical procedure known as YV plastic repair of the bladder neck. This surgery causes serious lesions to the internal sphincter, causing bladder-neck closure defects. In adult life these patients present with a decreased ejaculate volume (< 1 ml), and retrograde ejaculation. This diagnosis may be confirmed by finding sperm in the urine after ejaculation. Puberty usually occurs between the ages of 11 and 12 years in boys. If puberty onset is precocious, this may indicate an adrenogenital syndrome. On the other hand, if puberty is delayed, it may be secondary to an endocrinopathy, such as in Klinefelter’s syndrome or idiopathic hypogonadism.

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Testicular torsion may occur in the newborn, infant or adolescent, and may lead to testicular atrophy. An estimated 30–40% of men with a history of testicular torsion have difficulty in achieving parenthood, due to the alterations in semen variables19,20. Although in the past changes of spermatogenesis were attributed to a possible rupture in the blood–testis barrier, testicular biopsies in contralateral testes have shown that these also present with histological alterations, supporting the hypothesis that testes prone to torsion, as well as their contralateral counterpart, already demonstrate defects in spermatogenesis21. Between adolescence and adult life, information regarding sexual behavior and risks related to STDs is very important. Even though its importance has declined, urethritis is still an important source of infection to the prostate, epididymides and testes. The most common urethritis-causing agents are: Neisseria gonorrhoeae, Chlamydia trachomatis, Ureaplasma urealyticum and Trichomonas vaginalis. In the United States, C. trachomatis is the principal agent in causing non-gonococcic urethritis and acute epididymitis, and 10–25% of men are asymptomatic, sometimes presenting with an increase in seminal leukocytes22. Viral orchitis may also impair testicular function, especially if the onset is postpubertal. Mumps may cause unilateral orchitis in 30% of male patients and bilateral orchitis in 10%, and these patients will possess a decrease in testicular volume and consistency23. An important reminder is that a prolonged fever on its own can be a source of damage to spermatogenesis. Therefore, these effects will not be observed immediately, since the duration of the spermatogenic cycle is 74 days in men, a period during which type B spermatogonia will differentiate into mature sperm, in addition to 15 days of sperm transport through the excretory system until they are ready for ejaculation. Thus, if damage to the testis from fever or medication is suspected, seminal analysis should be performed after 90 days.

Substance abuse has been linked to male infertility in many studies, and it is well documented that alcohol24,25, tobacco26, marijuana27, cocaine28,29 and anabolic steroids30,31 may also cause testicular dysfunction. Alcohol has been shown to decrease serum testosterone levels, and this is due to its effects on three different levels: the hypothalamus, the pituitary gland and the testicular Leydig cells. While it directly alters Leydig cell function, and thus leads to the observed lower testosterone levels, alcohol may also have a negative impact on hypothalamic hormone production and the pituitary production, release and function of luteinizing and follicle stimulating hormones24,25. Tobacco, on the other hand, may cause a number of alterations, such as testicular atrophy, altered sperm morphology, low sperm motility, decreased semen volume, impaired spermatogenesis, poor acrosome reaction and sperm-penetrating ability, increased amounts of oxidative DNA damage, a higher risk for chromosome 13 aneuploidies and elevated serum prolactin and estradiol levels26. Marijuana and cocaine have both been shown to interfere with spermatogenesis, decreasing sperm concentration and motility and increasing the number of sperm with altered morphology27,28, while high doses of cocaine may cause erectile dysfunction29. Finally, exogenous testosterone and its metabolite, estrogen, lead to the suppression of gonadotropin releasing hormone (GnRH) production in the hypothalamus. This leads to a decreased release of luteinizing hormone (LH) from the pituitary gland, and thus to lower testicular testosterone production in the Leydig cells31. After ceasing use of these substances, spermatogenesis is expected to be back to normal within 3–6 months. Although uncommon, the pituitary suppression caused by steroidal drugs may be irreversible32. There are medications that may interfere with spermatogenesis, affecting quality and quantity of the ejaculate. Antidepressive therapy may increase blood prolactin levels, which in turn will decrease

EVALUATION OF THE SUBFERTILE MALE

the production of gonadotropins33. Calcium channel blocker antihypertensives (nifedipine, diltiazem) may block the acrosome reaction and prevent sperm–egg binding34,35, while alpha-blocker antihypertensives (prazosin, terazosin, phenoxybenzamine) may lead to retrograde ejaculation or even aspermia36. Other drugs are known to impair spermatogenesis, depending on the dosage and length of treatment. Some examples are sulfasalazine37,38 and cimetidine39. Testicular cancer, Hodgkin’s disease and leukemia represent three of the most frequent oncological diseases in young adult males. Their incidence is highest in the age group 15–35 years. A growing number of young men are treated successfully for cancer by chemotherapy and radiotherapy. Testicular cancer, for example, a major concern in male infertility, is currently treated by orchiectomy associated with chemotherapy, radiotherapy and retroperitoneal lymphadenectomy, with survival rates reaching upwards of 90%40. The benefits of these therapies come at a price, and this may be temporary or permanent infertility41. Testicular damage caused by cytotoxic drugs was first described in humans in 1948, when azoospermia was reported in men following treatment with nitrogen mustard42. Many other drugs have been shown to be gonadotoxic, and the agents most commonly implicated are: alkylating agents (cyclophosphamide, chlorambucil, busulfan, procarbazine, mustine, melphalan), antimetabolites (cytarabine, 5-fluorouracil, methotrexate), vinca alkaloids (vinblastine, vincristine), cisplatin and analogs, and topoisomeraseinteractive agents (bleomycin, doxorubicin, danorubicin, actinomycin)43. Although efforts have been made to modify protocols in order to minimize effects on fertility, the chances of fathering children after treatment remain difficult to predict44. Alkylating agents are known to be the drugs most deleterious to spermatogenesis, and they cause a cumulative effect. When a dose of > 400 mg/kg of alkylating agents is used, 30% of

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prepubertal boys and 70–95% of adult men will present with gonadal dysfunction45,46. Radiotherapy, on the other hand, may cause permanent azoospermia if doses of > 400 cGy are used, while a dose of > 300 cGy may cause various degrees of oligozoospermia47. These deleterious effects of chemo- and radiotherapy on spermatogenesis may last for up to 4 or 5 years, and therefore seminal analyses performed before this period of time should be considered inconclusive. Statistically, 50% of patients with a history of testicular cancer are oligozoospermic, and 7–10% azoospermic before receiving any treatment48. Post-therapeutic spermatogenic output will depend on the type of chemo- or radiotherapy utilized. Cryopreservation of sperm has provided hope for fertility preservation in cancer patients. These men should be referred to a licensed spermbanking unit as soon as possible, to collect one or various samples for freezing. Sperm banking is currently the only proven method of preserving fertility in cancer patients, although hormonal manipulation to enhance spermatogenic recovery and banking of testicular germ cells are both possibilities for the future43. Besides the negative effects of chemo- and radiotherapy on testicular function, retroperitoneal lymphadenectomy may cause ejaculatory dysfunction. Most of these patients will present with aspermia due to interruption of the sympathetic nodal nervous chain or its peripheral nerves, such as the sacral plexus or the hypogastric nerves49. Recently, nerve-sparing techniques have been used more, and these side-effects have become less common50. Patients presenting with postsurgical aspermia or, more rarely, retrograde ejaculation may revert to anterograde ejaculation after treatment with sympathomimetic drugs (e.g. ephedrine sulfate)51. If the pharmaceutical approach fails, semen can simply be retrieved from the urine and, after pH and osmolarity control, be used for intrauterine insemination (IUI) or ICSI. Nevertheless, the best way to preserve fertility in a male patient with testicular cancer during reproductive age is through

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gamete cryopreservation prior to the oncological treatment, especially with the advent of ICSI, which allows the use of very few sperm to achieve fertilization. Finally, systemic diseases may also affect the male reproductive tract. Diabetes and multiple sclerosis may both cause ejaculatory and sexual dysfunction, for example. Respiratory diseases associated with infertility may be caused by immotile cilia syndrome, or Kartegener’s syndrome, in which sperm concentration is normal but they are immotile due to defects in the flagellum.

PHYSICAL EXAMINATION A general physical examination should consider weight, height and arm span. Careful observation of all the systems may reveal important signs contributing to male-factor infertility diagnosis. This is especially true since altered function in many organs may alter reproductive potential in the man. As an example, an infertile patient without any specific complaint presenting with inadequate virilization, such as sparse facial and pubic hair and gynecomastia, may have hypogonadism or Klinefelter’s syndrome. On the other hand, inadequate virilization and anosmia are associated with Kallmann’s syndrome. Although chronic diseases are not usually found when evaluating a patient for infertility, early or mild alterations in adrenal function, chronic alcoholism or diabetes may be detected by careful physical examination. However, the genital physical examination, performed under ideal temperature (> 23oC) and light conditions, will provide the most important information regarding the pathogenesis of infertility in the male partner. The examination initiates with the patient in the upright position. This will allow better evaluation of the penis, scrotal size, testicular position, symmetry of testicular structures and, no less important, the venous return condition in the

pampiniform plexus. Regarding the penis, insertion of the urinary meatus is the most important aspect, since hypospadias renders the patient unable to place the ejaculate within the vaginal vault. A small scrotum or the scrotum of an obese man is more difficult to palpate, and therefore these patients may require scrotal ultrasonography. When examining the testes, size, consistency and regularity should be recorded. Patients with normal seminal analysis usually have testicles of 4.5 cm in length by 2.5 cm in height, with a minimum volume of 15 ml, as assessed by the Prader orchidometer. Not surprisingly, testicular volume and consistency usually predict seminal analysis results, especially taking into account that 85% of testicular volume is represented by the seminiferous epithelium. On the other hand, patients with small testes tend to present with varying degrees of oligozoospermia or even with azoospermia. If during the prepubertal phase the boy does not undergo normal gonadal development, in adult life he will most likely present with small (< 8 ml) and hardened testes. Such is true in Klinefelter’s syndrome. However, if the gonads develop normally but suffer injuries, such as in viral or bacterial orchitis, they show decreased size and consistency. Following examination of the testes, the epididymides should be evaluated using one hand to hold the testis while the other gently palpates the epididymal head, body and cauda between the thumb and the index finger. Epididymal volume, consistency, regularity, cysts and distance between the testis and the epididymis should be noted at this time. The further is the epididymis from the testis the more prone it is to abnormalities, while epididymal volume reflects testicular production and effusion. It is quite common to observe cysts in the epididymal head, but they are usually smaller than 0.7 cm and have no clinical meaning. However, when these cysts are larger or more numerous they may obstruct the epididymis and block sperm passage.

EVALUATION OF THE SUBFERTILE MALE

Moving on through the male reproductive tract, the deferent ducts should be evaluated. These are firm, cylindrical structures measuring 3 mm in diameter, and are easily distinguished from the other structures in the spermatic cord. Under ideal room-temperature conditions and with a cooperative patient the deferent ducts are always identified. For that reason, deferent duct agenesis is, in most cases, diagnosed solely through the physical examination, and ancillary examinations and exploratory surgery are not necessary. Various degrees of epididymal malformation, as well as agenesis or hypoplasia of the seminal vesicles, usually accompanies uni- or bilateral absence of the vas deferens. If the physical examination shows a thickening or hardening of the vas deferens, this may be a sign of previous infection, usually caused by STDs. Vasectomized patients present with dilated and painful epididymides. The last structure analyzed in the physical examination is the pampiniform plexus of the spermatic cord, observing possible asymmetries, bulges or growths. With the patient in the upright position, testicular volumes are measured and expected to be symmetrical. A grade II or III varicocele, usually on the left side, will reduce the testis volume and shift its axis from vertical to horizontal. Admitting that varicocele causes alterations in spermatogenesis, seminiferous tubule diameter will decrease, as well as testicular volume. It is common to find testicular asymmetry in unilateral varicocele patients, and the ipsilateral testis will be at least 2 ml smaller in volume than its contralateral counterpart. For diagnosis of a grade I varicocele, the patient will have to perform a Valsalva maneuver. Careful examination before and during a Valsalva maneuver will allow the examiner to palpate any engorgement of the pampiniform plexus. Subclinical varicoceles are not diagnosed by a physical examination, and their clinical meaning is currently questioned. The spermatic cord should be examined up to the point at which it exits the scrotum, and any abnormality, such as a spermatic cord cyst or inguinal–scrotal hernia, should be observed.

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The history and physical examination, with especial attention paid to the genitalia, are essential components not only in diagnosing male factor infertility but also in determining the management and prognosis for the infertile couple. Although seminal analysis is still the most important examination in male factor evaluation, it does not possess the necessary standardization, due to the lack of proper guidance given to the patient when ordering the examination and the lack of protocol and quality assurance among different laboratories. As a result, comparisons from different laboratories are very difficult. It is important to keep in mind that seminal analysis will not determine whether a man is fertile or not, especially because fertility is a couple phenomenon, of which pregnancy is the ultimate proof. Detailed descriptions of the current methodologies and interpretation of semen analysis are discussed in other chapters. Although semen analysis initiates the investigation of the infertile man, it cannot provide all the answers to questions regarding his fertility potential. It is never enough to repeat that semen analysis alone will not allow determination of the patient’s true fertility potential, but the average results from separate seminal analyses will allow the physician to estimate this potential. With these considerations in mind, in our institution we group patients into three categories (Table 8.2), according to their potential for natural conception. This table should serve only as a starting point for discussing the patient’s condition, and, as mentioned previously, the cut-off rates shown are still widely debatable. According to the World Health Organization, the reference value for semen volume is 2.0 ml52. If the patient produces no semen at all after an orgasm, he has aspermia. This may be due to clinical issues, such as bilateral sympathectomy, bilateral retroperitoneal lymphadenectomy, antihypertensive drugs which block the sympathetic tone, transurethral or open surgical resections of the bladder neck or prostate, extensive pelvic surgery and diabetic neuropathy.

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Table 8.2 Semen classification based on an average between two samples Potential for natural conception Variables

High

Moderate

Low

Volume (ml)

1–2

Concentration (× 106/ml)

> 20

10–20

< 10

Motility (% motile)

> 50

40–50

< 40

Morphology (% normal)

> 14* > 30**

4–14* 20–30**

< 4* < 20**

Classified according to Kruger strict criteria, 1986; classified according to World Health Organization, 1999

Hypospermia without spermatozoa in semen with a pH of less than 7.4 could be due to ejaculatory duct obstruction or congenital absence of seminal vesicles. Hypospermia with spermatozoa in the ejaculate with a pH of less than 7.4 could be due to obstruction of the seminal vesicle opening by a mucus-like plug; this obstruction may dissolve spontaneously53.

MALE REPRODUCTIVE ENDOCRINOLOGY From the clinical point of view, the most important hormones related to male fertility are follicle stimulating hormone (FSH), luteinizing hormone (LH) and testosterone. FSH acts primarily on Sertoli cells, stimulating the production of androgenbinding protein (ABP), which in turn binds to testosterone and intensifies its action on the seminiferous tubules54. Sertoli cells also produce inhibins A and B, which inhibit pituitary secretion of FSH, and even if the testis presents only spermatogonia, inhibin production is sufficient to decrease FSH levels to normal. FSH levels are therefore limited in predicting spermatogenic integrity55. LH acts on Leydig (or interstitial)

cells, where it stimulates the synthesis of testosterone. Only 2% of circulating testosterone is in the unbound form (free testosterone), and thus capable of producing its effects. Around 30% of circulating testosterone is bound to a specific globulin – sex hormone-binding globulin – and 68% is bound to albumin and other non-specific proteins56. Testosterone will, in the same manner as FSH, stimulate Sertoli cell function and therefore promote spermatogenesis. Testosterone is also converted into dihydrotestosterone in peripheral tissue, where it is responsible for the manifestation of secondary male sex characteristics57. Prolactin is secreted by the pituitary gland, and its production is inhibited by dopamine and stimulated by thyroid stimulating hormone (TSH). Although prolactin does not exert a direct action on spermatogenesis, chronic hyperprolactinemia is known to alter GnRH action, leading to altered secretion of FSH and LH and, consequently, decreased libido, sexual dysfunction, gynecomastia and alterations of spermatogenesis58. The male endocrine profile (FSH, LH and testosterone) is not necessary in patients with a normal seminal analysis. Patients with a sperm concentration as low as 10 × 106 cells/ml have been shown to be able to achieve paternity by natural conception59. On the other hand, patients with a sperm concentration of fewer than 5 × 106 cells/ml demonstrate significantly lower fertility rates60. A hormonal profile is therefore useful in severely oligozoospermic and azoospermic patients, as shown in Table 8.3. If we consider only circulating FSH levels, a few practical conclusions related to oligozoospermia or azoospermia may be reached: • Normal FSH: the alteration is either posttesticular (obstructive) or testicular (normogonadotropic hypogonadism). If post-testicular, hormonal treatment is unnecessary, and fertilization may be achieved through vasectomy reversal or ICSI (e.g. congenital bilateral absence of the vas deferens) using epididymal or testicular (TESE) aspiration61. If the

EVALUATION OF THE SUBFERTILE MALE

Table 8.3

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Association between serum hormone levels and origin of oligozoospermia or azoospermia

Type of oligozoospermia or azoospermia

Serum hormone levels

Post-testicular (obstructive) Vasectomy Congenital bilateral absence of the vasa deferentia

FSH, LH and testosterone usually normal

Pre-testicular (usually from hypothalamic or hypophyseal disorders, also known as hypogonadotropic hypogonadism) Tumors Hyperprolactinemia Kallmann’s syndrome

Low FSH, usually low LH and testosterone

Testicular (hypergonadotropic hypogonadism) Genetic syndromes (Klinefelter’s, myotonic dystrophy) Embryological malformations (cryptorchidism) Cytotoxic drugs (chemotherapy) Sequelae from viral diseases (mumps) Testicular (normogonadotropic hypogonadism) Androgen resistance Sertoli cel only syndrome and maturation arrest Y-chromosome microdeletions

Elevated FSH, variable LH and testosterone

Normal FSH, elevated LH and testosterone Normal FSH, LH, and testosterone Normal FSH, LH, and testosterone

FSH, follicle stimulating hormone; LH, luteinizing hormone

alteration is testicular, thus signifying androgen resistance, hormonal treatments could be beneficial to the patient. If this is not the case, sperm could be retrieved by masturbation (if oligozoospermic) or TESE (if azoospermic) for ICSI62. • Low FSH: the alteration may be either hypothalamic or hypophyseal, and treatment involves correcting these primary alterations. Sometimes, simultaneous hormonal treatment is necessary, such as in Kallmann’s syndrome. • High FSH: anamnesis and karyotyping may both help to define diagnosis in these patients, and treatment will depend on the etiology and presence of sperm in the ejaculate or in the testis63. If viable sperm are found, fertilization may be achieved through ICSI. If not, the couple may have to use donor semen or adoption as an option for constituting a family.

IMAGING THE REPRODUCTIVE TRACT There are several imaging resources that may be used to investigate the male reproductive tract for abnormalities, but the most frequently used are ultrasonography64 and nuclear magnetic resonance (NMR)65. Computerized tomography has been less and less indicated in clinical practice, due mainly to the fact that, besides using ionizing radiation, it does not render superior pelvic images when compared with transrectal ultrasound (TRUS) or magnetic resonance imaging (MRI). Nowadays deferentography is hardly used, mostly because there is a risk of iatrogenic deferent lesions at the puncture site. However, when there is doubt regarding vas deferens injury from previous hernia repair, deferentography is the imaging method of choice for confirming the clinical suspicion.

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Scrotum With the patient in a standing position in a warm room, an attentive physician is able to inspect and examine all the structures inside the scrotum. These include testicular and epididymal volume, consistency and regularity, the presence or absence of the vasa deferentia as well as their diameters and clinical varicocele. Since the role of subclinical varicocele is currently rather controversial66, and physical examination of the scrotum provides most of the information we need, imaging resources are not used very often to evaluate scrotal content. As an exception, scrotal ultrasonography may be helpful when evaluating obese patients or patients with a short scrotum.

vesiculography can be performed70. A large number of spermatozoa in the seminal vesicle fluid reinforce the diagnosis. While complete ejaculatory duct obstruction is relatively easy to diagnose and is accepted by everyone, partial duct obstruction is suggested by some, and is not as easy to demonstrate. Usually, when a patient is oligozoospermic and/or asthenozoospermic with a lower ejaculate volume, without any other clinical or laboratory finding, he is investigated for partial ejaculatory duct obstruction. Aspiration puncture of the seminal vesicles could be important in these patients, especially if performed immediately following ejaculation, because the partial obstruction will lead to impaired efflux from the seminal vesicles, and a large number of sperm may be found in the aspirate.

Ductal obstruction Although complete obstruction of the deferent ducts is very rare, it should be investigated because it is treatable67. A good imaging resource in this situation is high-frequency transrectal ultrasonography (TRUS), because it produces excellent images of the ejaculatory ducts, seminal vesicles and prostate68. It is also considered a simple, readily available and inexpensive examination. Another imaging approach is MRI, which can be performed either with or without a rectal probe. Although it offers very good spatial reconstitution of the necessary structures, it is not readily accessible, and costs will be significantly increased. However, in contrast to TRUS, MRI does not depend on examiner skill69. Patients with a normal scrotum examination who present with a low volume of ejaculate (< 1 ml) and seminal fluid devoid of fructose and coagulation might have complete ejaculatory duct obstruction. If submitted to TRUS, they may exhibit dilated ejaculatory ducts and/or seminal vesicles (greater than 1.5 cm in anteroposterior diameter). But it is important to keep in mind that normal vesicle size does not necessarily rule out the possibility of ductal obstruction. Under TRUS guidance, seminal vesicle aspiration and

Pituitary gland In male infertility, the most common indication for carrying out computerized tomography or MRI of the brain is in imaging the pituitary gland for diagnosis of hypogonadotropic hypogonadism65. Even if very unusual in an infertilityclinic setting, hypogonadism associated with gonadotropic insufficiency deserves special attention, as it is one of the few alterations in male infertility with specific and effective clinical treatment.

VARICOCELE Varicocele is defined as an abnormal increase in scrotal volume due to dilated veins in the pampiniform plexus. Although it is present in 15–25% of the male population, its prevalence can reach 40% in infertile men71,72. Most patients are asymptomatic, but some may present with testicular pain which increases following physical activities or long periods in the upright position. However, the pain is relieved upon adopting the supine position, which explains why patients do not usually refer to pain in the morning.

EVALUATION OF THE SUBFERTILE MALE

Diagnosis is performed through careful physical examination in a warm room (> 23oC), with the patient in the upright position. If there is an observable or palpable dilatation in the pampiniform plexus before or during a Valsalva maneuver, diagnosis is confirmed, and the varicocele classified as grade I, II or III, according to the intensity of the dilatation: • Grade I varicoceles are visible with difficulty, but easily palpable during a Valsalva maneuver; • Grade II varicoceles are visible, and there is significant venous gorging during the Valsalva maneuver; • Grade III varicoceles are easily visible, with great reflux during the Valsalva maneuver. There is enough evidence in the literature to demonstrate that varicocele can cause macroscopic, microscopic and functional alterations to the testes73,74. Varicocele usually develops earlier and more intensely on the left side, because venous return is more difficult due to anatomical peculiarities in the internal spermatic drainage system on this side75,76. Macroscopic alterations are evident in adolescents, because these patients present with a delay in development of the left testis. This delay will eventually lead to testicular asymmetry, a difference in volume of more than 2 ml between the testes in the adult77,78. Histologically, patients with varicocele demonstrate a loss of maturational stratification, characterized by: loss of desmosomes, adluminal compartment structural disorganization, maturation arrest in the different stages of spermatogenesis, early release of spermatids into the lumen and, as a consequence, thinning of the seminiferous epithelium and increase of the tubular lumen79,80. As far as testicular function is concerned, the consequences of venous ectasia may be observed in two compartments: interstitial and intratubular. The World Health Organization (WHO), through a multicentric study comparing young patients with and without varicocele, observed that varicocele patients possess lower blood

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testosterone levels, characterizing impaired steroidogenesis and Leydig cell dysfunction. Alterations in the seminiferous tubules cause changes in seminal variables and lead to a decrease in sperm concentration, motility and normal morphology81. There is also evidence showing that sperm from patients with varicocele possess a lower ability to bind tightly to the human oocyte zona pellucida82. The negative effects of varicocele on the testes have been shown over the past few decades, either through clinical83,84 or experimental studies85. Although many theories have been proposed and much has been hypothesized, it is not known how venous reflux and ectasia lead to testicular malfunction. Many studies regarding varicocele and infertility evaluate variables between men with and without varicocele or assess pre- and postvaricocelectomy data, but a definite explanation for the negative impact of varicocele on gametogenesis or for its bilateral effects has yet to be found86.

Etiology Testicular blood flow and hyperthermia

The involvement of testicular blood flow in varicocele etiopathogeny is highly concordant with studies related to hyperthermia, but many controversies have yet to be explained87,88. Some groups have shown that this increase in blood flow is present on both sides, even in unilateral varicoceles89. Even though a change in testicular blood flow direction associated with varicocele has not been defined, it is very important to recognize that an increase in blood flow is most compatible with hyperthermia, and that contralateral organs may respond in a similar fashion to their ipsilateral counterparts when an injury is present, due to hormonal and neural mechanisms. Testicular hyperthermia is considered the most important mechanism leading to the secondary alterations associated with varicocele90. The scrotum is maintained at a lower temperature than

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body temperature owing to five important anatomical traits: (1) dartos muscle, (2) cremaster muscle, (3) countercurrent heat-exchange mechanism, (4) the absence of adipose tissue and (5) the presence of many sweat glands. Two systems play a main role in thermal regulation. The scrotal system, with the dartos and cremaster muscles, assists the countercurrent heat-exchange mechanism. This in turn allows heat exchange from the arterial to the venous system, thus maintaining thermal homeostasis90. Varicocele impairs this heatexchange mechanism, and therefore hinders the cooling of arterial blood before it enters the testis. This alteration in blood flow prevents the testis from maintaining a lower temperature. An increase in testicular temperature may have a direct effect on spermatogenic germ cells, altering metabolism, Sertoli cell function, DNA synthesis, apoptosis rates and nutrient content and oxygen tension in the testicular environment, as well as decreasing enzymatic activity and leading to vascular alterations due to an increase in arteriovenous shunting91,92. The higher testicular temperature associated with androgenic suppression will also act concurrently, altering different stages of spermatogenesis, generally lowering the sperm concentration93.

Spermatogenesis and apoptosis Spermatogenesis is a continuous proliferative process that leads to the production of millions of sperm each day. Apoptosis, or programmed cell death, is present in both physiological and pathological situations, and will determine, during gametogenesis, the sperm concentration in fertile and infertile men. Apoptosis is associated with nuclear DNA fragmentation94, and is present throughout spermatogenesis, occurring in spermatogonia, spermatocytes and spermatids95. Since the process of apoptosis has been extensively studied and documented96–102, and although it is known that heat stress, androgen deprival and accumulation of toxic substances in the testes all contribute to increase, apoptosis

rates, more studies regarding the molecular mechanisms underlying varicocele-induced activation of apoptosis need to be done103. Apoptosis-induced DNA fragmentation can currently be evaluated through many different techniques, of which the TUNEL (terminal deoxynucleotide transferase-mediated dUTP nick-end labeling) and the Comet (or single cell gel electrophoresis) assays are noteworthy. In a recent study, DNA fragmentation rates were significantly increased in adolescents with grades II and III bilateral varicocele104.

Sperm motility Many men with varicocele present lower sperm motility, and this may or may not be accompanied by alterations in other sperm variables105. Most studies have focused on three basic causes of this lower sperm motility: an increased concentration of reactive oxygen species (ROS), the presence of antisperm antibodies and deficient mitochondrial activity106. ROS concentration in sperm is inversely related to motility107. Although ROS are normally present, and even necessary at low concentrations, excessive ROS are present during leukospermia or an increased presence of abnormal sperm, such as sperm with cytoplasmic droplets108. Men with varicocele present a higher concentration of ROS and a lower antioxidant capability109. This is demonstrated by the fact that these men demonstrate a defect in mitochondrial oxidative phosphorylation, with a low concentration of the mitochondrial coenzyme Q10, an important antioxidant110, and a deficiency in superoxide dismutase (SOD) and catalase111.

Sperm morphology Although classical alterations in sperm morphology associated with varicocele are an increase in fusiform and amorphous cells112, recent data assessed through strict criteria demonstrate that there is a decrease in normal sperm

EVALUATION OF THE SUBFERTILE MALE

morphology113. Specific studies evaluating sperm donors exposed to cadmium, shown to cause stress to the testes, demonstrated an increase in the expression of heat shock protein (HSP), which, among other characteristics, possesses actin-like sequences. Since it is not yet known whether HSPs act as protectors of actin or inhibit its polymerization114, and since patients with varicocele exposed to environmental agents possess higher cadmium concentrations, future studies may help us to understand how HSPs participate in sperm morphology determination. Another important finding is an increase in cells with midpiece cytoplasmic droplets, which lead to lower sperm motility in these patients with varicocele115. It is also important to assess the acrosome reaction in patients with varicocele, along with functional tests that evaluate the acrosome and its ability to bind to the zona pellucida. A study of adolescents with varicocele found that sperm from these patients possessed decreased binding capacity to the zona pellucida (hemizona assay, HZA), when compared with adolescents without varicocele82.

Acrosome reaction Alterations of sperm function seem to be more relevant than morphology and concentration in varicocele patients, and this involves the acrosome reaction and zona pellucida binding116. Sperm from patients with varicocele demonstrate an altered calcium influx mechanism117. Cofactors such as metals may exacerbate this alteration, and there is a wide variety of individual response, ranging from no alteration to infertility. Since this variability may be explained by qualitative and quantitative differences in protein expression, studies have set out to find candidate genes, to evaluate susceptibility for this defect118. The acrosome reaction is calcium-dependent, and few motile sperm are able to complete this exocytosis119. Following cholesterol efflux there is an influx of calcium ions, which will stimulate

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mannose receptor externalization and initiate exocytosis through myosin activity117. Calcium influx is controlled by voltage-dependent channels120, and it has been demonstrated that men with varicocele exhibit a deletion of amino acids in the calcium channel pore, thus providing a genetic cause for infertility. These channels may be altered when environmental agents are present, since they may also transport zinc, cadmium, cobalt, nickel, lead and aluminum121. Since the testicular cadmium concentration is higher and the zinc concentration lower in men with varicocele118, it has been suggested that cadmium may negatively affect calcium channels. It has not yet been defined whether varicocelectomy or zinc supplementation will reverse the effects of cadmium on these channels106. In spite of the enormous progress that has been achieved related to varicocele and its consequences on spermatogenesis, many doubts still remain. Prospectively designed studies, which are currently scarce, would not only help us to understand better the intrinsic mechanisms through which varicocele affects male fertility, but also shed light on the present uncertainties regarding treatment.

AZOOSPERMIA Defined as the complete absence of sperm in the seminal fluid after centrifugation, azoospermia represents a very important topic in male infertility, and, as such, deserves special attention. It is present in 1% of all men and in approximately 15% of infertile men122,123. The first issue regarding azoospermia is to be certain that we really are dealing with azoospermia and not severe oligozoospermia. The distinction between these two entities is not only a semantic issue but rather fundamental, since the presence of just a few spermatozoa could represent the difference between being a genetic father or not. This has become particularly true since the introduction of ICSI in 1992. For this reason, the WHO guideline recommends that if no

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spermatozoa are found in three aliquots of 10 µl of semen, the whole specimen should be centrifuged at 3000 g for 15 min and the resulting pellet examined thoroughly52. Moreover, it is not unusual to detect sperm in the specimen of a patient initially considered azoospermic124. Therefore, azoospermia should not be definitely assumed unless two separate samples are scrutinized in this way. Azoospermia may be due to a variety of conditions, and a history, physical examination, hormonal profile, genetic and imaging resources will be necessary not only to establish the cause but also to direct the couple towards the best treatment option suitable. Some causes are potentially correctable; other conditions are irreversible but still possibly treatable by assisted reproductive techniques using the husband’s semen; and, finally, some causes are irreversible and not amenable to any form of treatment, demanding donor semen or adoption in order to constitute a family. This section discusses the evaluation of some specific conditions associated with azoospermia.

from a therapeutic perspective they are similar to patients with primary testicular failure, and, as such, are not clinically treatable. Finally, patients with low FSH, LH and testosterone have secondary hypogonadism, and represent one of the very few occasions where specific therapy may be effective. However, patients with congenital or acquired hypogonadotropic hypogonadism are rarely seen in an infertility clinic, but when they do present they should be tested for deficiencies of other pituitary hormones (thyroid-stimulating, adrenocorticotropic and growth hormones)126. Patients with an altered gonadotropin profile and anosmia or hyposmia are candidates for Kallmann’s syndrome. A careful neurological examination, including visual field testing, serum prolactin measurements and radiological images of the pituitary fossa may reveal a pituitary adenoma. It is especially noteworthy that, although it is unusual for infertility due to hyperprolactinemia to occur in men without impotence and hypoandrogenization, hyperprolactemia does occur without any detectable hypothalamic or sellar alteration.

Azoospermia with small testicles

Congenital absence of vasa deferentia

Azoospermia associated with bilateral small testicles may be caused by either primary or secondary testicular failure. The differentiation between these two very distinct clinical situations is feasible using the initial results of the endocrine tests. Patients who sustain elevated FSH and LH and low testosterone levels have primary testicular insufficiency in both Leydig and germ-cell compartments. Elevated gonadotropins distinguish primary testicular failure from hypothalamic– pituitary diseases. Klinefelter’s syndrome and its variants represent a typical example of primary testicular failure. These alterations, confirmed by karyotyping, account for 14% of these cases of azoospermia125. On the other hand, some patients might present azoospermia with elevated FSH but normal LH and testosterone. Although they do not exhibit total panhypogonadal dysfunction,

Considering that the scrotal contents are very easily reached and knowing that the vas deferens is a fairly solid structure, 3 mm in diameter, the diagnosis of unilateral or bilateral vasal agenesis is possible through physical examination. Ancillary examinations or even surgical exploration is not necessary to confirm the diagnosis, but may be useful for seeking associated abnormalities. Approximately 25% of men with unilateral vasal agenesis and 10% of men with congenital bilateral absence of the vasa deferentia (CBAVD) have unilateral renal agenesis documented by abdominal ultrasonography127. Moreover, since the seminal vesicles and vasa deferentia are formed by the Wolffian ducts, a variable degree of seminal vesicle abnormalities is expected. Most patients with vasal agenesis submitted to transrectal ultrasonography (TRUS) will exhibit seminal-vesicle hypoplasia or

EVALUATION OF THE SUBFERTILE MALE

agenesia. For the same embryological reasons, it is possible to explain why a patient with unilateral absence of the vas deferens may have segmental abnormality of the contralateral vas deferens and seminal vesicle and present with azoospermia. In terms of seminal analysis, patients with CBAVD commonly show a decrease in ejaculate volume, fructose content and semen pH128,129. Another remarkable clinical aspect about CBAVD is its association with mutations of the cystic fibrosis transmembrane conductance regulator (CFTR) gene. Almost all patients manifesting cystic fibrosis have CBAVD, and almost 70% of men with CBAVD have mutation of the CFTR gene. It is also important to mention that routine laboratory methods may fail to identify all CFTR abnormality in a man with CBAVD, and the presence of a mutation cannot be ruled out. Assuming that we cannot be a 100% sure that a man with CBAVD does not harbor a genetic abnormality in the CFTR gene, if his semen is to be used, it is very important to test his wife for CFTR gene mutations. The chance that she may be a carrier is estimated to be 4%130.

Azoospermia due to obstruction/spermatogenesis failure When bilateral testicular atrophy and vasal agenesis are excluded, azoospermia may occur due to ductal obstruction at some level in the reproductive system, or abnormal spermatogenesis. To determine the etiology of the azoospermia, we must rely upon FSH measurements, ejaculate volume and testicular biopsy. Normal ejaculate volume

Patients with normal ejaculate volume may present either ductal obstruction or abnormalities of spermatogenesis, and the FSH level could be used to direct the next step. If the FSH level is high (greater than twice the upper limit range), the patient has severe germ- and Sertoli-cell dysfunction, and there is no need to perform a testicular biopsy for diagnostic purposes. However, if

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testicular sperm extraction with ICSI is being considered, a testicular biopsy may be indicated, initially, for prognostic purposes. On the other hand, patients should be warned that the presence of sperm in a previous biopsy specimen does not assure that sperm will be found on the day of ICSI. For that reason, the role of prognostic biopsy in patients with a very high FSH concentration has been considered rather controversial. However, when a patient has a normal serum FSH level, a testicular biopsy can lead to the diagnosis, as normal serum FSH levels do not guarantee normal spermatogenesis. Testicular biopsy may be unilateral or bilateral, and a consensus about this issue has not yet been reached. If performed unilaterally, a testicular biopsy should be done on the best testis. Testicular biopsy can be performed either by a standard open incision technique or by percutaneous methods. An open surgical biopsy performed under general anesthesia can provide enough testicular tissue for histological and cryopreservation purposes. The presence of sperm in the fresh specimen may avoid the need for repeat surgery. If normal testicular histology is confirmed, obstruction at some level in the semen pathway must be present, and the location of the obstruction may be determined. Vasectomy is, without any doubt, the most common cause of ductal obstruction. After that, bilateral epididymal obstruction is considered the most important cause of obstructive azoospermia and microscopic surgical exploration may show dilated epididymal tubules. Low ejaculate volume

Patients with azoospermia and a very low ejaculate volume (< 1 ml) may have gonadotropin insufficiency, CBAVD or ejaculatory duct obstruction (EDO). Ejaculatory dysfunction does not cause azoospermia, but rather aspermia or hypospermia with oligozoospermia. The determination of additional seminal parameters, such as pH and fructose concentration, may be useful in determining the presence of

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total EDO, as the seminal vesicles produce an alkaline secretion containing fructose. However, caution should be taken, because the results of semen pH and fructose testing may be misleading if they are not properly performed. The method of choice for determining EDO is transrectal ultrasonography (TRUS)70,131. Although vasography is considered an alternative method, TRUS is minimally invasive and prevents the possible risk of vasal injury associated with vasography. For a more detailed description, the reader is directed to the above section on ‘Imaging the reproductive tract’.

GENETIC EVALUATION Screening for genetic alterations in infertile men is usually recommended in cases of severe oligozoospermia, non-obstructive azoospermia and in azoospermia due to congenital bilateral absence of the vasa deferentia. The most common genetic tests used for evaluating the origin of these alterations are: karyotyping, screening for Y-chromosome microdeletions and mutations in the cystic fibrosis genes. Genetic screening may also be recommended for patients with varicocele or cryptorchidism, since more than one factor may be present.

Karyotype It has been known for decades that constitutional chromosome abnormalities are more prevalent in infertile men than in fertile men132, and these are inversely related to sperm concentration. An estimated 5% (2–8%) of infertile men present with chromosomal alterations132–134, but in the azoospermia group this number may reach 15%, and this is mostly due to 47,XXY aneuploidy, or Klinefelter’s syndrome, the most common chromosomal abnormality in men with severe infertility135. Almost all men with a 47,XXY karyotype are azoospermic, while 46,XY/47,XXY mosaic men may show a limited number of sperm in their ejaculates. During testicular sperm extraction

(TESE), sperm is found in 50% of 47,XXY men136, and most of them are 23,X or 23,Y, although there is an increase in 24,XX or 24,XY cells. While the majority of chromosome alterations in azoospermic men are sex chromosome-related, a wide array of abnormalities has been described, such as reciprocal and Robertsonian translocations, inversions, duplications and deletions. Some preliminary studies have shown that there is an increase in prenatally detected sex chromosome abnormalities during gestations from ICSI, when compared with gestations from natural conception137. It has also been well described that infertile men possess an increase in chromosome alterations both in somatic cells and in gametes138,139. Knowing that when men possess these alterations there is an increased risk of abortion, or of children being born with genetic and congenital alterations, karyotyping is recommended for patients presenting with azoospermia or with severe oligozoospermia before performing ICSI.

Y-chromosome microdeletions Since the original work by Tiepolo and Zuffardi140, many studies have demonstrated an association between male infertility and the presence of microdeletions in the long arm of the Ychromosome (Yq)141,142. Karyotyping will not reveal these microdeletions, and therefore molecular techniques such as the polymerase chain reaction (PCR) must be used. There are three loci (chromosomal regions) associated with spermatogenesis in this region, and they have been termed azoospermia factors: AZFa, AZFb and AZFc. Many candidate genes have been isolated in infertile men: DBY and USPY9 in AZFa143,144, RBMY1 in AZFb145 and DAZ in AZFc146. Other genes have been identified in Yq, but their contribution to the AZF phenotype has yet to be determined. Y-chromosome microdeletions may lead to primary testicular insufficiency, which is characterized by azoospermia or severe oligozoospermia.

EVALUATION OF THE SUBFERTILE MALE

Around 60% of Y-chromosome microdeletions occur in region AZFc, while 15% occur in AZFb and 5% in AZFa. The other 20% involve more than one AZF region141. Between 10 and 15% of infertile men present Yq microdeletions142. In patients with idiopathic severe oligozoospermia this figure may rise to 18%, and in idiopathic azoospermia to 20%. The region in which the microdeletion occurs may also determine how spermatogenesis will be affected. AZFa microdeletions are associated with complete absence of germ cells, in a syndrome known as Sertoli cell only syndrome (SCOS). AZFb microdeletions, on the other hand, determine maturation arrest. Finally, AZFc microdeletions, unlike AZFa and AZFb, are not associated with any specific phase of spermatogenesis. Phenotypical alterations may range from azoospermia to, more typically, oligozoospermia147. The presence of AZFa and AZFb microdeletions greatly decreases the chances of finding sperm in the testes, and therefore screening for Y-chromosome microdeletions is very useful in determining the prognosis for patients with non-obstructive azoospermia148. Men with important seminal alterations associated with clinical conditions such as varicocele or cryptorchidism may also present Yq microdeletions149–151. Patients carrying AZFc microdeletions are not necessarily azoospermic, and thus are candidates for ICSI using sperm from the ejaculate or the testes. In these cases, Y chromosome-bearing sperm will transport the microdeletions. Male children born from these patients will therefore also possess these deletions152–154. Some recent articles have suggested an increased risk of altered gonadal formation and Turner’s syndrome (45,X0) in children from Yq-microdeletion patients155–158, and this leads to important ethical issues. Patients with non-obstructive azoospermia or severe oligozoospermia should be screened for Y-chromosome microdeletions even if signs of testicular lesions are present, since both may occur simultaneously, and ICSI may allow transmission of these deletions. On the other hand, patients with a sperm count of more than 10 × 106

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sperm/ml do not need to be screened, since Yq microdeletions are very rare in this case.

CFTR gene Cystic fibrosis (CF) is one of the most common recessive autosomal diseases in Caucasians, with a prevalence of one affected person per 2500 births. One in 25 individuals is an asymptomatic heterozygote. CF is caused by a mutation in the gene encoding for the cystic fibrosis transmembrane conductance regulator protein (CFTR). The most common mutation in the CFTR gene is the deletion of a phenylalanine in position 508 (∆508), but more than 1000 different mutations have been identified, according to the CFTR online database (www.genet.sickkids.on.ca/cftr). Congenital bilateral absence of the vasa deferentia (CBAVD) is in many cases considered an incomplete or mild form of CF. Around 70–80% of these men are heterozygotes for CFTR mutations159,160. Another mutation associated with CBAVD is the presence of five thymines in intron 8 (the ‘wild-type’ has seven or nine), designated the ‘5T allele’. This alteration leads to the nontranscription of exon 9, and thus to low levels of CFTR161. CBAVD is diagnosed in 1.5% of all cases of male infertility. Most (60%) heterozygote mutations are compound mutations (different mutations in each copy of the CFTR gene)162. Congenital unilateral absence of the vas deferens (CUAVD) is also related to CFTR mutations. Although the prevalence of mutations in these patients varies significantly (10–73%)163,164, it has been established that the occurrence of CUAVD is due in part to the production of a defective CFTR protein. Therefore the clinical manifestation of patients with a CFTR mutation may be azoospermia or oligozoospermia, associated with CBAVD and CUAVD, respectively. It is important to note that these patients demonstrate normal spermatogenesis. They are therefore candidates for ICSI by collecting sperm from the epididymides or the testes. If their female partner is a heterozygote for the CFTR mutation,

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their children are at risk of presenting classic cystic fibrosis. Therefore, if a patient presents with CBAVD or CUAVD, both partners in the couple should be investigated for CFTR mutations, and appropriate genetic counseling should be provided.

CONCLUSIONS We may expect that 30% of infertile couples are so due to a significant isolated male factor, and associated male and female factors are present in an additional 20%. Although male factors contribute to half of the cases of infertility, the pathophysiological mechanisms of male infertility are so poorly understood that most infertile men are described as idiopathic oligo/astheno/teratozoospermia rather than having an etiological diagnosis. As a consequence, there is no scientific basis for clinical treatment, except for gonadotropin deficiency. The use of assisted reproductive technologies, particularly ICSI, has become the rule, instead of the exception, and physicians and patients have shifted focus from the cause of infertility to the ultimate goal, pregnancy. To reverse the present situation, we need to improve current male-factor diagnostic tools, emphasizing genetics and post-receptor mechanisms, which will open new venues for protein- or gene-based therapies directed towards the underlying cause and mechanisms of male infertility.

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varicocele-associated infertility (VAI). Fertil Steril 2000; 74: 555 Platt B, Busselberg D. Combined actions of Pb+2, Zn+2 and Al+3 on voltage-activated calcium channel currents. Cell Mol Neurobiol 1994; 14: 831 Willott GM. Frequency of azoospermia. Forensic Sci Int 1982; 20: 9 Jarow JP, Espeland MA, Lipshultz LI. Evaluation of the azoospermic patient. J Urol 1989; 142: 62 Odival T Jr, et al. Search and identification of spermatozoa and spermatids in the ejaculate of nonobstructive azoospermic patients. Int Braz J Urol 2005; 31: 42 Rao MM, Rao DM. Cytogenetic studies in primary infertility. Fertil Steril 1977; 28: 209 Sokol RZ, Swerdloff RS. Endocrine evaluation. In Lipshultz LI, Howards SS, eds. Infertility in the Male, 3rd edn. St Louis: Mosby-Year Book, 1997: 210 Schlegel PN, Shin D, Goldstein M. Urogenital anomalies in men with congenital absence of the vas deferens. J Urol 1996; 155: 1644 Anguiano A, et al. Congenital bilateral absence of the vas deferens. A primarily genital form of cystic fibrosis. JAMA 1992; 267: 1794 Chillon M, et al. Mutations in the cystic fibrosis gene in patients with congenital absence of the vas deferens. N Engl J Med 1995; 332: 1475 The Male Infertility Best Practice Policy Committee of the American Urological Association, and the Practice Committee of the American Society for Reproductive Medicine. Report on evaluation of the azoospermic male. Fertil Steril 2004; 82 (Suppl): 131 Jarow JP. Seminal vesicle aspiration in the management of patients with ejaculatory duct obstruction. J Urol 1994; 152: 899 Chandley AC. The chromosomal basis of human infertility. Br Med Bull 1979; 35: 181 Hargreave T. Genetically determined male infertility and assisted reproduction techniques. J Endocrinol Invest 2000; 23: 697 Gekas J, et al. Chromosomal factors of infertility in candidate couples for ICSI: an equal risk of constitutional aberrations in women and men. Hum Reprod 2001; 16: 82 de Braekeleer M, Dao TN. Cytogenetic studies in male infertility: a review. Hum Reprod 1991; 6: 245 Palermo GD, et al. Births after intracytoplasmic injection of sperm obtained by testicular extraction

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from men with nonmosaic Klinefelter’s syndrome. N Engl J Med 1998; 338: 588 Van Opstal D, et al. Determination of the parent of origin in nine cases of prenatally detected chromosome aberrations found after intracytoplasmic sperm injection. Hum Reprod 1997; 12: 682 Meschede D, et al. Chromosome abnormalities in 447 couples undergoing intracytoplasmic sperm injection – prevalence, types, sex distribution and reproductive relevance. Hum Reprod 1998; 13: 576 Colombero LT, et al. Incidence of sperm aneuploidy in relation to semen characteristics and assisted reproductive outcome. Fertil Steril 1999; 72: 90 Tiepolo L, Zuffardi O. Localization of factors controlling spermatogenesis in the nonfluorescent portion of the human Y-chromosome long arm. Hum Genet 1976; 34: 119 Foresta C, Moro E, Ferlin A. Y-chromosome microdeletions and alterations of spermatogenesis. Endocr Rev 2001; 22: 226 Pryor JL, et al. Microdeletions in the Y-chromosome of infertile men. N Engl J Med 1997; 336: 534 Foresta C, Ferlin A, Moro E. Deletion and expression analysis of AZFa-genes on the human Y-chromosome revealed a major role for DBY in male infertility. Hum Mol Genet 2000; 9: 1161 Sun C, et al. An azoospermic man with a de novo point mutation in the Y-chromosomal gene USP9Y. Nat Genet 1999; 23: 429 Elliott DJ, et al. Expression of RBM in the nuclei of human germ cells is dependent on a critical region of the Y-chromosome long arm. Proc Natl Acad Sci USA 1997; 94: 3848 Reijo R, et al. Diverse spermatogenic defects in humans caused by Y-chromosome deletions encompassing a novel RNA-binding protein gene. Nat Genet 1995; 10: 383 Vogt PH. Molecular genetics of human male infertility: from genes to new therapeutic perspectives. Curr Pharm Des 2004; 10: 471 Hopps CV, et al. Detection of sperm in men with Ychromosome microdeletions of the AZFa, AZFb and AZFc regions. Hum Reprod 2003; 18: 1660 Moro E, et al. Y-chromosome microdeletions in infertile men with varicocele. Mol Cell Endocrinol 2000; 161: 67 Foresta C, et al. Y-chromosome microdeletions in cryptorchidism and idiopathic infertility. J Clin Endocrinol Metab 1999; 84: 3660

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151. Krausz C, et al. A high frequency of Y-chromosome deletions in males with nonidiopathic infertility. J Clin Endocrinol Metab 1999; 84: 3606 152. Kent-First MG, et al. The incidence and possible relevance of Y-linked microdeletions in babies born after intracytoplasmic sperm injection and their infertile fathers. Mol Hum Reprod 1996; 2: 943 153. Page DC, Silber S, Brown LG. Men with infertility caused by AZFc deletion can produce sons by intracytoplasmic sperm injection, but are likely to transmit the deletion and infertility. Hum Reprod 1999; 14: 1722 154. Oates RD, et al. Clinical characterization of 42 oligospermic or azoospermic men with microdeletion of the AZFc region of the Y-chromosome, and of 18 children conceived via ICSI. Hum Reprod 2002; 17: 2813 155. Siffroi JP, et al. Sex chromosome mosaicism in males carrying Y-chromosome long arm deletions. Hum Reprod 2000; 15: 2559 156. Jaruzelska J, et al. Mosaicism for 45,X cell line may accentuate the severity of spermatogenic defects in men with AZFc deletion. J Med Genet 2001; 38: 798 157. Papadimas J, et al. Ambiguous genitalia, 45,X/46,XY mosaic karyotype, and Y-chromosome

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microdeletions in a 17-year-old man. Fertil Steril 2001; 76: 1261 Patsalis PC, et al. Effects of transmission of Y-chromosome AZFc deletions. Lancet 2002; 360: 1222 Quinzii C, Castellani C. The cystic fibrosis transmembrane regulator gene and male infertility. J Endocrinol Invest 2000; 23: 684 Casals T, et al. Heterogeneity for mutations in the CFTR gene and clinical correlations in patients with congenital absence of the vas deferens. Hum Reprod 2000; 15: 1476 Chu CS, et al. Genetic basis of variable exon-9 skipping in cystic fibrosis transmembrane regulator mRNA. Nat Genet 1993; 3: 151 Patrizio P, et al. Aetiology of congenital absence of vas deferens: genetic study of three generations. Hum Reprod 1993; 8: 215 Dork T, et al. Distinct spectrum of CFTR gene mutations in congenital absence of vas deferens. Hum Genet 1997; 100: 365 Casals T, et al. Extensive analysis of 40 infertile patients with congenital absence of the vas deferens: In 50% of cases only one CFTR allele could be detected. Hum Genet 1995; 95: 205.

9 The basic semen analysis Roelof Menkveld

INTRODUCTION

by a description of the acrosome and the presence of vacuoles in the sperm head by Williams et al. in 19346. Over this time period, different methods were proposed for the evaluation of semen samples with the inclusion of various semen parameters and standards for normality4,5,7–12. Further standardization and minimum requirements for the methodology of a semen analysis performance and ‘normal’ semen variable standards were established in 1951 by the American Fertility Association13. This was followed by the contributions of MacLeod and Gold14, Freund15,16 in the 1960s and Eliasson in the 1970s17,18, especially with regard to sperm morphology. In order to obtain better world standardization of semen analysis, the first World Health Organization (WHO) manual was published in 198019, followed by the 198720, 199221 and 1999 editions22. Requirements for a complete extended semen analysis as performed today are undergoing changes according to the demands of time and new developments in the fields of spermatology and andrology, as well as assisted reproductive technologies (ART). Today, a complete basic semen analysis must also include screening tests for the presence of antisperm antibodies, such as the mixed antiglobulin reaction (MAR) test23, and a leukocyte peroxidase test24 aimed at identifying the presence of polymorphonuclear leukocytes.

The scientific approach to establish a male’s fertility potential by means of the semen analysis started in 1677, with van Leeuwenhoek’s letter to the Royal Society of London describing the discovery of the human spermatozoon by Johan Ham. According to Schirren1, van Leeuwenhoek stated that in the case of a sterile marriage, the microscope could solve the problem as to the responsible partner. A more scientific approach to the semen analysis procedure was introduced by the end of the 19th century, when Lode2 performed the first dilutions of semen samples before performing a sperm count with the aid of a hemocytometer, finding a mean sperm concentration of 60.88 × 106/ml for the four males investigated. In 1941, Hotchkiss3 published a basic grading system for sperm motility evaluation that was modified by MacLeod and Heim4 in 1945 to a system in which the motility and progressive activities were recorded separately, the motility in 10% units and the forward progression on a scale of 0–4. Belding5 made one of the first contributions towards sperm morphology evaluation as we know it today by suggesting a classification for abnormalities of the head, midpiece and tail, which could be indicated as a single abnormality or a combination of abnormalities. This was followed 141

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Other tests are still mentioned in the 1999 WHO manual22, including the sperm–mucus penetration test25, performed with good periovulatory human cervical mucus or with human mucus replacements such as bovine mucus26 or hyaluronate27, the sperm–cervical mucus contact test28, the zona-free hamster-ovum penetration test29 and the hemizona assay test30, which are performed to a lesser and more selective extent, for example when indicated by unexplained poor ART results. Newly developed tests such as the DNA status of the spermatozoa31, the acrosome reaction test32, the reactive oxygen species (ROS) activity of the spermatozoa and especially leukocytes33 and the antioxidation capacity of seminal plasma34 have recently attracted more attention. However, owing to new developments and advances in ART procedures, especially intracytoplasmic sperm injection (ICSI), McDonough35 doubted the future role of the standard basic semen analysis, and wrote: ‘Traditional sperm analysis as a clinical test may become nothing more than an ancestral heirloom. It may be performed spasmodically by those who know how to do it, like a 1940-air show or laparotomy, to remind us of the good old days. We have come to the end of something. Surely someone will want to carve a headstone for traditional sperm analysis or perhaps a mausoleum will be more fitting.’ It is difficult to agree with the above concepts. Even in the light of new developments such as in vitro fertilization (IVF) and especially ICSI, semen analysis has and will be the most important test in the initial investigation of a male’s fertility potential. It is therefore extremely important that a semen analysis should be performed skillfully and properly. If the necessary background data are known, including a short personal and medical history, so that the results can be interpreted correctly, the basic (complete) semen analysis will always remain the cornerstone of the initial investigation of a male’s fertility potential, as part of a couple’s basic infertility investigation. A complete semen analysis can be divided into the following four categories: (1) background

data, (2) physical analysis, (3) microscopic analysis and (4) additional procedures. Biochemical analyses and functional tests should be performed on repeated semen analysis when indicated, for instance after unexpected poor results with ART.

THE BASIC SEMEN ANALYSIS Specimen handling Semen samples present a possible biohazard since they may contain harmful viruses, e.g. human immunodeficiency virus (HIV), hepatitis B and herpes. Therefore, semen samples should always be handled with care, as if infected, and the wearing of protective gear is advised (gloves, masks and spectacles). Further information is given in the 1999 WHO manual22, based on the work of Schrader36.

Background data A semen analysis cannot be interpreted unless some basic facts are known, namely the method by which the sample was produced, the time lapse between production and analysis, days of abstinence and the type of container used, as these factors can have an influence on the results, as discussed below. These factors are the so-called background data, and should also include data from a succinct medical history taken when the semen sample was received. Methods for the production of semen

Today, it is expected that the semen sample should be collected in a specially equipped, and if necessary air-conditioned, room at or in close proximity to the laboratory, especially when special tests are involved37,38. This method has the advantage that the exact time of semen production and the time lapse between production and investigation are known, and observations such as the presence of coagulation and the occurrence of liquefaction can be made. The way in which the sample is

THE BASIC SEMEN ANALYSIS

produced is also controlled. Many patients may produce a sample by coitus interruptus or by using a spermicidal condom if the sample is produced at home. Coitus interruptus has the disadvantage that the first part of the sample may be lost. An indication that the sample may have been produced by coitus interruptus will be the presence of vaginal epithelium cells38. When the patient collects the sample at the laboratory, a relationship can be built with the patient, information is easy to obtain and the patient’s enquiries can be answered. It often happens that a sample is brought to the laboratory and left on a counter without any information. The results of such a semen analysis cannot be evaluated or interpreted because the method of production, days of abstinence and time of ejaculation are not known. For patients having problems producing a semen sample by masturbation, a wide range of special condoms are available, such as silastic condoms (e.g. Seminal Collection DeviceTM, HDC Corp., Milipitas, CA, USA)39, the Seminal PouchTM made of polyethylene (Milex Products Inc., Chicago, USA), condoms made of polyurethane (Male-Factor PakTM; FertiPro NV, Beernem, Belgium) and complete kits (Hy-geneTM Kit, FertiPro) for transportation of the sample to the laboratory22. Normal latex condoms should not be used for semen collection as they may impair sperm motility due to their spermicidal properties. Semen samples should thus be produced by masturbation into a clean plastic container that is sterile-packed at shipment, or otherwise should be separately sterilized at the laboratory. The patient is instructed to urinate and then to wash his hands with soap and water and the glans of the penis with water alone, before producing the sample. The patient should be asked about the precise period of abstinence, as well as a short medical history. Questions regarding his medical history should include information on the occurrence of any previous infections or illnesses, especially in the past 3 months, if it is his first visit, or since his

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previous semen analysis when a repeat analysis is being performed. Also included in this medical history should be questions on any recent medication or anesthesia in the past 3 months, any previous history of operations of the urogenital tract, especially involving the bladder, an orchidopexy, orchiectomy, varicocelectomy or testicular biopsy, or whether he has had any severe injuries of the testicles or orchitis. A note should also be made about his smoking and drinking habits40. Containers

In the early years, glass containers were used, but this practice should be discouraged, owing to the possibility of virus contamination and the fact that the glass containers have to be washed and sterilized after use. There is also the possibility that the container may break while being washed, or even when the man is producing the semen sample40. The ideal container is a 60–100-ml widemouth plastic jar made of polypropylene, with a screw cap that fits tightly to prevent any loss of semen when it is transported. In our experience, some types of plastic (e.g. polystyrene) have the disadvantage that they may cause increased viscosity, or may be toxic to the spermatozoa and may influence motility. Before the introduction of new containers in a laboratory, these should always first be tested for any negative effects on the semen sample and ART outcome39,40. Abstinence

The profound effect of abstinence on semen parameters, especially semen volume and sperm concentration, is well known41. It is therefore important that a fixed period of abstinence should be prescribed so that optimum results can be expected, the semen analysis is performed according to more standardized conditions and the results of different semen analyses can be compared with each other. If this is not done, it is impossible to know whether differences between semen parameters of different semen samples from the same patient are due to normal variation, a difference in days of abstinence or both. The

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variation of 2–7 days suggested in the WHO manual22 is too long42, and should be standardized to 3–4 days40,43. The question is now raised whether the period of abstinence should be expressed as days or the exact number of hours42. For routine semen analysis the number of days will be acceptable, but for medical trials the exact number of hours is advised. After production of the semen sample the container is placed in an incubator at 37°C until complete liquefaction has occurred. The sample is then ready for evaluation, and is usually transferred to a graduated conical test-tube for further processing.

Physical parameters Parameters describing the appearance of the sample are classified by Freund44 and Zaneveld and Polakoski45 as physical parameters, and include the color, liquefaction and viscosity, while coagulation and odor can also be added to this category. Although strictly speaking a biochemical characteristic, pH is also included in this group. All these parameters are simple to evaluate and are mainly determined by visual examination. Coagulation

This is an important aspect of semen analysis that is ignored by many investigators, mainly because many semen samples are still produced at home instead of at the laboratory. Human semen is ejaculated in a liquefied state, but is quickly transformed into a semisolid state or coagulum, probably under the influence of the enzyme protein kinase46 secreted by the seminal vesicles. In a normal situation, nearly the whole sample is transformed into the coagulated state, and only a very small part remains liquefied. This is generally regarded as the first portion of the ejaculate, containing the major part of the motile sperm fraction. In cases where coagulation does not occur it may be the result of congenital absence of the vas deferens and the seminal vesicles, as the coagulating enzymes originate from the seminal vesicles,

and is then also associated with the absence of fructose in the seminal plasma. Liquefaction

In a normal sample, liquefaction occurs within 10–20 minutes. This is caused by a proteolytic enzyme fibrinolysin secreted by the prostate47, as well as two other proteolytic enzymes, fibrinogenase and aminopeptidase48. Liquefaction therefore serves as an indicator of normal prostatic function. After complete liquefaction the sample will appear homogeneous in composition and color. Small roundish particles may still be present in some samples; however, this can be regarded as normal, and they will usually dissolve within an hour. If liquefaction takes more than 20 minutes or does not occur at all, it is a sign that the prostate is not functioning normally, usually as a result of previous prostatitis. In some cases this non-liquefaction of semen may be a cause of infertility, as the spermatozoa are not released from the coagulum40. Viscosity

As long ago as 1934, Cary and Hotchkiss7 described the consistency of semen as slightly more viscous than water. The most convenient way to determine viscosity is by means of a modified pipette method37. The semen is drawn into a Pasteur pipette and slowly released in a drop-wise fashion. The viscosity is regarded as normal when single drops are formed that are released within a distance of 20 mm from the point of the pipette. If threads are longer than 20 mm the viscosity can be regarded as increased40. It is also important to distinguish between a delayed period of liquefaction (non-homogeneous appearance) and an increase in the viscosity (homogeneous but ‘sticky’). Increased viscosity may be the result of abnormal prostatic function due to an infection in the genital tract, prostate or seminal vesicles49, or an artifact as a result of the use of an unsuitable type of plastic container, frequent ejaculation or the psychological state of the

THE BASIC SEMEN ANALYSIS

patient. A constant increase in viscosity may be regarded as a cause of infertility45 for in vivo conception, and can also have an adverse effect on the determination of spermatozoa concentration and motility. Biochemical means should be used to reduce high semen viscosity, for example α-amylase50 and chymotrypsin51, while another method is the addition of an equal volume of a medium such as saline, phosphate-buffered saline or culture medium, followed by repeat pipetting with a wide-bore pipette52. In these cases, care should be taken that the sperm concentration is correctly calculated, taking into consideration the extra dilution effect of the added fluid53. Volume

The most common method still used today to determine the volume is by transferring the sample to a 15-ml graduated conical tube and reading the volume to the nearest 0.1 ml. Determination of the volume can also be performed by means of weighing samples, taking the total weight of the sample and container minus the container weight determined beforehand. The weight is expressed as the nearest 0.1 ml, taking 1 g equal to 1 ml53. The normal volume of an ejaculate after 3–5 days of sexual abstinence is 2–6 ml. Hotchkiss11 stressed the importance of a normal volume, as this is needed for good buffering function of the seminal pool against the acid secretions of the vagina. If the volume of a semen sample is smaller than 1.0 ml, it is important to establish whether a complete sample was collected. This is important, as the first portion containing the major amount of sperm with the best motility is often lost. A low volume may, however, also be the result of an obstruction due to a previous infection of the genital tract, or of congenital absence of the seminal vesicles and vas deferens; this condition will be associated with the absence of fructose45. A small volume may also be due to retrograde ejaculation, especially if the patient has had any previous surgery of the prostate or the bladder neck.

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Retrograde ejaculation can be diagnosed by investigation of the urine after ejaculation. Color

By paying attention to the color of the semen sample, an indication of possible pathology of the semen can already be obtained. Cary and Hotchkiss7 described the color of normal semen as opaque and grayish, which will change to yellowish with an increase in the days of abstinence. Hotchkiss11 noticed that fresh blood will give semen a reddish color and old blood a brownish color, which may be caused by recent inflammation. In cases of inflammation a more yellowish color may exist, while samples with a low sperm concentration will usually have a transparent and watery consistency. Schirren1 found that certain types of medicine such as antibiotics might discolor the semen. Odor

Although semen has a strong, distinctive odor, derived from the prostatic secretions, this parameter is seldom used. The odour is sometimes compared to that of the flowers of the chestnut or St John’s bread tree. It is thought that the odor is caused by oxidation of the spermine secreted by the prostate. Only with absence of the odor or when an uncharacteristic odor is present should a note be made, as this is usually associated with an infection45, or is the result of a long period of abstinence37. pH

Preference should be given to pH measurement using a special pH indicator paper (range 6.4–8.0; Merck #9557), for hygiene reasons and also the possibility that sexually transmitted diseases may be transferred when using the glass-electrode method. After liquefaction, a drop of semen is placed on the indicator strip and immediately compared against a color scale. The pH of a normal ejaculate may vary between 7.2 and 7.822. In cases of acute prostatitis, vesiculitis or bilateral epididymitis, the pH will always be more than

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8.045. In cases of chronic infection of the above organs, the pH will always be below 7.2, and can be as low as 6.6. With an obstruction of the ejaculatory duct or in cases where only prostatic fluids are secreted, the pH will also be less than 7.0, and if the sample is azoospermic this low pH may also indicate the presence of bilateral congenital absence of the vas deferens22.

Microscopic analysis Wet preparation examination

After completion of the physical examination, the centrifuge tubes are placed on a cradle or roller system39 for the duration of all subsequent procedures. This can be done at room temperature or at 37°C. After 10 minutes of gentle mixing, a drop of semen is taken with a positive-displacement pipette and placed on a precleaned glass slide kept at 37°C until use. The size of the drop of semen will depend on the size of the coverslip used, so that the depth of fluid between the microscope slide and the coverslip is about 20 µm, to allow maximum free movement of the spermatozoa and still optimum visibility with a 40× objective. The standard drop size most often used is 10 µl for a 20 × 20-mm coverslip. Complete guide tables for different sizes of coverslips and corresponding drop sizes to be used can be found in several publications22,39,53. The preparation is left for a few minutes to stabilize before examination. General appearance All examinations of wet preparations are done with phase-contrast optics, first at a 100× or 150× or low-power field (LPF) magnification (with 10× or 15× objectives) to obtain an overall view, and then at 400× or highpower field (HPF) magnification (with a 40× objective). The examination starts with scanning through ten LPFs to get an impression of the general appearance of the sample. The impression obtained here will dictate all subsequent procedures, such as the performance of a MAR test23 if enough motile spermatozoa are present, or a vital

staining test54 when the motility is low. An estimation of the number of spermatozoa per HPF is made, which will be used to determine the dilution of the sample for calculation of the sperm concentration and drop size to be used for preparing smears for sperm morphology evaluation. Agglutination and presence of other cells The sample is also examined for the presence of sperm agglutination. Two types of agglutination can be observed. In the first instance, agglutination can be due to non-specific factors where, in most cases, non-motile spermatozoa adhere to cells present in the seminal plasma; when this occurs it is termed aggregation39. The second is specific agglutination, caused by antisperm antibodies, which consists mostly of motile spermatozoa clumps with only minimal involvement of other cells or debris39. Agglutination is described as negative (–), occasional (±), slight (+), moderate (++) or severe (+++)40, or as an appropriate percentage to the nearest 5%39,53. A note is also made of the presence of other cells, such as round cells, and the presence of spermine phosphate crystals, recorded in the same way as for agglutination. The presence of any organisms is also recorded. Analysis of quantitative parameters

The parameters classified under this heading are those that are regarded by many investigators to constitute a complete or standard semen analysis, and include estimation of the percentage and grade of sperm motility, the vital staining procedure to determine the percentage of live spermatozoa, if indicated due to poor motility, the spermatozoa concentration and the morphology of the spermatozoa. The MAR test23 and a leukocyte peroxidase test24 should now also be included as routine procedures55. Motility and forward progression Motility is now mostly determined in one of two manners. The first is by manual observation of the sample with phase-contrast optics. More recently,

THE BASIC SEMEN ANALYSIS

automated computer-assisted semen analysis (CASA) techniques have been introduced with varying degrees of success22,39. This is discussed briefly under a separate heading dealing with CASA. For the manual method, the wet preparation slide, as prepared for the initial examination, can be used and the evaluation is performed as described in the WHO22 and European Society for Human Reproduction and Embryology (ESHRE)53 manuals. The exact aliquot of semen to provide a depth of 20 µm is of importance due to the rotary and spiral movement pattern of progressive motile spermatozoa. If the time interval between the initial wet preparation and observation for motility is too long, a new preparation should be made, and examination of the wet preparation should begin as soon as the flow of the semen drop has ceased. If this has not occurred within a minute, a new preparation should be made and examined53. Spermatozoa are classified according to the rapidity of their forward progressive motility into four grades, from grade a to grade d, as follows: • Grade a = rapid progressive motility; • Grade b = slow or sluggish progressive motility; • Grade c = non-progressive motility; • Grade d = immotile. Definitions of rapid and slow forward motility will differ, depending on whether the motility evaluation is performed at room temperature or at 37°C by means of a hot stage fitted on the microscope. For rapid motility at 37°C, the spermatozoa should travel ≥ 25 µm per second, and at room temperature ≥ 20 µm per second, i.e. the distance of five and four sperm heads, respectively, as spermatozoa move more rapidly at 37°C22. If the forward progression is < 5 µm per second, for both room temperature and 37°C determinations, spermatozoa are regarded as having a non-progressive grade c motility. Between these limits, spermatozoa will be regarded as having a grade b or slow forward motility.

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At least 200 spermatozoa should be counted in five separate high-power magnification fields with the aid of phase-contrast microscopy. The percentages of the different categories must add up to 100%. The count should be repeated on a separate wet preparation. The results of the two counts are then averaged, provided that they are within acceptable limits that can be calculated according to a method provided in the ESHRE manual53. Poor motility or asthenozoospermia can be caused by several factors. One reason may be artifacts caused by the wrong method of collection, such as use of a condom which may be spermtoxic, contamination by vaginal secretions, the use of lubricants56, an incomplete sample, a long delay in transportation of the sample to the laboratory or exposure to extreme temperatures. Artifacts can also be caused by technical factors such as cold shock due to use in the laboratory of cold containers, slides and pipettes, the use of unsuitable, contaminated or wet containers, storage of the sample at an adverse temperature57,58 or wrong thickness of the wet preparation (< 10 µm), hindering the free rotational movement of the spermatozoa59. Poor motility can also be due to structural abnormalities of the midpiece60, or the short-tail61 and immotile cilia or Kartagener’s syndrome62. Poor motility may also be caused by unfavorable environmental conditions during the formation and maturation of spermatozoa before they are released from the Sertoli cells63,64, or during transport through the epididymis65 and ductal system, or via abnormal functions of the prostate or seminal vesicles caused by acute infections or inflammation of the accessory glands. Other factors that can cause poor motility are the presence of hematospermia, a varicocele, chromosomal aberrations, bacterial infections and an abnormal pH58,66, as well as the presence of certain metals or metal ions67. Sperm concentration In 1929, the well-known article of Macomber and Sanders68 was published in which their sperm counting technique, which forms the basis for most of the techniques still

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used today, was described. A 1:20 dilution was made with the aid of a white blood-cell pipette, and the count performed on a hemocytometer. The diluting fluid consisted of a 5% sodium bicarbonate solution to which 1% formalin was added. Over the years, the pipettes used to prepare the dilutions have changed, with the aim of measuring and delivering the semen aliquot to be diluted as accurately as possible. Concern about delivering the true measured volume of the semen aliquot is due to the higher viscosity of semen compared with water. Instead of the white blood-cell pipette, Eliasson18 used micropipettes to make a 1:50, 1:100 or 1:200 dilution. Van Zyl69 introduced the use of a glass tuberculin syringe instead of the white blood-cell pipette. With this method it is possible to make a 1:10, 1:20 or a 1:100 dilution. Menkveld et al.70 demonstrated that the results of the tuberculin syringe method compared well with the results of the white blood-cell pipette (WCP) method. In 1979, Makler introduced a special sperm-counting chamber, which was improved in 198071. With the Makler chamber it is possible to carry out sperm counts directly on undiluted semen samples, after immobilization of the spermatozoa in a hot water bath at ± 60°C. The exact amount of semen delivered to the chamber for this apparatus is not as critical as for the preparation of dilutions for counting using the standard hemocytometer. Depending on the observed sperm number per high-power field, a 1:10, 1:20 or a 1:50 dilution will be made, while some laboratories also make use of a 1:100 dilution40. It is now advised that positive-displacement pipettes should be used to deliver the semen aliquot, and a normal airdisplacement pipette to deliver the dilution fluid22,53. For a 1:10 dilution, 900 µl of the dilution fluid is placed in a small tube and 100 µl of the semen sample is transferred to the dilution fluid by means of the positive-displacement pipette. The sperm suspension is thoroughly mixed by vortex, and both sides of a hemocytometer with improved Neubauer ruling is carefully filled without spilling

the suspension over the sides of the chamber. The hemocytometer is left in a moist Petri dish for about 10 minutes for the spermatozoa to settle on the bottom of the chamber40. The number of spermatozoa in the upper left corner block (consisting of 16 smaller blocks) of the central grid, used for counting red blood cells, is counted, to determine the proportion of blocks from the 25 in the grid that should be considered for counting. A reference table is given in the WHO22 and ESHRE53 manuals indicating the number of blocks from the 25 to be included so that in all instances the number of spermatozoa counted will be more than 200. The counting procedure is repeated on the other side. By the use of a table also provided in the two manuals22,53, the actual concentrations are calculated, depending on the initial dilution and the number of blocks counted per side of the hemocytometer. It is very important to note that the two tables with conversion factors in the WHO22 and ESHRE53 manuals differ, due to the fact that the WHO manual (incorrectly) first obtains the mean of the two counts. The two counts obtained are compared to establish whether the results are within acceptable limits by means of a formula also provided in the two manuals. If the counts are not within acceptable limits, the whole counting procedure should be repeated. Estimation of the sperm concentration with the aid of computerized equipment (CASA) is gaining ground, and is now used as the routine method in many laboratories22,39. Differences still exist as to what can be regarded as a normal sperm concentration, and many different so-called normal cut-off values have been proposed, including 60 × 106 by Macomber and Sanders68, 20 × 106 by Eliasson18 and by MacLeod and Gold72 and 10 × 106/ml by Van Zyl69 and Van Zyl et al.73,74. Sperm morphology evaluation The 1999 WHO manual22 recommends that sperm morphology evaluation should be performed according to strict Tygerberg criteria. The principles for the evaluation of sperm morphology by strict Tygerberg

THE BASIC SEMEN ANALYSIS

criteria were laid down by Menkveld41 and Menkveld et al.75, while the clinical application for the in vitro situation was demonstrated by Kruger et al.76. In a follow-up study Kruger et al.77 also described the prognostic categories with strict criteria for in vitro fertilization outcome, i.e. the poor-prognosis or P-group with ≤ 4% morphologically normal spermatozoa, the good-prognosis or G-group with 5–14% morphologically normal spermatozoa and the normal group with ≥ 15% morphologically normal spermatozoa76,77. Sperm morphology evaluation according to strict criteria uses a holistic approach, starting with the preparation of clean microscope slides, the correct preparation of thin semen smears, the correct methodology for evaluation of the slides, i.e. the correct optics and magnification to be used, the correct number of spermatozoa to be evaluated and, most important of all, the criteria for a morphological normal spermatozoon as based on biological evidence41,78,79. Morphological evaluation of the semen smear can also include evaluation of the semen cytology. Therefore, two slides are prepared, one thicker smear for semen cytology evaluation and a thin smear for sperm morphology evaluation. It may also be beneficial to prepare one or two extra smears that can be kept in case the original smear is unsuitable for evaluation, or for back-up purposes18,73. Morphological evaluation of spermatozoa The morphological evaluation of spermatozoa as discussed here is based on the methodology described by Menkveld41, Kruger et al.76, Menkveld et al.75 and Menkveld and Kruger78. If indicated, due to oiliness or dirt, slides must be thoroughly cleaned before use, first washed in a detergent, rinsed in clean water and then rinsed in alcohol and airdried78. For the morphology evaluation smear, a small drop of semen is used so that a very thin smear is prepared. As a result, all the spermatozoa will be within one focus level and each sperm can be visualized separately and no more than 5–10 spermatozoa will be present per visual field at oil magnification (1000 or 1250×). The size of the

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drop will depend on the sperm concentration; for high concentrations a small drop is used (~5 µl), for normal concentrations a drop of ~10 µl will be used and for low concentrations a drop of not more than 15 µl, as with these thicker smears the semen may wash off with the staining procedure78. The thickness of the smear can also be controlled by altering the angle and speed of the microscope slide used to make the smear11,17. The slides are left until they appear to be just air-dried, i.e. only a few minutes, and are then immediately fixed in methanol or ether–alcohol (50:50) and can be stored for later reference or until staining. The modified Papanicolaou technique should be the preferred method for staining the smears22,39,75. Alternative staining methods exist, such as the rapid blood-staining methods80, and the Spermac stain method81,82. The Spermac stain is also a rapid staining method and gives excellent staining of the acrosome region and sperm tails81. The rapid blood-staining methods, such as Diff-Quik®80, cause the spermatozoa to swell slightly, and thereby may cause slight alterations to the form of the spermatozoa giving rise to bigger size measurements, which should be kept in mind when using these stains. Problems with background staining can also occur when too much seminal plasma is present on the slide83. The criteria used for a morphologically normal spermatozoon are based on the appearance of spermatozoa seen in good cervical mucus drawn from the endocervical canal shortly after intercourse for the performance of a postcoital test. These spermatozoa have a very homogeneous appearance, with only small biological variations75,78,84. According to the strict Tygerberg criteria41,75, a normal spermatozoon is defined as one having an oval form with a smooth contour and a clearly visible and well-defined acrosome, with homogeneous light blue staining. The tail should be apically inserted without any abnormalities of the neck/midpiece region; there should be no tail abnormalities; and there should be no cytoplasmic residues at the neck region or on the tail. Measurements for an abnormal cytoplasmic droplet

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and normal sperm size as seen with Papanicolaou staining are based on those described by Eliasson in 197117. The size of a normal acrosome was described as covering between 40 and 70% of the anterior part of the sperm head, with abnormal cytoplasmic droplets being present when larger than 50% of a normal-sized sperm head, which will measure 3.0–5.0 µm in length and 2.0–3.0 µm in width. The midpiece should not be longer than 1.5 times the length of a normal head and about 1 µm thick. The tail should be about 45–50 µm long and without any sharp bends17. For a spermatozoon to be classified as morphologically normal, with strict Tygerberg criteria41,75, the whole spermatozoon must be normal, as suggested by Eliasson17. However, in contrast to the views of earlier workers14,15,17, borderline or slightly abnormal spermatozoa are considered to be abnormal according to strict Tygerberg criteria41,75. This was proposed to keep allowable sperm morphology variations as small as possible, in agreement with biological variations seen in the cervical mucus75. However, the measurements as proposed by Eliasson17 and in other publications22,75 are in need of re-evaluation, as the range allowed especially for the normal head length of 3.0–5.0 µm is probably too wide. Our own experience indicates that the head length for normal spermatozoa may vary between 4.0 and 4.5 µm, with a mean of 4.07 ± 0.19 µm and a mean width of 2.98 ± 0.14 µm, as measured with a built-in microscope eyepiece micrometer (Menkveld, unpublished data). We have shown in several publications that males presenting with large-headed spermatozoa of > 5.0 µm in length, and with a proportional increase in width, and/or large acrosomes can be associated with poor in vitro fertilization results76,85 and decreased sperm functional abilities85. The presence and size of, and terminology for, cytoplasmic droplets or cytoplasmic residues are also controversial. Originally, it was stated by Eliasson17, the WHO manuals19,20 and Menkveld et al.75 that a normal cytoplasmic droplet present

on spermatozoa should be < 50% of a normal sperm head. This has been changed to < 30% in the 1999 WHO manual22. Recently, Cooper et al.86 and Cooper87 addressed this issue of the presence and size of the cytoplasmic bodies, as well as the correct terminology to be used. From the publication by Cooper87, it is clear that the retention of cytoplasmic material on spermatozoa as seen in air-dried and stained smears can be associated with impaired sperm function. It is also clear from this article and from our own experience34 that no amount of cytoplasmic material should be present on a normal spermatozoon at all, and if observed it should be regarded as an abnormality, regardless of the size or amount of cytoplasmic material present. In the article by Cooper87, it is suggested that the correct term to use if cytoplasmic material is present should be ‘excess cytoplasmic residues’, or just cytoplasmic residues. At least 200 spermatozoa should be evaluated in duplicate per slide with the highest magnification possible, i.e. 1000×, but preferably 1250×. In case of any doubt about the dimensions of a spermatozoon, the size can be measured with a micrometer. The spermatozoa should preferably not be evaluated in one area but in several areas, to increase the accuracy of the evaluation78. The latest WHO manuals21,22 recommend that spermatozoa should be classified only as normal or abnormal. A note should be made if a specific abnormality occurs in a frequency of > 20%. However, as indicated above, an abnormal spermatozoon can have only one specific abnormality or any combination of two or up to four abnormalities. To reflect this, the teratozoospermia index (TZI) was introduced as an indication of the mean number of abnormalities per abnormal spermatozoon21,22. The TZI value will therefore always be between 1 and 4. However, in the 1999 WHO manual22, cytoplasmic residues were omitted as an abnormality, and the TZI value was indicated as being between 1 and 3. This was in contrast to the ESHRE manual, which maintained that this is not correct and the value should be between 1 and 453.

THE BASIC SEMEN ANALYSIS

Evaluation of semen cytology For the evaluation of the semen cytology, i.e. investigation of the presence of different cells and organisms, a thicker smear is prepared. A small drop of egg albumin may be added to ensure better adherence of the cells to the slide. However, the more intense staining of the albumin background can sometimes make it difficult to identify the round cells present. The smears are fixed immediately in 1:1 solution of ether–alcohol for 30 minutes and stained together with the slide for morphology evaluation78. The slides are screened at a low magnification (15×), and if any cells or organisms are observed a 40× objective is used to make a better diagnosis. Cells looked for are especially polymorphonuclear white blood cells, monocytes and epithelium cells. With good staining, germinal epithelium cells, sometimes called precursors, can also be identified. The presence of identified cells, especially polymorphonuclear white blood cells (WBC) is recorded separately in a semiquantitative way by means of plus and minus signs as follows: no cells/HPF –; occasional cells/HPF ±; 1–5 cells/HPF +; 5–10 cells/HPF ++; > 10 cells/HPF +++78. A good correlation has been found between WBC identified in this manner and granulocyte white blood cells counted by means of the leukocyte peroxidase method, with ≥ +WBC/HPF correlating to ≥ 0.25 × 106 leukocytes/ml semen, a value found to be of pathological importance78,88. More details of the cytological evaluation of semen smears and the origin89 of different cell types and the identification of these cells have been published by others55,89–91. The mixed antiglobulin reaction (MAR) test A MAR test as described by Jager et al.23 must be included in all semen analysis as a routine procedure, as a screening test for the possible presence of antispermatozoa antibodies if a sufficient number of motile spermatozoa are present. The original MAR test as described by Jager et al.23 makes use of a suspension of sensitized R1R2 erythrocytes. The erythrocytes are sensitized by washing them three times with a phosphate-buffered saline

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(PBS) solution, pH 7.5. The suspension is mixed 5 : 1 with a strong incomplete anti-D serum (Behring ORRA 20/21) and incubated at 37°C for 30 minutes. After incubation the suspension is again washed three times in PBS and suspended to a hematocrit of 5–10%. This suspension can be kept at 4°C for a few days23. However, today, most laboratories make use of commercial products (MarScreen, Bioscreen, New York, USA; SpermMar, FertiPro)22, in which the erythrocytes are substituted by latex particles92. For the latex MAR test, a drop of semen is placed on a clean glass slide followed by a drop of antiserum to human immunoglobulin (IgG) and a drop of the sensitized latex particle suspension. Care should be taken that the drops do not touch each other, as this can influence the outcome of the test. The drops are thoroughly mixed with a coverslip and then covered by the same coverslip. The test is read after 10 minutes at room temperature. No interpretation is made if latex agglutinates are not observed. The test is reported as negative if no latex particles are observed bound to motile spermatozoa, doubtful when < 10% of motile sperm have latex particles bound to them, positive if 10–90% of motile spermatozoa show latex particles bound and strongly positive if > 90% of motile spermatozoa show latex particles bound. In all cases of a positive MAR test (> 10% binding), blood and seminal plasma can be obtained for subsequent testing of antisperm antibody titers with the microagglutination93 and immobilization94 tests at a later stage. However, these tests are now also performed very rarely in most modern andrology laboratories, and have been substituted by the Immunobead test for IgA, IgG and IgM (Irvine Scientific, Santa Anna, USA; Laboserv GmbH, Am Boden, Staufenberg, Germany)53. Commercial kits are also available for the direct determination of IgA and IgM antisperm antibodies in semen, although IgM antibodies are seldom found on spermatozoa and the clinical relevance is not clear. IgA, on the other hand, is the antisperm antibody of most clinical importance, as

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spermatozoa coated with IgA antibodies are not capable of penetrating the cervical mucus in vivo, and may be an important reason for long-standing unexplained infertility, which is easily treatable by intrauterine insemination (IUI) of the husband’s washed spermatozoa. Some laboratories perform the IgG and IgA MAR test simultaneously on the semen sample, while others only do the IgA MAR test if the IgG MAR test is positive, as IgA antisperm antibodies are seldom found on their own95. Detection and role of leukocytes Ejaculates usually contain cells other than spermatozoa, called round cells, compiled of, for example, white blood cells and germinal epithelium cells, the latter contributing up to 90% of all round cells in fertile males, with a mean concentration of 0.12 × 106/ml semen, as found by Ariagno et al.96. The inclusion of a test for the identification of granular white blood cells must now be regarded as part of the standard basic routine semen analysis, as the presence of leukocytes is associated with the production of ROS55, causing DNA damage and reduced pregnancy rates with ART33. Many procedures are available for the detection of leukocytes in semen, but, based on the available literature, the leukocyte peroxidase test is indicated as a basic test for this purpose24. Peroxidase test for detecting leukocytes. The solution for performing the leukocyte peroxidase test (Endtz test)97 is prepared by dissolving 125 mg benzidine and 150 mg cyanosine (phloxine) in 50 ml 95% alcohol which is then further diluted with 50 ml distilled water. This solution can be stored in a light-protected bottle. A 3% hydrogen peroxide solution is also prepared. Before the test is performed, 250 µl of the stock solution is mixed with 20 µl of the peroxide solution. For the test itself, one drop of semen is mixed with one drop of the above working solution on a clean glass slide, covered with a coverslip and examined microscopically after 2 minutes, and the number of brown cells per high-power field estimated. Neutrophil granulocytes (leukocytes) stain brown.

Granules of basophil and eosinophil granulocytes stain reddish brown to violet, while lymphocytes and precursors stain light pink, as they are peroxidase-negative22,24. The concentration of peroxidase-positive cells can be counted with the aid of a hemocytometer, and expressed as 106/ml semen. The WHO manual22 suggests that the presence of > 1 × 106 granulocytes/ml semen should be regarded as the presence of leukocytospermia, possibly based on the work of Comhaire et al.98. Other methods for the detection of leukocytes. The suitability of the leukocyte peroxidase test as a screening test for leukocytes has been questioned, and more sophisticated methods have been proposed55. However, it has been demonstrated that polymorphonuclear granulocytes are the most prevalent WBC in semen22,55, and these cells are mainly responsible for the production of ROS99,100. As the leukocyte peroxidase test detects only granular WBC, the procedure can be considered a suitable and reliable routine test for this purpose97. The identification of lymphocytes and monocytes is possible with the aid of monoclonal antibodies against the common leukocyte antigen CD45, whereby granulocytes, lymphocytes and macrophages can be detected, while other monoclonal antibodies allow the selective staining of other WBC subpopulations101–104. According to Wolff 99, immunocytochemistry can be considered the gold standard for the detection of WBC, but these methods are time-consuming and laborintensive, and thus more suitable as a research tool than a routine method. The use of flow cytometry with the aid of monoclonal antibodies can also be considered. Ricci et al.105 described this method as a simple, reproducible procedure, capable of accurately detecting leukocytes in semen and categorizing the different WBC subpopulations without any preliminary purification procedures of the semen samples. Another (indirect) method is the detection of polymorphonuclear leukocyte (PMN) elastase. This enzyme is released by

THE BASIC SEMEN ANALYSIS

activated granulocytes, and can be measured in fresh or frozen seminal plasma. The method is objective, but costly and time-consuming. A strong correlation has been found between elastase levels and WBC numbers in semen106. Esfandiari et al.107 used the nitroblue tetrazolium reduction test for the identification of leukocytes and the assessment of ROS production by leukocytes and spermatozoa. The most basic way of detecting WBC in semen samples is by direct observation of semen smears using bright-field light microscopy with the aid of the Papanicolaou, or the Bryan– Leishman staining108 technique, as discussed in the preceding morphology section. However, the cytological identification of leukocytes and germinal epithelium cells has always been regarded as an insufficient method in much of the literature109. The argument is the inability of most observers to diagnose accurately the various leukocyte subpopulations, or even the inability to distinguish between the different WBC forms and immature germinal epithelium cells. However, positive identification of both groups is possible with a good staining method such as Papanicolaou, although thorough theoretical knowledge, practical training and extensive experience are required78,89–91,110–112. Cut-off values for leukocytospermia. Controversy exists about what can be regarded as leukocytospermia. The WHO manual defines leukocytospermia as the presence of excessive numbers of white blood cells (WBC) or leukocytes in the human ejaculate, which are predominantly granulocytes and more specifically of the neutrophil subtype, and states that in a normal ejaculate the number of WBC should be < 1 × 106/ml21,22. Politch et al.113 already concluded that more research is needed to establish thresholds for pathological levels of WBC in semen for both a mono-antibody-based immunohistological method and the peroxidase method. New cut-off values as low as 0.25 × 106 and even 0.2 × 106 WBC/ml have been proposed88,108. Controversy about leukocytospermia. The fact that the presence of leukocytes in semen may have

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a negative impact on semen parameters and sperm function was addressed as early as 1980 by Comhaire et al.98 and in 1982 by Berger114, and in 1990 by Wolff et al.115. However, in 1992 and 1993, Tomlinson et al. published three articles with an opposite view103,104,116. The first article indicated that seminal leukocytes may play a positive role in male fertility by the removal of morphologically abnormal spermatozoa, the second suggested that the presence of immature germ cells but not leukocytes in semen is associated with reduced success of in vitro fertilization and the third, a prospective study, suggested that leukocytes and leukocyte subpopulations in semen are not a cause for male infertility103,104,116. In 1995, Aitken and Baker concluded that there does not appear to be a convincing case for believing that seminal leukocytes are ‘good Samaritans’. They mentioned that on the other hand leukocytes may often be present without an obvious effect but that it must always be kept in mind that WBC may pose a risk depending on the circumstances that led to their infiltration of the semen sample and that the potentioal of WBC to act as negative terrorists must not be ignored117. Since then, several more articles with opposing views have been published, as well as reports suggesting that inflammation of the male reproductive tract causing leukocytospermia may be a temporary and self-limiting episode, and that this phenomenon is probably common even in fertile males118,119. Matters are further complicated by reports of a very poor relationship between the presence of bacteriospermia and leukocytospermia and male genital tract inflammation120. Comhaire et al. reported that leukocytospermia may be associated with inflammatory reactions of the male genital tract due to the presence of bacteria, and found that ejaculates with > 106 peroxidase positive cells/ml semen contained significantly more pathogenic bacteria isolates, compared with a group of men with < 106 peroxidase-positive cells/ml semen98. Punab et al. also found a positive correlation between the WBC count and the number

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of different micro-organisms, and also between the WBC count and the total count of microorganisms in the semen samples they investigated for both leukocytospermia and bacteriospermia108. In contrast, Rodin et al. found that leukocytospermia was a poor marker for the presence of bacteriospermia121, while Eggert-Kruse found no significant association between leukocytospermia and bacteriospermia122, and neither did Cottell et al.123. The origin of bacteriospermia is still complex. Bacteriospermia usually occurs due to one or more of the following three reasons: (1) normal colonization, (2) contamination of the semen sample or (3) a urogenital infection123. The male genital tract is usually bacteria-free, but the urethra may be colonized by a variety of micro-organisms. It is not clear to what extent these bacteria, which are usually considered as commensal organisms, can contribute to an inflammatory process124. Matters are furthermore complicated due to the possibility of contamination of semen samples by nonpathogenic commensals of the skin or glans penis123. It is therefore not clear to what extent bacteriospermia is indicative of male genital infection per se, and different results for the relationship between leukocytospermia and bacteriospermia have been published, as mentioned above. To add to the controversy, the incidence of leukocytospermia as found in different infertile male populations varies widely from 6.8 to 44.3%125. Influence of leukocytospermia on semen parameters and sperm function. As mentioned in the above paragraph on ‘Controversies about leukocytospermia’, contradictory reports have been published in the literature on the effect of leukocytospermia or even the presence of leukocytes on semen parameters and the functional ability of spermatozoa. For instance, Kaleli et al. found a significant positive correlation between leukocyte counts, as determined using the leukocyte peroxidase test, and increased hypo-osmotic swelling test scores, higher sperm concentrations and enhanced acrosome reactions126. These favorable effects were

especially noted at seminal leukocyte concentrations of between 1 and 3 × 106/ml semen. Kiessling found that semen samples with evaluated concentrations of leukocytes contained a significantly higher frequency of spermatozoa with ideal morphology127. Eggert-Kruse et al. did not find any significant association between the presence of leukocytospermia and the production of antisperm antibodies in semen of the IgA and IgG types as detected using the red blood-cell MAR test122. Neither did Rodin et al. find a negative or positive effect on semen parameters and sperm function in the presence of leukocytospermia121. Many reports on the negative effects of leukocytospermia on sperm function have been published, for instance by Chan et al., who showed that, in the presence of leukocytospermia, hyperactivation of spermatozoa, but not sperm motility, was negatively affected128. Negative correlations between leukocyte concentrations and progressive sperm motility, normal sperm morphology and the hypo-osmotic swelling test have also been reported129, as well as a negative effect on normal sperm morphology, with an increase in the incidence of the stress-related phenomenon of elongated spermatozoa112. From the most recent literature, it is now clear that the main negative effect of leukocytospermia is the production of ROS, causing DNA fragmentation and damage of spermatozoa as detected with the TUNEL (terminal deoxynucleotide transferase-mediated dUTP nick-end labeling) and sperm chromatin structure assays31,33,130,131. Henkel et al. found that DNA fragmentation due to leukocytospermia did not correlate with in vitro fertilization rates, but found a significantly reduced pregnancy rate in IVF and ICSI patients inseminated with spermatozoa for semen samples containing high numbers of TUNEL-positive spermatozoa. This would imply that spermatozoa with damaged DNA are able to fertilize an oocyte, but at the time that the parental genome is switched on, further development of the embryo stops, leading to a lower pregnancy rate31,33. This

THE BASIC SEMEN ANALYSIS

is in agreement with earlier work published by Aitken et al. showing that the incidence of spontaneous pregnancies was negatively correlated with the generation of ROS in a prospective study performed in a group of oligozoospermic patients, where about half the population exhibited increased ROS activity132, and is also confirmed by the work of Fedder110. Therefore, the main negative effect of the presence of leukocytospermia seems to be high ROS production, especially by the WBC but also by spermatozoa themselves, which then causes poor sperm functional ability either by ROS action on the sperm membrane where they interact with polyunsaturated fatty acids or by DNA damage or fragmentation. Source of leukocytospermia. According to Barratt et al. leukocytospermia has a heterogeneous etiology, including infections, inflammations and autoimmunity, making the immediate cause for this condition quite complex and unclear133. In most cases, leukocytes present in semen are presumed to originate from some sort of infection in the male genital tract, but most men with leukocytospermia have negative cultures of samples obtained from the seminal tract134,135. Purvis and Christiansen found that, often, the source of white blood cells in semen is the testicle/epididymis, and that this may be of significance, since spermatozoa are exposed to the potentially damaging influence of leukocytes for much longer periods in the epididymis than in other parts of the tract, leaving more time for DNA damage to occur136. It is thought that in some males the origin of leukocytospermia may be sources outside the genital tract, and a wide range of these factors that may cause leukocytospermia have been reported109,118,120,136–141. Trum et al. reported that leukocytospermia was associated with a history of gonorrhea120. Close et al. found that current cigarette smokers, marijuana users and heavy alcohol users showed a statistically significant greater number of leukocytes in the seminal fluid than did non-users, in a group of 164 men investigated for infertility

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problems142. The increase in round cells and leukocytes in semen samples from smokers was confirmed by a study of Trummer et al.143. It has also been reported that clomiphene citrate treatment of a group of males with low serum testosterone levels may have led to leukocytospermia144. Although it was not correlated with the presence of leukocytospermia, Bieniek and Riedel reported that the same bacteria could be found in the semen samples of men who were diagnosed with bacterial foci in their teeth, oral cavities and jaws, and that after 6 months, following dental treatment in about two-thirds of these men, their semen samples proved to be sterile and the semen parameters such as sperm concentration, motility and morphology had clearly improved, while the semen parameters of the control group remained poor137. Treatment of leukocytospermia. In many reports, antibiotics have routinely been used to treat leukocytospermia, but this is also a controversial matter as several studies have obtained differing results118,134. A meta-analysis of the effectiveness of treatment with broad-spectrum antibiotics of men suffering from leukocytospermia and/or bacteriospermia was performed by Skau and Folstad139. In total, 23 clinical studies were identified, but only 12 studies were included for analysis. Their results indicated that the most used antibiotics were doxycycline, erythromycin and trimethoprim in combination with sulfamethoxazole, and treatment resulted in significant improvements in semen quality. When improvements in the results for different semen parameters were expressed as weighted effect size, the smallest effect was found for sperm concentration, with a mean weighted effect size of 0.16, followed by semen volume and sperm motility, with a mean weighted effect of 0.20, followed by an improvement in normal sperm morphology, with a weighted effect size of 0.22, and the best response to antibiotic treatment was a significant reduction in the concentration of leukocytes in semen samples, with a mean weighted effect size of 0.23.

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A literature survey with emphasis on antibiotic treatment for leukocytospermia only was performed, and 12 articles dealing with the topic were identified110,118,119,145–153. Ten of the articles reported a positive response, that is, a reduction in seminal leukocyte concentrations110,118,145–147,149–153. Some of the articles also reported an improvement in semen parameters, and four110,118,145,153 reported the occurrence of pregnancies as a result of the antibiotic treatment. In a case study of a male with azoospermia, antibiotic treatment for leukocytospermia resulted not only in a decrease of the leukocyte concentration, but also in the appearance of spermatozoa; however, two ICSI treatment cycles were unsuccessful150. Interesting was the observation by Branigan and Muller that higher ejaculation frequencies enhanced the disappearance of leukocytes from semen samples145, and this was confirmed by Yamamoto et al.152. Only two articles reported that significant reductions in the leukocyte concentrations were not obtained119,148. There may be several reasons for not obtaining a positive response with antibiotic treatment for leukocytospermia. One reason may be that different end-points are set for successful treatment results, as illustrated by the two cases found in the literature survey referred to above, reporting a negative result. Although there was a (significant) reduction in seminal leukocyte concentrations after antibiotic treatment, this did not meet the end-point of total eradication of leukocytospermia set by the authors119,148. Other reasons may be as postulated by Purvis and Christiansen, who proposed two reasons for difficulties in showing positive antibiotic treatment-effects in infertile males presenting with leukocytospermia118. The first is that the therapy may not have been appropriate for the organism(s) responsible for the infection, or that the dose or duration may have been inadequate. According to the authors, only certain antibiotics, the most important being ciprofloxacin, have the capacity to penetrate the accessory sex glands in high

enough concentrations. The encouragement of frequent ejaculation during antibiotic treatment is important, as the higher turnover of secretions would be anticipated to encourage passage of the antibiotics into the glandular lumen of affected organs and thereby increase the efficiency of the treatment. The second is that pathological changes in the reproductive tract, due to the presence of infection and responsible for poor semen quality, may become permanent (e.g. epididymal stenosis causing a delay in transit time of spermatozoa or seminiferous tubule failure caused by orchitis). Antibiotic treatment can therefore be expected to have a positive effect on sperm quality only if the chronic infection is still active and the pathological organism is still present, and where the degree of damage is still limited. Eggert-Kruse et al. are of the very forcible opinion that patients with symptoms of genital tract infections (leukocytospermia) should be treated as soon as possible, often as partner therapy, to avoid the severe sequelae of ascending infections102. However, they and others warn strongly that antibiotic treatment should be used with caution and used only when clearly indicated, especially in healthy individuals102,154,155, the reasons being that the non-critical use of antibiotics may result in resistant strains of bacteria, and that certain antibiotics may also have a possible toxic effect on spermatogenesis. The working mechanism of antibiotic treatment in the improvement of semen parameters is not yet quite clear. One mechanism suggested by Skau and Folstad is that antibiotic treatment may cause a reduction in the level of cytotoxic cells present in the testes, causing a reduction in immune activity in the testes, resulting in a higher number of morphologically normal spermatozoa, and less DNA damage139. The effect of treatment can also lead to pregnancies without clear alterations in semen quality but after the disappearance of leukocytes from the ejaculate, possibly because the source of ROS production and thus DNA damage has been eliminated.

THE BASIC SEMEN ANALYSIS

Sperm vital staining test Where previously it was normal procedure to perform a vital staining test on every semen sample with a sperm concentration of > 1.0 × 106/ml, it is now mostly performed in cases with progressive sperm motility of < 30%. The method as described by Eliasson54, based on the method described by Blom156, is generally used. A drop of semen is placed on a spot plate and mixed with one drop of 1% aqueous eosin Y solution. After 15 seconds two drops of 10% aqueous nigrosin solution are added and thoroughly mixed. A drop of this mixture is transferred to a clean glass slide and a thin smear made and air-dried. The smears are examined with a 100× oil magnification. Red cells or any sperm cells not totally white are regarded as dead, and the results are expressed as the percentage of live (white) sperm. It is important to note that, although the staining solutions are referred to as aqueous eosin Y and nigrosin solutions, this refers to the type of eosin Y and nigrosin. Both eosin Y and nigrosin should be dissolved in phosphatebuffered solutions to prevent hypo-osmotic swelling of the sperm tails and the induction of sperm death due to the hypo-osmotic stress caused when the solutions are prepared with water, which thus can give false-negative (low vitality) results157. The performance of a vital stain technique is an important tool to distinguish between live but motionless and dead spermatozoa. Motionless but still alive spermatozoa can be found, for example, in cases of Kartagener’s syndrome, or may be caused by cold shock. In cases where all spermatozoa are found to be dead by vital staining, the condition is called necrozoospermia.

Additional procedures Azoospermia

When examination of the wet preparation indicates that the semen sample contains no spermatozoa, i.e. azoospermia, the following steps are performed. The sample is centrifuged in a conical disposable plastic centrifuge tube for 10 minutes

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at 3000 g. The supernatant is carefully drawn off and discarded, and the pellet suspended in a small amount of medium and re-examined microscopically using phase-contrast optics. The results are interpreted as follows: • No spermatozoa observed = azoospermia; • Spermatozoa present = cryptozoospermia, or sometimes called severe oligozoospermia (< 1 × 106 spermatozoa/ml)39. Moderate oligozoospermia

In cases where the spermatozoa concentration is < 5.0 × 106/ml, the remaining semen after completion of all procedures can be centrifuged at ± 200 g for 10 minutes. The pellet is suspended in a small volume of medium and a small drop used to prepare a standard smear. The smear is air-dried and stained for use in cases where there are too few spermatozoa present on the original morphology smear. Semen biochemistry

Semen biochemistry is not usually performed as part of the standard basic semen analysis procedure. However, in cases of azoospermia, or in the presence of round cells in the wet preparation, certain tests can be performed to aid in the diagnosis of azoospermia, or identification of granular white blood cells when round cells are present. These biochemical tests are usually carried out on seminal plasma obtained by centrifugation of the semen sample in a conical disposable test-tube at 3000 g for 30 minutes. A test for α-glucosidase can be performed to identify a possible obstruction at the site of the epididymis, as this enzyme is produced exclusively by the epididymis158. When the α-glucosidase value is reduced, it can be interpreted that the azoospermia may be due to an obstruction at the level of the epididymis. In cases of azoospermia, the fructose content of the seminal plasma can also be determined, either biochemically or directly on the semen sample as a bench test (as

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discussed below). Fructose is inter alia an indicator of the secretory function of the seminal vesicles, and low fructose levels may indicate congenital dysgenesis (absence) of the seminal vesicles and vas deferens. A kit (Fructose Test; FertiPro) for the spectrophotometric determination of fructose in semen/seminal plasma is commercially available22. In cases where round cells are observed in the wet preparation, a PMN-elastase assay can be performed159. This enzyme is secreted by activated granulocytes, and can be measured in fresh or frozen seminal plasma. The method is objective and convenient, but costly and time-consuming, as 20 samples must be tested at the same time, but a strong correlation has been found between elastase levels and WBC numbers in semen. An increased value of > 290 ng/ml semen is a strong indication of the presence of leukocytes, or a silent inflammation of the genital tract106,160. Colorimetric bench method for fructose determination

A bench method for the quick determination of fructose has been described by Amelar, based on the Selivanoff method161. In this test, 5 mg of resorcinol is added to 33 ml of concentrated hydrochloric acid and diluted to 100 ml with distilled water. An aliquot of 0.5 ml semen is added to 5 ml of the reagent in a heat-resistant glass tube and heated to boiling point. In the presence of fructose, an orange-red coloring will appear within 60 seconds after boiling. Special care should be taken when performing this procedure by wearing protective clothing, especially glasses for protection of the eyes, due to the vicious boiling161. Semen cultures

In all cases where a semen analysis is done for the first time, swabs should be prepared for culturing of aerobic bacteria and for Ureaplasma and Mycoplasma, especially in cases for ART procedures. The patient should be instructed to pass urine and then to wash his hands with soap and

water and the glans penis with water alone. The semen is produced into individually packed and sterilized containers. Unfortunately there is poor correlation between bacteriospermia and leukocytospermia, and the validity of culturing semen samples has been questioned55. It is also evident that it is difficult to culture semen samples, and some laboratories prescribe specialized procedures in order to obtain optimal results22.

Computer-assisted semen analysis The past decade has seen the development of many computer-based systems to analyze semen samples more accurately, objectively and efficiently. Although systems for the measurement of sperm concentration22,39,162,163 and normal morphology164 exist, the motility parameter has received the most attention22,163. These systems have primarily enabled the critical analysis of sperm head kinematics, while flagellar kinematics remains a future challenge. To date, numerous parameters of sperm head motion have been identified, of which 11 have been officially accepted and standardized. Computer-assisted semen analysis (CASA) has also been invaluable in the characterization of hyperactivated motility22. Three of the most generally used CASA parameters are: (1) curvilinear velocity (VCL), i.e. the measure of the rate of travel of the centroid of the sperm head over a given time period (this is calculated from the sum of the straight lines joining the sequential positions of the sperm head along the sperm’s track); (2) straight-line velocity (VSL) (this represents the straight-line distance between the first and last centroid positions for a given time period); (3) linearity of forward progression (LIN) (this is reported as the ratio VSL/VCL expressed as a percentage, and represents the value of 100 cells swimming in a perfectly straight line)22,39,163. In an attempt to standardize CASA results stringent guidelines for the correct operational procedures for CASA systems have been proposed by the ESHRE Andrology Special Interest

THE BASIC SEMEN ANALYSIS

Group165,166. The guidelines include several recommendations, namely, the need for internal and external quality control, adequate training, including that offered by the different manufacturers, and correct operational procedures. The group have advised that in any manuscript or report the technical operational procedures should be clearly spelled out. These should include the image acquisition rate, which is recommended as 50 Hz, tract sampling time, recommended to be a minimum of 0.5 seconds, indication of the type of smoothing algorithm employed, the number of cells sampled, recommended to be > 200 in at least six fields, the type of chamber used, recommended depth 10–20 µm, and also some data on the instrument used, such as the model and software version numbers and microscope optics and magnification22,39,156,166.

Quality control in the andrology laboratory In the modern andrology laboratory, the importance of a distinct quality assurance (QA) policy has become very evident in the past decade, and every laboratory should have such a program in place, including measures for quality control (QC). QA is the larger picture of QC and examines overall laboratory quality, including good laboratory administration of personnel and laboratory procedures, communications skills between all role players and introduction of remedial actions taken when indicated, and documentation of procedures and programs. Detailed descriptions of QA and QC programs and methods can be found in the WHO 1999 manual22, the ESHRE manual53 and various articles165,166 and textbooks39. The 1999 WHO manual22 and the ESHRE manual53 also place great emphasis on especially QC, which must include an internal (IQC) and an external (EQC) leg. The IQC program should include aspects of control of equipment and replicate assessments of the main semen parameters between and within technologists. It can also

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include sampling of monthly averages and other more sophisticated actions such as assessing systematic differences between technicians. There are several EQC programs in different countries and continents run by governments, for example UK NEQAS (UK National External Quality Assessment Service), or national and international society programs such as those of the European Academy of Andrology (EAA) and ESHRE. These programs are limited due to practical logistical difficulties in sending out large numbers of the same samples as well as cost factors. A problem encountered with the different programs is that the standards between them differ, and also they do not all follow the same procedures with regard to standardization. It is known that some national programs use Diff-Quik™stained smears and others Papanicolaou-stained smears, which may give different results. Other problems have been encountered with sperm morphology as demonstrated by Cooper et al.167, indicating that users of the ESHRE EQC program are much more strict in their sperm morphology scoring compared with users of the EAA and UK NEQAS programs. Disagreements between motility grades a and b have also been found between the three schemes. Better standardization can be achieved by continued training of laboratory personnel, as done by the ESHRE Special Interest Group in Andrology with their basic semen analysis training program168, but continued interaction between laboratories and the training facility is also important to assure continued standardization169.

INTERPRETATION OF SEMEN ANALYSIS RESULTS As mentioned previously, a semen analysis result cannot be interpreted correctly unless all factors that may have an influence on the results are known. It is also important when repeat semen analyses are performed and compared with

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previous results that there are a number of factors that may add to the normal biological variation of the results170. Some of them are discussed below.

Sources of variation affecting semen parameters Many sources of variation are known, but only some of those that can cause large variations in semen parameters, such as sexual abstinence, seasonal influences or illness, are discussed briefly. Today, it is an accepted fact that abstinence can have a pronounced but varied effect on the semen parameters. This can vary from a small influence on sperm morphology to a statistical significant effect on sperm motility, sperm concentration and semen volume. This is due to the fact that production of spermatozoa and the secretions of the accessory glands that form the seminal plasma are daily ongoing processes37,40,41,45. It is generally accepted that the human is not a seasonal breeder and that spermatogenesis is a continuous and active process throughout the year. However, a few studies have been reported in which a possible seasonal influence has been investigated171–175. From the literature, it appears that this influence is the result of increased summer temperatures and is mainly an influence on sperm concentration and/or sperm morphology. On the other hand, it has also been speculated that day length rather than temperature may be a reason for seasonal fluctuations175. Henkel et al. found significant seasonal changes in chromatin condensation and sperm count175. Best chromatin-intact values with a mean maximum value of 86.2% aniline blue-negative spermatozoa were found in January, and the highest mean sperm concentration of 68.75 × 106/ml semen was found in April in a group of patients investigated in Germany. For a control group of patients from the Southern hemisphere, a seasonal change shift by 4–5 months was observed for maximum chromatin condensation, but no trend for sperm concentration could be observed.

Mention is often made in articles or chapters on male infertility that a common cold, a bout of influenza or other febrile illnesses will have an adverse effect on spermatogenesis. Therefore, it is important that this is queried in the questionnaire to be completed with every semen analysis1,37. MacLeod published several articles demonstrating the effect of a viral infection with an increased body temperature as well as the effect of chickenpox on semen quality176,177. He found that sperm concentration, motility, forward progression and morphology were all impaired. The same effect was observed by Menkveld and Kruger40,41. The effect can be quite drastic, and is an important factor when evaluating semen analysis results. Two cases presented by Menkveld and Kruger40,41 illustrated that the motility, speed of forward progression and percentage of morphologically normal spermatozoa were the first parameters to show negative effects of the illness. The sperm concentration was not immediately negatively affected, probably due to storage of spermatozoa in the genital tract. This may suggest, therefore, that sperm morphology and movement can be altered while in the genital tract, especially the epididymis65,178. The negative effect of the illness is longest reflected in the sperm morphology, which may indicate that spermatogenesis and spermiogenesis are very sensitive as far as the whole process of morphogenesis is concerned. The adverse environmental effects investigated above seem to have their most pronounced effects on sperm morphology40,41,179. Menkveld et al.40,41,179, like MacLeod14,176,177, came to the conclusion that sperm morphology is a very sensitive parameter that will reflect any adverse influence on the body/testes in a short time. Menkveld and Kruger speculated that any illness or infection might cause a temporary decrease in the percentage of morphologically normal forms, after which it will return to its original value40,41. However, if the testes are repeatedly attacked by adverse influences or conditions, this may start to cause histological changes in the lamina propria and basal membrane or Sertoli cell function, which will

THE BASIC SEMEN ANALYSIS

then adversely influence spermatogenesis. This negative effect will first be reflected in a gradual lowering of the percentage of morphologically normal spermatozoa179,180, with an increase in the percentage of elongated sperm as well as an increase in the number of immature forms. This will then be followed by a decrease in the sperm concentration40,41,180.

Number of semen analyses to be performed Zaneveld and Polakoski45 advocated that if a patient produces a normal sample with the first semen analysis, then there is no need to perform a further semen analysis. However, if the sample is a borderline case or classified as abnormal according to the specific laboratory’s standards, it will be necessary to do more semen analyses before a final diagnosis can be made. In these cases they recommend that three semen analyses, with 3–5 weeks’ intervals, should be carried out. Some authors feel that there will always be some variation from sample to sample, and that it is therefore necessary to perform at least two semen analyses before a diagnosis can be made181–183. Others have stated that several or 3–4 semen analyses over a period of 3 months, representing a complete cycle of spermatogenesis, are required to make an estimation of a patient’s fertility potential18,37,73. An aspect that must now seriously be considered is the cost factor. Due to the increasing costs, the tendency is to keep the number of semen analyses per patient to a minimum. A good policy, therefore, in a case where the first semen sample is classified as normal according to the specific laboratory’s standards, will be to suffice with one semen analysis and to repeat the semen analysis only if indicated due to a long time interval or due to a recent medical event. In a case where the semen analysis is abnormal, the analysis can be repeated two or three times within a period of 3 months so that a good semen profile of the patient can be obtained.

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Interpretation of results The evaluation of a semen specimen must be based on an overall picture that relates seminal volume, spermatozoa concentration, motility and sperm morphology and the results of additional tests such as the MAR test23, leukocyte peroxidase test and biochemical results24. It must be kept in mind that even when results are far below the normal values of a laboratory, conceptions can still occur, although the time to reach this goal may be longer in such cases compared with cases where normal semen values are observed39,184. A distinction should also be made according to the reason the semen analysis was requested. Results of a semen analysis that may establish a poor prognosis for in vivo fertilization may still be adequate for in vitro fertilization. Calculating an index of the total concentration of morphologically normal motile spermatozoa may be of use for in vitro fertilization but is of little relevance for in vivo fertilization, as volume plays an important part in these calculations. It is known that oligozoospermia is frequently associated with a large semen volume, which must be regarded as an abnormal parameter, and this abnormal factor can therefore not be used to calculate such an index and compensate for the low sperm concentration. Large semen volumes are associated with semen loss from the vagina after intercourse, resulting in a large percentage of the available spermatozoa also being lost. Much has been written about interrelationships between semen parameters and the compensating interaction of semen parameters37,185. Although there may be a general tendency186 that high sperm concentrations are associated with higher percentages of motility and normal morphology, Menkveld et al.41,75,180 have shown that there are exceptions, especially as far as sperm morphology is concerned. With regard to the compensating interaction of semen parameters, the above-mentioned argument also holds true. In cases where the volume is within the normal range, a certain degree of compensating interac-

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tion may occur, but this will be limited. It was observed43,73 while calculating normal values and minimal values for conception, based on the occurrence of conceptions in an infertile population (which incidentally should be used and not so-called ‘normal populations’), that a single and consistently very abnormal semen parameter could be associated with no, or only a sporadic, occurrence of conception in apparently normal women43,187.

Standards for normal semen parameters and fertility Normal standards of semen parameters for the basic semen features, i.e. volume, motility, sperm concentration and morphology, have from time to time been published43 and also reviewed in the WHO manuals19–22. In the 1999 WHO manual22, the term ‘normal values’ was changed to ‘reference values’. The values published in the WHO manuals19–22 were mostly obtained through studies done on so-called normal or fertile populations, and were not the lowest values necessary to achieve spontaneous pregnancies. This means that spontaneous pregnancies in normal relationships can also be obtained with lower semen parameter values than those indicated in the manuals. Many authors, especially those not working in the field of andrology, do not take this fact into consideration, and confuse normality with fertility. This results in a situation whereby if the semen parameters (variables) are not within the normal range, as given in the WHO manuals19–22, males are regarded as infertile, i.e. not capable of conception. This can lead to social problems and stress among couples, for example in cases where spontaneous pregnancies actually occur after such a pronouncement has been made. The differences between standards for normality and fertility have been demonstrated by Van Zyl188, Van Zyl et al.73,74 and Menkveld and Kruger41,43. Results of semen analyses of males who had recently impregnated their wives were

classified according to the then internationally accepted normal standards17 by Van Zyl et al.73 and according to the 1987 WHO manual20 normal values by Menkveld and Kruger41,43, and compared with classifications based on the values used for fertility at Tygerberg Hospital189. Van Zyl et al.73 found that only 18.8% of the men were classified as normal or fertile according to the then normal international criteria, as against 68.4% of men according to the Tygerberg values. Menkveld and Kruger41,43 found corresponding values of 20.5% and 64.5%, respectively. The Tygerberg normal values were based on comparison of the values of each separate semen parameter with those in spontaneous pregnancies obtained in a group of apparently normal women attending the infertility clinic at Tygerberg Hospital73,74,187. The lowest value for each semen parameter, above which no significant increase in pregnancy rate per interval group occurred, was taken as the normal value for fertility for each semen parameter. It was found that, based on these semen parameter values, males could be divided into one of three groups, fertile or normal, subfertile and infertile. Fertile is regarded as an optimal chance for spontaneous conception in vivo, subfertile as a reduced chance and infertile as a small chance. These values were found to be also applicable to in vitro fertilization189. Recently, a number of studies have been published in which the semen parameter values of males from so-called fertile populations were compared with the semen parameter values of males from subfertile populations, in order to determine minimum cut-off values for the different semen parameters, to establish a male’s fertility potential190–194. Guzick et al.192, similar to Van Zyl et al.73,74, Menkveld41 and Menkveld and Kruger189, found that men’s fertility potential could be classified into one of three groups based on their semen parameters as possibly fertile or normal, subfertile and infertile. A summary of the proposed values for the different classes as found in the various

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Table 9.1 Cut-off values of semen parameters for the classification of a male’s possible fertility potential, as found in the recent literature, based on a comparison of fertile versus subfertile populations. The Tygerberg Hospital values are based on pregnancies observed Author/semen parameters

Infertile

Subfertile

Ombelet et al.190 concentration (106/ml) progressive motility (%) morphology (% normal) Guzick et al.192 concentration (106/ml) motility (% motile) morphology (% normal)

Fertile

34.0 45.0 10.0 (SC) 13.5–48.0 32.0–63.0 9.0–12.0

> 48.0 > 63.0 > 12.0 (SC)

Günalp et al.193 concentration (106/ml) progressive motility (%) morphology (% normal)

9.0 14.0 5.0

42.0 12.0 (SC)

Menkveld et al.194 motility (% motile) morphology (% normal) morphology (% normal) AI (% normal) TZI (0–4)

20.0 21.0 3.0 3.0 2.09

45.0 31.0 (WHO) 4.0 (SC) 3.0 1.64

2.0–9.9 10.0–29.0 5.0–14.0 < 1.0 and > 6.0

≥ 10.0 ≥ 30.0 ≥ 15.0 1.0–6.0

Tygerberg Hospital values* concentration (106/ml) motility (% motile) morphology (% normal) volume (ml)

< 13.5 < 32.0 < 9.0

< 2.0 < 10.0 < 5.0

Based on publications of Van Zyl69,188, Van Zyl et al.73,74,187, Menkveld41, Menkveld and Kruger40,43,189 and Kruger et al.76,77; AI, acrosome index; TZI, teratozoospermia index; SC, strict Tygerberg Criteria41,75; WHO, 1992 World Health Organization criteria21

studies190,192–194 is presented in Table 9.1. For the Tygerberg classification, the male is categorized based on his poorest semen parameter189. It is believed that in the subfertile group some compensating interaction between the different semen parameters may occur. However, if a specific semen parameter falls in the infertile category, the impairment is so severe that one or even more good semen parameters cannot compensate for the single poor parameter189; nevertheless, even in these cases, a spontaneous in vivo pregnancy is still possible.

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37. Mortimer D. The male factor in infertility. Part I: Semen analysis. Curr Probl Obstet Gynecol Fertil 1985; 8: 4 38. Alexander NJ. Male evaluation and semen analysis. Clin Obstet Gynecol 1982; 25: 463 39. Mortimer D. Practical Laboratory Andrology. Oxford: Oxford University Press 1994 40. Menkveld R, Kruger TF. Basic semen analysis. In Acosta AA, Kruger TF, eds. Human Spermatozoa in Assisted Reproduction, 2nd edn. Carnforth: Parthenon Publishing, 1996: 53 41. Menkveld R. An investigation of environmental influences on spermatogenesis and semen parameters. PhD dissertation (in Afrikaans), Faculty of Medicine, University of Stellenbosch, South Africa, 1987 42. Elzanaty S, Malm J, Giwercman A. Duration of sexual abstinence: epididymal and accessory sex gland secretions and their relationship to sperm motility. Hum Reprod 2005; 20: 221 43. Menkveld R, Kruger TF. Basic semen analysis. In Acosta AA, et al. eds. Human Spermatozoa in Assisted Reproduction. Baltimore: Williams and Wilkins, 1990: 68 44. Freund M. Performance and interpretation of the semen analysis. In Rolands M, ed. Management of the Infertile Couple. Springfield: Charles C Thomas, 1968: 48 45. Zaneveld LJD, Polakoski KL. Collection and the physical examination of the ejaculate. In Hafez ESE, ed. Techniques of Human Andrology. Amsterdam: Elsevier, North-Holland Biomedical Press, 1977: 147 46. Mandal A, Bhattacharyya AK. Studies on the coagulational characteristics of human ejaculates. Andrologia 1985; 17: 80–6 47. Amelar RD. Coagulation, liquefaction and viscosity of human semen. J Urol 1962; 87: 187 48. Mann T. Biochemical appraisal of human semen. In Joël CA, ed. Fertility Disturbances in Men and Women. Basel: Karger, 1971: 146 49. Portnoy L. The diagnosis and prognosis of male infertility: a study of 44 cases with special reference to sperm morphology. J Urol 1946; 48: 735 50. Vermeiden JPW, et al. Pregnancy rate is significantly higher in in vitro fertilization procedure with spermatozoa isolated from nonliquefying semen in which liquefaction is induced by α-amylase. Fertil Steril 1989; 51: 149 51. Tucker M, et al. Use of chymotrypsin for semen preparation in ART. Mol Androl 1990; 2: 179

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52. Hancock P, McLaughlin E. British Andrology Society guidelines for the assessment of post vasectomy semen samples. J Clin Pathol 2002; 55: 812 53. Kvist U, Björndahl L. Manual on Basic Semen Analysis. ESHRE Monographs. Oxford: Oxford University Press, 2002 54. Eliasson R. Supravital staining of human spermatozoa. Fertil Steril 1977; 28: 1257 55. Menkveld R. Leukocytospermia. In Daya S, Harrison RF, Kempers RD, eds. Advances in Fertility and Reproductive Medicine. International Congress Series. Amsterdam: Elsevier, 2004; 1266: 218 56. Goldenberg RL, White R. The effect of vaginal lubricants on sperm motility in vitro. Fertil Steril 1975; 26: 872 57. Carruthers GB. Assessment of the semen. In Philipp EE, Carruthers, GB, eds. Infertility. London: William Heinemann Medical Books, 1981: 195 58. Appell RA, Evans PR. The effect of temperature on sperm motility and viability. Fertil Steril 1977; 28: 1329 59. Makler A. The thickness of microscopically examined seminal samples and its relationship to sperm motility estimation. Int J Androl 1978; 1: 213 60. Folgerí T, et al. Mitochondrial disease and reduced sperm motility. Hum Reprod 1993; 8: 1863 61. Barthelemy C, et al. Tail stump spermatozoa: morphogenesis of defect. An ultrastructural study of sperm and testicular biopsy. Andrologia 1989; 22: 417 62. Eliasson R, et al. The immotile-cilia syndrome. A congenital ciliary abnormality as an etiologic factor in chronic airway infection and male sterility. N Engl J Med 1977; 297: 1 63. Atherton RW. Evaluation of sperm motility. In Hafez ESE, ed. Techniques of Human Andrology. Amsterdam: Elsevier, North-Holland Biomedical Press, 1977: 173 64. MacLeod J, Pazianos A, Ray BS. Restoration of human spermatogenisis by menopausal gonadotrophins. Lancet 1964; 1: 196 65. Purvis K, Brekke I, Tollefsrud A. Epididymal secretory function in men with asthenoteratozoospermia. Hum Reprod 1991; 6: 850 66. Bar-Sagie D, Mayevsky A, Bartoov B. A fluorometric technique for simultaneous measurement of pH and motility in ram semen. Arch Androl 1981; 7: 27 67. Kesserü E, León F. Effect of different solid metals and metallic pairs on human sperm motility. Int J Fertil 1974; 19: 81

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68. Macomber D, Sanders MB. The spermatozoa count. Its value in the diagnosis, prognosis and treatment of sterility. N Engl J Med 1929; 200: 981 69. Van Zyl JA. A review of the male factor in 231 infertile couples. S Afr J Obstet Gynecol 1972; 10: 17 70. Menkveld R, Van Zyl JA, Kotze TJvW. A statistical comparison of three methods for the counting of human spermatozoa. Andrologia 1984; 16: 554 71. Makler A. The improved ten-micrometer chamber for rapid sperm count and motility evaluation. Fertil Steril 1980; 33: 337 72. MacLeod J, Gold RZ. The male factor in fertility and infertility. II. Spermatozoa counts in 1000 men of known fertility and in 1000 cases of infertile marriage. J Urol 1951; 66: 436 73. Van Zyl JA, et al. The importance of spermiograms that meet the requirements of international standards and the most important factors that influence semen parameters. In Proceedings of the 17th Congress of the International Urological Society. Paris: Diffusion Dion Editeurs, 1976; 2: 263 74. Van Zyl JA, et al. Oligozoospermia: a seven-year survey of the incidence, chromosomal aberrations, treatment and pregnancy rate. Int J Fertil 1975; 20: 129 75. Menkveld R, et al. The evaluation of morphological characteristics of human spermatozoa according to stricter criteria. Hum Reprod 1990; 5: 586 76. Kruger TF, et al. Sperm morphologic features as a prognostic factor in in vitro fertilization. Fertil Steril 1986; 46: 1118 77. Kruger TF, et al. Predictive value of abnormal sperm morphology in in vitro fertilization. Fertil Steril 1988; 49: 112 78. Menkveld R, Kruger TF. Evaluation of sperm morphology by light microscopy. In Acosta AA, Kruger TF, eds. Human Spermatozoa in Assisted Reproduction, 2nd edn. Carnforth: Parthenon Publishing, 1996: 109 79. Mortimer D, Menkveld R. Sperm morphology assessment – historical perspectives and current opinions. J Androl 2001; 22: 192 80. Kruger TF, et al. A quick, reliable staining technique for human sperm morphology. Arch Androl 1987; 18: 275 81. Oettlé EE. Using a new acrosome stain to evaluate sperm morphology. Vet Med 1986; 81: 263 82. Menkveld R. Appendices. In Menkveld R, et al., eds. Atlas of Human Sperm Morphology. Baltimore: Williams & Wilkins, 1991: 115

83. Menkveld R, et al. Effect of different staining and washing procedures on the results of human sperm morphology evaluation by manual and computerised methods. Andrologia 1997; 29: 1 84. Menkveld R, et al. Basic principles and practical aspects. In Menkveld R, et al., eds. Atlas of Human Sperm Morphology. Baltimore: Williams & Wilkins, 1991: 6 85. Menkveld R, et al. Acrosomal morphology as a novel criterion for male fertility diagnosis: relation with acrosin activity, morphology (strict criteria) and fertilization in vitro. Fertil Steril 1996; 65: 637 86. Cooper TG, et al. Cytoplasmic droplets are normal structures of human sperm but are not well preserved by routine procedures for assessing sperm morphology. Hum Reprod 2004; 19: 2283 87. Cooper TG. New debate. Cytoplasmic droplets: the good, the bad or just confusing? Hum Reprod 2005; 20: 9 88. Menkveld R, Kruger TF. Sperm morphology and male urogenital infections. Andrologia 1998; 30 (Suppl 1): 49 89. Johanisson E, et al. Evaluation of ‘round cells’ in semen analysis: a comparative study. Hum Reprod Update 2000; 6: 404 90. Riedel H-H, Semm K. Leucospermia and male fertility. Arch Androl 1980; 5: 51 91. Riedel H-H. Techniques for the detection of leukocytospermia in human semen. Arch Androl 1980; 5: 287 92. Mahmoud A, Comhaire F. Debate continued. Antisperm antibodies. Use of the mixed agglutination reaction (MAR) test using latex beads. Hum Reprod 2000; 15: 231 93. Friberg J. A simple and sensitive micro-method for demonstration of sperm-agglutinating activity in serum from infertile men and women. Acta Obstet Gynecol Scand 1974; 36 (Suppl): 21 94. Isojima S, Shun T, Ashitaka Y. Immunologic analysis of sperm-immobilizing factor found in sera of women with unexplained sterility. Am J Obstet Gynecol 1968; 101: 677 95. Menkveld R, et al. Detection of sperm antibodies on unwashed spermatozoa with the immunobead test: a comparison of results with the routine method and seminal plasma TAT and SCMC test. Am J Reprod Immunol 1991; 25: 88 96. Ariagno J, et al. Shedding of immature germ cells. Arch Androl 2002; 48: 127 97. Endtz AW. A rapid staining method for differentiating granulocytes from ‘germinal cells’ in Papanicolaou-stained semen. Acta Cytol 1974; 18: 2

THE BASIC SEMEN ANALYSIS

98. Comhaire F, Verschraegen G, Vermeulen L. Diagnosis of accessory gland infection and its possible role in male infertility. Int J Androl 1980; 3: 32 99. Wolff H. Methods for the detection of male genital tract inflammation. Andrologia 1998; 30 (Suppl 1): 35 100. Shekarriz M. Positive myeloperoxidase staining (Endtz test) as an indicator of excessive reactive oxygen species formation in semen. J Assist Reprod Genet 1995; 12: 70 101. Wolff H, Anderson DJ. Immunohistologic characterization and quantitation of leukocyte subpopulations in human semen. Fertil Steril 1988; 49: 497 102. Eggert-Kruse W, et al. Differentiation of round cells by means of monoclonal antibodies and relationship with male fertility. Fertil Steril 1992; 58: 1046 103. Tomlinson MJ, et al. The removal of morphologically abnormal sperm forms by phagocytes: a positive role for seminal leukocytes? Hum Reprod 1992; 10: 517 104. Tomlinson MJ, et al. Round cells and sperm fertilizing capacity: the presence of immature germ cells but not seminal leukocytes are associated with reduced success of in vitro fertilization. Fertil Steril 1992; 58: 1257 105. Ricci G, et al. Leukocyte detection in human semen using flow cytometry. Hum Reprod 2000; 15: 1329 106. Ludwig M, et al. Evaluation of seminal plasma parameters in patients with chronic prostatitis or leukocytospermia. Andrologia 1998; 30 (Suppl 1): 41 107. Esfandiari N, et al. Utility of nitroblue tetrazolium test for assessment of reactive oxygen species production by seminal leukocytes and spermatozoa. J Androl 2003; 24: 862 108. Punab M, et al. The limit of leukocytospermia from the microbiological viewpoint. Andrologia 2003; 35: 271 109. Wolff H. The biological significance of white blood cells in semen. Fertil Steril 1995; 63: 1143 110. Fedder J. Nonsperm cells in human semen: with special reference to seminal leukocytes and their possible influence on fertility. Arch Androl 1996; 36: 41 111. Leib Z, et al. Reduced semen quality caused by chronic abacterial prostatitis: an enigma or reality? Fertil Steril 1994; 61: 1109 112. Menkveld R, et al. Morphological sperm alternations in different types of prostatitis. Andrologia 2003; 35: 288

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113. Politch JA, et al. Comparison of methods to enumerate white blood cells in semen. Fertil Steril 1993; 60: 372 114. Berger RE, et al. The relationship of pyospermia and seminal fluid bacteriology to sperm function as reflected in the sperm penetration assay. Fertil Steril 1982; 37: 557 115. Wolff H, et al. Leukocytospermia is associated with poor semen quality. Fertil Steril 1990; 53: 528 116. Tomlinson MJ, Barratt CLR, Cooke ID. Prospective study of leukocytes and leukocytes subpopulations in semen suggest they are not a cause of male infertility. Fertil Steril 1993; 60: 1069 117. Aitken RJ, Baker HWG. Seminal leukocytes: passengers, terrorists or good Samaritans? Hum Reprod 1995; 10: 1736 118. Purvis K, Christiansen E. Infection in the male reproductive tract. Impact, diagnosis and treatment in relation to male fertility. Int J Androl 1993; 16: 1 119. Yanushpolsky EH, et al. Antibiotic therapy and leukocytospermia: a prospective, randomised, controlled study. Fertil Steril 1995; 63: 142 120. Trum JW, et al. Value of detecting leukocytospermia in the diagnosis of genital tract infection in subfertile men. Fertil Steril 1998; 70: 315 121. Rodin DM, Larone D, Goldstein M. Relationship between semen cultures, leukospermia, and semen analysis in men undergoing fertility evaluation. Fertil Steril 2003; 79 (Suppl 3): 1555 122. Eggert-Kruse W, et al. Induction of immunoresponse by subclinical male genital tract infection? Fertil Steril 1996; 65: 1202 123. Cottell E, et al. Are seminal fluid microorganisms of significance or merely contaminants? Fertil Steril 2000; 74: 465 124. Willén M, et al. The bacterial flora of the genitourinary tract in healthy fertile men. Scand J Urol Nephrol 1996; 30: 387 125. Omu AE, et al. Seminal immune response in infertile men with leukocytospermia: effect on antioxidant activity. Eur J Obstet Gynecol Reprod Biol 2000; 86: 195 126. Kaleli S, et al. Does leukocytospermia associate with poor semen parameters and sperm function in male fertility? The role of different seminal leukocytes concentrations. Eur J Obstet Gynecol Reprod Biol 2000; 89: 185 127. Kiessling AA, et al. Seminal leukocytes: friends or foes? Fertil Steril 1995; 64: 196 128. Chan PJ, et al. White blood cells in semen affect hyperactivation but not sperm membrane integrity

168

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130.

131.

132.

133.

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136. 137.

138.

139.

140.

141.

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in the head and tail regions. Fertil Steril 1994; 61: 986 Arata de Bellabarba G, et al. Nonsperm cells in human semen and their relationship with semen parameters. Arch Androl 2000; 45: 131 Aitken RJ, et al. Differential contribution of leukocytes and spermatozoa to generation of oxygen species in the ejaculates of oligozoospermic patients and fertile donors. J Reprod Fertil 1992; 94: 451 Alvarez JG, et al. Increased DNA damage in sperm from leukocytospermic semen samples as determined by the sperm chromatin structure assay. Fertil Steril 2002; 78: 319 Aitken RJ, Irvine DS, Wu FC. Prospective analysis of sperm oocyte fusion and reactive oxygen species generation as criteria for the diagnosis of infertility. Am J Obstet Gynecol 1991; 164: 542 Barratt CLR, Bolton AE, Cooke ID. Functional significance of white blood cells in the male and female reproductive tract. Hum Reprod 1990; 5: 639 Anderson DJ. Should male infertility patients be tested for leukocytospermia? Fertil Steril 1995; 63: 246 Hillier SL, et al. Relationship of bacteriologic characteristics to semen indices in men attending an infertility clinic. Obstet Gynecol 1990; 75: 800 Purvis K, Christiansen E. The impact of infection on sperm quality. J Br Fertil Soc 1995; 1: 31 Bieniek KW, Riedel H-H. Bacterial foci in teeth, oral cavity, and jaw – secondary effects (remote action) of bacterial colonies with respect to bacteriospermia and subfertility in males. Andrologia 1993; 25: 159 Busolo F, Zanchetta R, Cusinato R. Microbial flora in semen of asymptomatic infertile men. Andrologia 1984; 16: 269 Skau PA, Folstad I. Do bacterial infections cause reduced ejaculate quality? A meta-analysis of antibiotic treatment of male infertility. Behavioral Ecol 2003; 14: 40 Eggert-Kruse W, et al. Antisperm antibodies and microorganisms in genital secretions – a clinical significant relationship? Andrologia 1998; 30 (Suppl 1): 61 Mi´c i´c S, Petrovic S, Dotli´c R. Seminal antisperm antibodies and genitourinary infections. Urology 1990; 35: 54 Close CE, Roberts PA, Berger RE. Cigarettes, alcohol and marijuana are related to pyospermia in infertile men. J Urol 1990; 144: 900

143. Trummer H, et al. The impact of cigarette smoking on human semen parameters and hormones. Hum Reprod 2002; 17: 1554 144. Matthews GJ, Goldstein M, Schlegel HJM. Nonbacterial pyospermia: a consequence of clomiphene citrate therapy. Int J Fertil Menopaus Stud 1995; 40: 187 145. Branigan EF, Muller CH. Efficacy of treatment and recurrence rate of leukocytospermia in infertile men with prostatitis. Fertil Steril 1994; 62: 580 146. Branigan EF, Spadoni LR, Muller CH. Identification and treatment of leukocytospermia in couples with unexplained infertility. J Reprod Med 1995; 40: 625 147. Erel TE, et al. Antibiotic therapy in men with leukocytospermia. Int J Fertil 1997; 42: 206 148. Krisp A, et al. Treatment with levofloxacin does not resolve asymptomatic leukocytospermia – a randomised controlled study. Andrologia 2003; 35: 244 149. Maruyama DK, et al. Effects of white blood cells on the in vitro penetration of zona-free hamster eggs by human spermatozoa. J Androl 1985; 6: 127 150. Montag M, Van der Ven H, Haidl G. Recovery of ejaculated spermatozoa for intracytoplasmic sperm injection after anti-inflammatory treatment of an azoospermic patient with genital tract infection: a case report. Andrologia 1999; 31: 179 151. Malallah YA, Zissis NP. Effect of minocycline on the sperm count and activity in infertile men with high pus cell counts in their seminal fluid. J Chemother 1992; 4: 286 152. Yamamoto M, et al. Antibiotic and ejaculation treatments improve resolution rate of leukocytospermia in infertile men with prostatitis. Nagoya J Med Sci 1995; 58: 41 153. Ekwere PD, Etuk EH. Semen quality among subfertile males following treatment with ofloxacin. Curr Ther Res 1991; 50: 425 154. Bar-Chama N, Goluboff E, Fisch H. Infection and pyospermia in male infertility. Is it really a problem? Urol Clin North Am 1994; 21: 469 155. Schlegel PN, Chang TSK, Marshall FF. Antibiotics: potential hazard to male fertility. Fertil Steril 1991; 55: 235 156. Blom E. A one-minute live–dead sperm stain by means of eosin–nigrosin. Fertil Steril 1950; 1: 176 157. Björndahl L, et al. Andrology Lab Corner. Why the WHO recommendations for eosin–nigrosin staining techniques for human sperm vitality assessment must change. J Androl 2004; 25: 671

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158. Comhaire F, et al. Why do we continue to determine α-glucosidase in human semen? Andrologia 2002; 34: 8 159. Maegawa M, et al. Concentration of granulocyte elastase in seminal plasma is not associated with sperm motility. Arch Androl 2001; 47: 31 160. Blenk H, Hofstetter A. Complement C3, coeruloplasmin and PMN-elastase in the ejaculate in chronic prostato-adnexitis and their diagnostic value. Infections 1991; 19 (Suppl 3): 138 161. Amelar RD. Infertility in Men. Philadelphia: FA Davies, 1966: 13 162. Knuth UA, Yeung C-H, Nieschlag E. Computerized semen analysis: objective measurement of semen characteristics is biased by subjective parameter setting. Fertil Steril 1987; 48: 118 163. Mortimer D, Goel N, Shu MA. Evaluation of the CellSoft automated semen analysis system in a routine laboratory setting. Fertil Steril 1988; 50: 960 164. Kruger TF, et al. A new computerized method of reading sperm morphology (strict criteria) is as efficient as technician reading. Fertil Steril 1993; 59: 202 165. ESHRE Andrology Special Interest Group. Guidelines on the application of CASA technology in the analysis of spermatozoa. Hum Reprod 1998; 13: 142 166. ESHRE Andrology Special Interest Group. Consensus workshop on advanced diagnostic andrology techniques. Hum Reprod 1996; 11: 1463 167. Cooper TG, et al. Semen analysis and external quality control schemes for semen analysis need global standardization. Int J Androl 2002; 25: 306 168. Björndahl L, et al. ESHRE basic semen analysis courses 1995–1999: immediate beneficial effects of standardized training. Hum Reprod 2002; 17: 1299 169. Franken DR, et al. Monitoring technologist reading skills in a sperm morphology quality control program. Fertil Steril 2003; 79 (Suppl 3): 1637 170. Álvarez C, et al. Biological variation of seminal parameters in health subjects. Hum Reprod 2003; 18: 2082 171. Mortimer D, et al. Annual patterns of human sperm production and semen quality. Arch Androl 1983; 10: 1 172. Tjoa WS, et al. Circannual rhythm in human sperm count revealed by serially independent sampling. Fertil Steril 1982; 38: 454 173. Saint Pol P, et al. Circannual rhythms of sperm parameters of fertile men. Fertil Steril 1989; 51: 1030

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174. Rojansky N, Brzezinski A, Schenker JG. Seasonality in human reproduction: an update. Hum Reprod 1992; 7: 735 175. Henkel R, et al. Seasonal changes in human sperm chromatin condensation. J Assist Reprod Genet 2001; 18: 371 176. MacLeod J. The clinical implications of deviations in human spermatogenesis as evidenced in seminal cytology and experimental production of these deviations. In Proceedings of the Fifth Congress on Fertility and Sterility, Stockholm, June 16–22. Excerpta Medica International Congress Series. 1966; 133: 563 177. MacLeod J. Effect of chickenpox and of pneumonia on semen quality. Fertil Steril 1951; 2: 523 178. Fredricsson B. On the development of different morphologic abnormalities of human spermatozoa. Andrologia 1978; 10: 43 179. Menkveld R, et al. Possible changes in male fertility over a 15-year period. Arch Androl 1986. 17: 143 180. Menkveld R, Kotze TJvW, Kruger TF. Relationship of human sperm morphology with other semen parameters as seen in different reference populations. Hum Reprod 1992; 6 (Suppl 1): 96 181. Mehan DJ, Chehval MJ. A clinical evaluation of a new silastic seminal fluid collection device. Fertil Steril 1977; 28: 689 182. Amelar RD, Dubin L, Schoenfeld C. Semen analysis: an office technique. Urology 1973; 2: 605 183. Taylor PJ, Martin RH. Semen analysis in the investigation of infertility. Can Fam Phys 1981; 27: 113 184. Jouannet P, et al. Male factors and the likelihood of pregnancy in infertile couples. 1. Study of sperm characteristics. Int J Androl 1988; 11: 379 185. MacLeod J, Gold RZ. The male factor in fertility and infertility. VIII. A study of variation in semen quality. Fertil Steril 1952; 7: 387 186. MacLeod J. A possible factor in etiology of human male infertility. Fertil Steril 1962; 13: 29 187. Van Zyl JA, Kotze TJvW, Menkveld R. Predictive value of spermatozoa morphology in natural fertilization. In Acosta AA, et al., eds. Human Spermatozoa in Assisted Reproduction. Baltimore: Williams & Wilkins, 1990: 319 188. Van Zyl JA. The infertile couple. Part II. Examination and evaluation of semen. S Afr Med J 1980; 57: 485 189. Menkveld R, Kruger TF. Basic semen analysis: the Tygerberg experience. In Acosta AA, et al., eds. Human Spermatozoa in Assisted Reproduction. Baltimore: Williams & Wilkins, 1990: 164

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190. Ombelet W, et al. Semen parameters in a fertile versus infertile population: a need for change in the interpretation of semen testing. Hum Reprod 1997; 12: 987 191. Zinaman MJ, et al. Semen quality and human fertility: a prospective study with healthy couples. J Androl 2000; 21: 145 192. Guzick DS, et al. Sperm morphology, motility and concentration in fertile and infertile men. N Engl J Med 2001; 345: 1388

193. Günalp S, et al. A study of semen parameters with emphasis on sperm morphology in a fertile population: an attempt to develop clinical thresholds. Hum Reprod 2001; 16: 110 194. Menkveld R, et al. Semen parameters including WHO and strict criteria morphology, in a fertile and subfertile population: an effort towards standardisation of in vivo thresholds. Hum Reprod 2001; 16: 1165

10 Advances in automated sperm morphology evaluation Kevin Coetzee, Thinus F Kruger

INTRODUCTION

can be attributed to the subjective nature of evaluation and methodological inconsistencies. Despite the lack of confidence in the manually evaluated sperm morphology outcomes, the majority of clinics persist in the use of the standard, manually evaluated semen analysis1,10. Automated systems have the power to increase the objectivity, precision and reproducibility of sperm morphology evaluations, and add further value by providing accurate sperm kinematics measures. As attractive as this option may seem, not many automated systems have been introduced into routine andrology laboratories. The majority of systems currently in operation are used in more experimental situations, because of the objective biological resolution of the systems. The probable reasons for the resistance to routine application of the systems are: (1) the cost of the systems, (2) technical limitations of some of the systems (software and hardware) and (3) the limited number of technical and clinical studies published per system to prove their value11. Only through continued demonstration of the value of objective automated semen analysis outcomes in relation to fertility in large prospective randomized studies will the incentive increase to introduce automated systems into routine andrology laboratories12.

Normal sperm morphology has been shown to be predictive of male fertility, independent of other semen parameters. Two literature surveys were conducted to assess this value, both confirming the superior value of percentage normal sperm morphology, as compared with any other manually evaluated semen parameter1,2, when evaluated using standardized methodology under controlled conditions. In humans, normal fertile ejaculates contain spermatozoa exhibiting considerable morphological variations not only in the size and shape of the head and the acrosome, but also in the degree of nuclear vacuolation, size of persisting cytoplasmic droplets, midpiece disturbances and tail abnormalities1. Since 1950 many investigators have tried to create a standardized set of criteria for the assessment of human sperm morphology3–9. The major shortcoming underlying the universal acceptance of any of these criteria and/or guidelines has been the large interobserver, intraobserver and interlaboratory coefficients of variation observed. The value of manually evaluated sperm morphology outcomes has been questioned by many, owing to the lack of precision and reliability observed. Most of the variation inherent to manual evaluations

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AUTOMATED SYSTEMS Although this chapter’s focus is on automated systems, manual techniques and semiautomatic systems have been developed that can also be classified as objective systems. These techniques and systems are important in that they are often simple and economical to set up and use. Calamera et al.13 modified and described a manual method using only a video camera, monitor and microscope. An acetate overlay mask of normal sperm morphology was created by three independent observers using World Health Organization (WHO) 199214 guidelines and strict criteria. Similarly, Goulart et al.15 in a comparative study (manual vs. semiautomatic vs. automatic) developed a manual system in which the operator controlled all the settings (strict criteria) and the evaluation procedure, using a computer mouse. Semiautomatic methods for classifying human sperm based on objective measurements of head shapes and sizes have also been developed15,16, in which the operator can interactively control the evaluation procedure. In the study by Goulart et al.15, the semiautomatic system was found to be the most reliable and secure method for performing sperm analysis, as such a system allowed the operator to confirm or correct possible computer misidentification. Although these systems have demonstrated a certain degree of accuracy and reliability in the evaluation of sperm morphology, the limitation is the time required per evaluation. True automated systems consist of a microscope, a video camera, a computer, a frame grabber and morphology software. The systems work as follows. The video camera delivers the image (digitization) to the frame grabber, which stores it for analysis, and the image is evaluated by the morphology software and included for statistical analysis. Recognition of spermatozoa and exclusion of other cells depend on the software specifications (gates) for sperm shape, size and color (stain) intensity. Once spermatozoa are recognized

and separated from debris and other cells, metric measurements are performed on the sperm head, midpiece, acrosome and other cytological features. The software is normally programmed to recognize spermatozoa according to dimensions and criteria required by the authors17. These may depend on the staining procedure used (e.g. Papanicolaou vs. Diff-Quik) and the range of values of the classification systems used (e.g. strict criteria, WHO guidelines, biological selection criteria, etc.). Many variations (hardware and/or software) of the above configuration can be developed to eliminate a weakness and/or exploit a strong point (Table 10.1). Sofitikis et al.18 used a confocal laser scanning microscope instead of a normal light microscope to evaluate sperm morphology quantitatively. They were therefore able to use unstained semen samples to define the normal ranges of sperm morphometric parameters, to exclude the effect of the staining procedure. The initial systems relied only on morphometric measurements to classify spermatozoa into groups. Evaluation precision was improved by the Hamilton Thorne Research integrated visual optical system (IVOS), which introduced the signature method of including evaluation of the sperm head shape, shown to be of most clinical significance19,32. The ability of the systems to evaluate shape is important, because the correct cell head aspect ratio does not always guarantee normality. Other systems have also incorporated shape analysis methods in their evaluation procedure, for example the Hobson Sperm Tracker20. The sperm head automated morphometric analysis system (SHAMAS) used by Garrett et al.33 included another classification parameter, %Z: the percentage of sperm with characteristics which conform to those of sperm that bind to the zona pellucida of the human oocyte. These ‘zona pellucida preferred’ values indicate axial symmetry, narrow neck and large acrosomal area as important for sperm–zona binding, and therefore normal fertilizing potential.

Coetzee et al.17 Kruger et al.19,21,22 Laquet et al.23 Menkveld et al.24

Davis et al.12, Davis and Gravance25

MacLeod and Irvine26

Wang et al.27,28

Mundy et al.29

Garret and Baker30

Sofikitis et al.18

El-Ghobashy and West20

Goulart et al.15

FERTECH SMA

CellForm-Human

HDATA

Morphologizer II

MOP-Videoplan

Microsoft Professional Basic 7.0

Hobson Sperm Tracker

Zeiss imageprocessing system KS400 (Zeiss-Vision)

Zeiss, Germany

Hobson Tracker United, Sheffield, UK

Microsoft Corp., Redmond, WA

Kontron

Cryo Resources Ltd

Pyramid Technical Consultants, Waltham, MA

Microsoft Corp., Bellview, WA

FERTECH, Norfolk, VA

Company

Criteria and measurements

Strict criteria: head size and shape (signature method) and acrosome size

WHO 1987: length, width, area, perimeter and width/length ratio WHO 1987: length, width and area

WHO 1987: area, perimeter, length/width ratio, roundness, length and width NG: area, perimeter, head maximum diameter, head width, midpiece width, midpiece length and tail length WHO 1992: set of 32 morphometric parameters (size, shape and staining heterogeneity) ± 2SD of fertile men: length, width, length of midpiece, length of principle piece of sperm tail WHO 1999: head length 4–5 µm, width 2.5–3.5 µm, length/width ratio 1.5–1.75 and acrosome size 40–70% of total, including tail and acrosomal vacuoles Strict criteria, head size and shape

Instrument

IVOS, Hamilton Thorne Research, Beverly, MA

Combination system HTM-S 2030, Hamilton Thorne Research, Beverly, MA Combination system Combination system (SEM) Combination system Confocal scanning laser microscope, Lasertec, Yokohama, Japan Combination system

Combination system

NG, not given; SEM, scanning electron microscope; WHO, World Health Organization

Authors

Automated sperm morphology analyzers. Modified from reference 31

Software

Table 10.1

ADVANCES IN AUTOMATED SPERM MORPHOLOGY EVALUATION

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SLIDE PREPARATION AND STAINING In a world-wide survey conducted by Ombelet et al.34, it was confirmed that a wide variety of different methodologies were being followed for the evaluation of sperm morphology. The adopted and adapted methods included procedures for the preparation of semen samples and staining of sperm cells, as well as classification systems used to identify normal and abnormal cells. Ombelet et al.34 concluded that an urgent need to standardize sperm morphology evaluation methodology existed. Just as in the case of the visual evaluation of sperm morphology, users of computer-assisted sperm morphology analyzers must recognize that the principles of standardization and quality control are paramount to accurate evaluations35. Sample preparation and staining may significantly influence the precision and reliability of sperm morphology evaluations. The variation that may result from these procedures can to a large extent be overcome by an experienced technician using visual evaluation, but this may not be possible when an automated system is used24. By the very nature of the evaluation process in automated systems, there is no means of compensating for preparation defects and artifacts. For example, small differences in background shading relative to cell staining intensity can result in digitization errors, leading to incorrect classification or the inability to identify the cell as a sperm. Davis and Gravance25 found that the percentage of normal sperm detected by the CellFormHuman method was not different for washed specimens compared with unwashed controls. The technical variability arising from semen preparation and slide staining methods could, however, be reduced when specimens were washed and resuspended to a standard concentration (150–200 × 106) before smearing. Lacquet et al.23 also preferred using washed semen samples resuspended at a concentration of 100 × 106 cells/ml. Thin, evenly spread smears were made from this solution to ensure that approximately five cells were available per screen for analysis. It is now

preferred practice to prewash the semen sample and to adjust the concentration of the resultant sperm sample. A single- or double-wash procedure can be followed. If a single wash is performed the sample must be adequately diluted (≥ 1 : 5, semen/ medium) prior to centrifugation. Washing the semen sample may be essential for two reasons: (1) to remove as much of the acellular constituents (plasma) of the semen as possible and (2) to concentrate the sperm sample11. The presence of a high concentration of seminal plasma results in intense background staining and flaking during the staining procedure. A droplet, its size depending on the concentration of the resultant sperm sample, must be thinly smeared across a clean slide and allowed to air-dry (room temperature). This capability of being able to adjust the concentration of sample is especially important for oligozoospermic samples. The sample processing procedure must result in between 10 and 20 sperm per high-field magnification (5–10 sperm per computer screen) to optimize the reading time. The density of sperm required for automatic evaluations is therefore double that required for manual evaluations. The most commonly used stains or staining methods used for the evaluation of sperm morphology are hematoxylin stain, the Papanicolaou method, the Shorr method, the Spermac method or the Diff-Quik method. Morphometric measurements were found to be more accurate and precise when sperm were stained with GZIN than when stained with Papanicolaou or hematoxylin25. Lacquet et al.23 found no statistical difference in outcome between five different Diff-Quik (Hemacolor Kit, Merck) staining procedures. Menkveld et al.24, in a study comparing the effect of washing and staining methods (Papanicolaou, Shorr, DiffQuik and Spermac) on automated evaluation, obtained results comparable to manual evaluation by washing the semen samples once and staining with Diff-Quik stain. Wang et al.27, using a simplified Shorr staining procedure, found that less shrinkage of the spermatozoa occurred compared with the Papanicolaou staining procedure,

ADVANCES IN AUTOMATED SPERM MORPHOLOGY EVALUATION

resulting in higher length, width and length/width ratio means. Different staining procedures therefore result in different chromatic and physical appearances of sperm cells. This is certainly true for the Papanicolaou and Diff-Quik staining methods36. Dimension-specific software (Papanicolaou and Diff-Quik) has therefore been loaded into the Hamilton Thorne Research (IVOS) system. A study was hence conducted to determine the agreement between computer-analyzed normal sperm morphology values (n = 97) stained according to the Papanicolaou and Diff-Quik methods17. A significant bias of 1.6% was obtained in favor of higher normal sperm morphology percentages when the Diff-Quik method was used. One of these two methods had to be selected to standardize methodology for future automated evaluation studies. The Diff-Quik staining procedure was selected as the preferred staining method, because of its simplicity, short staining time and good contrast. This difference seen when using different stains also illustrates the importance of ensuring that the software program is developed according to the method of cell staining used. These results illustrate the importance of the standardization of procedures, and of selecting procedures that will result in optimal cell recognition and evaluation. The requirements are: thin, evenly spread smears (five sperm cells per screen) to ensure that all sperm are on the same focus plane, and a staining procedure that ensures minimal background staining, good contrast and good color differentiation. The reproducible production of high-quality slides will ensure that the time required to carry out normal sperm morphology evaluations is kept to the minimum.

EVALUATION PRECISION The manual evaluation of sperm morphology still continues, resulting in inaccurate and valueless measures, even though a better alternative exists. The coefficients of variation for repeat estimates

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by manual evaluation of normal sperm have been observed to be as high as 100% within and between laboratories. The average coefficients of variation for most laboratories are probably in the range 30–60%37. This high possible level of variation may no longer be acceptable, with increasing pressure for laboratories to implement strict quality control programs and be accredited according to the guidelines and conditions of accreditation bodies. If automated systems represent the only alternative, the question would have to be whether the available versions have reached the level of precision acceptable for routine implementation. Although inaccurate and imprecise, the visual evaluation of sperm morphology provides the only practical standard with which to compare the outcomes of automated evaluations of normal sperm morphology. For these systems to be accepted they must first demonstrate coefficients of variation smaller in magnitude than those obtained for visual evaluations. The strict criteria are unique in that the underlying philosophy of the classification system limits variation in the evaluation of sperm morphology. This is clearly illustrated by the study performed by Menkveld et al.32, in which relatively low coefficients of variation were obtained for repeat manual evaluations by experienced technicians, ranging between 5.21 and 27.76%. The goal should therefore be to develop systems that will produce coefficients of variation of < 10%. Davis et al.12, measuring the same sperm repeatedly by computer, obtained a < 1% overall coefficient of variation for repeated measures. In their study, Kruger et al.19 analyzed 255 cells three times in succession and obtained pairwise agreements of 0.85, 0.80 and 0.85 (K statistic > 0.75, i.e. excellent agreement). Davis et al.12 also partitioned the variance among other factors and obtained the following coefficients of variation: between men 1.84–4.17%, between slides 0.6–1.38%, between repetitions 0.16–1.10% and between sperm 6.59–11.39%. Sperm morphology outcomes, as determined by an automated system using stained smears from washed samples, was

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shown in a study by Garrett and Baker30 to have a coefficient of variation equaling < 4% for the same semen sample and < 7% with different batches of stain. The authors concluded that such results are superior to those of an experienced technician using manual evaluations. The average intraslide (three repeat measures) coefficient of variation for the automated evaluations of 100 cells and 200 cells was found to be 9.73% and 8.30%, respectively, when using the IVOS. The average interslide coefficient of variation obtained using the IVOS was, however, 15.39%38. The approximately 6–7% higher variation obtained for interslide evaluations, as compared with intraslide evaluations, may once again point to the importance of sample and slide preparation. The average coefficient of variation for repeat evaluations is known to be a function of both the number of sperm evaluated and the percentage of normal forms25,39, due to statistical presuppositions. Semen samples with low percentages of normal sperm (< 10%) will inherently exhibit higher variability in repeat analyses. Davis and Gravance25 concluded that at least 200 cells should be evaluated to obtain a stable estimate of the percentage of normal sperm. Analyzing the group of patients in whom the average normal sperm morphology outcome across the three evaluations was ≤ 10%, a coefficient of variation of 13.9% and 10.63%, for 100 and 200 cells, respectively38, was obtained. Greater confidence in normal sperm morphology outcomes will therefore be achieved if 200 or more cells are evaluated in patients with low normal sperm outcomes. The evaluation of 200 or more cells per sample (per slide) will become more feasible as the speed of processors used in automated systems increases. In a study comparing sperm morphology analyzed by a computer equipped with a morphologizer with that using the traditional manual method, Wang et al.27 found a significant correlation between the two methods (r = 0.52; p < 0.0001) for percentage of normal forms. Although the mean percentages of normal forms classified by the methods were not significantly

different (72.4% vs. 72.3%), the limits of agreement were relatively large (–20.5% to +20.7%). Davis et al.12, comparing manual with automated classification, obtained a 60% unambiguous agreement. They also found that the automated classification method always resulted in a lower percentage of normal sperm than the manual method: 50.9% compared with 61.9%. Kruger et al.22, evaluating 43 slide preparations blindly, found that 84% of the FERTECH’s evaluations compared well with the manual method. In a subsequent study, Kruger et al.19 correlated the percentage normal morphology (strict criteria) outcomes between manual and automated evaluations and found the limits of agreement to be between 12.1 and –15.5%. In the percentage normal sperm morphology range 0–20%, the limits of agreement were, however, narrower (8.4 to –6.6%). The Spearman correlation coefficient for this study was 0.85, which was similar to the correlation (r = 0.83) obtained between two observers performing manual evaluations. Using the 14% fertility cut-off point for strict criteria, Kruger et al.21 found that the automated system was able to classify 81.3% (65/80) of cases, similar to the manual method. Four identical automated instruments (CellTrak-S), two each at two sites, were used to analyze (archive) videotape material40. The coefficients of variation obtained for repeated measures were between 1 and 8% for each variable measured on all instruments. Kruger et al.41 examined intermachine variation for two IVOS set-ups (Tygerberg vs. Norfolk), evaluating the same slides. The comparison showed no difference in the mean percentage of normal forms (15.6% vs. 15.8%) produced by the two systems. Although a correlation coefficient of 0.92 was obtained, the coefficient of variation was, however, 20.65%. In a multicenter study in which 30 sperm morphology slides were evaluated at five independent centers using the IVOS, the magnitudes of variation (coefficients of variation) obtained ranged between 11.36 and 23.09%42. Although most of the major variables (sample preparation, cell

ADVANCES IN AUTOMATED SPERM MORPHOLOGY EVALUATION

staining and classification system) influencing the evaluation of sperm morphology were eliminated, a variation of > 15% was still obtained between outcomes produced at the different centers. The results observed show that there is good agreement between an experienced manual observer’s evaluations and automated evaluations. The results also show that the use of an automated system does not mean that all variation will be eliminated. The technologist performing the computer-assisted evaluation therefore still has an important role to play in limiting variation. Factors other than sample and slide processing that the technologist can control, and which may significantly influence outcomes, are focus and illumination.

FERTILITY PREDICTION VALUE The primary objective for developing any diagnostic tool for in vitro or in vivo human fertility diagnosis is the ability to determine accurately the fertility potential, to provide the infertile couple with realistic advice with regard to conception potential. Replacement of manual evaluations of sperm morphology with automated evaluations, therefore, also requires unequivocal proof that the outcomes have predictive value. Wang et al.28 were among the first to assess the usefulness of automated sperm morphology evaluation to predict the outcome of human sperm fertilizing capacity. Multivariate discriminant analysis was used to analyze the ability to predict the outcome of the zona-free hamster-oocyte assay. The eight variables selected were able to predict fertility capacity with 74% accuracy, compared with 84% when the manual method was used. Kruger et al.22 determined the prognostic value of the IVOS by evaluating 21 slides from Tygerberg Hospital and 21 slides from Norfolk’s in vitro fertilization (IVF) program. The fertilization rates for the two fertility groups, < 14% and > 14% normal forms, were 33.3% (15/45) and 76.6% (46/60), respectively, for manual evaluations and 46.8%

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(30/64) and 75.6% (31/41), respectively, for automated evaluations (Tygerberg slides). Evaluations performed on the Norfolk slides produced a similar result: 27.4% (14/51) and 90.0% (127/141) and 33.9% (18/53) and 88.4% (123/139) for the manual and computer analyses, respectively. Sofitikis et al.18, using fresh sperm and a confocal scanning laser microscope, found that when the percentage normal forms were ≥ 22%, fertilization occurred in 25 of 26 cases, while below this percentage only two of 15 cases fertilized oocytes. MacLeod and Irvine26 examined the value of both manual and computer-assisted semen analysis (WHO 198743) using the Hamilton Thorne HTM-S 2030 in predicting the in vivo fertility (‘normal’ women) of cryopreserved donor semen. When the post-thaw semen profiles were compared, pregnant versus not pregnant, there were differences in respect of both morphometry and movement characteristics determined by the HTM-S. When multiple logistic regression was used to predict the achievement of pregnancy, the conventional criteria of semen quality were of no value (χ2 = 6.67; p = 0.353). However, the automated assessment of morphometric and movement characteristics successfully predicted outcome in 86.9% of cases (χ2 = 44.3; p = 0.0021). The most important variables in the regression were morphometric attributes (mean minor axis, mean major axis and mean area), amplitude of lateral head displacement and average path velocity. Kruger et al.21, using an automated system, showed that in patients with ≤ 10 × 106 motile spermatozoa, normal sperm morphology and the number of oocytes were important predictors of fertilization. The normal sperm morphology outcomes produced by automated evaluations were also found to be significantly (p = 0.0001) correlated with fertilization by logistic regression. Except for one case, all other zero fertilization cases were found to be within the group with < 106/ml sperm and < 10% normal sperm morphology. The overall fertilization rates for the fertility subgroups were: 45.6% (37/81) for the

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group with ≤ 4% normal forms, 72.5% (87/120) for the 5–9% group, 82.1% (46/56) for the 10–14% group and 85.2% (69/81) for the > 14% group. In another study conducted by the Kruger group44, the automated normal sperm morphology outcomes were found to be significant predictors of both fertilization (p = 0.0419) in vitro and pregnancy (p = 0.0210), using logistic regression models. The fertilization rates across the 5% normal sperm morphology fertility cut-off point were 39.4% (≤ 5%) and 62.9% (> 5%), while the pregnancy rates were 15.2% (≤ 5%) compared with 37.36% (> 5%). The significance of the 5% normal sperm morphology fertility cut-off point established by the manual evaluation of sperm morphology, using the strict criteria, has therefore been confirmed by computer-assisted evaluations. In a study using the Hobson Sperm Tracker, a positive correlation was found between the fertilization rate (FR%) and the proportions of sperm with a normal (oval) head shape, sperm exhibiting acrosomal vacuoles, sperm with a normal acrosomal size (40–70% of total head area) and sperm undergoing the acrosome reaction (AR) after adding follicular fluid20. Multiple regression analysis revealed that by incorporating the above four parameters, the sensitivity of prediction of in vitro fertilization rate values was 79% and the specificity was 93%, with a positive predictive value of 96%. During 1997–99, 1191 infertile couples with no known barrier to conception were assessed by conventional semen analysis and automated measurements, including motility, concentration and morphology evaluations33. A SHAMAS (sperm head automated morphometric analysis system) analysis was performed on Shorrstained smears of washed semen. The analysis measures %C, which is similar to conventional manual percentage normal morphology, and %Z, the percentage of sperm with characteristics conforming to those of sperm that bind to the zona pellucida of the human oocyte. Binding to the zona pellucida is essential for fertilization, and the process is highly selective for sperm with axial symmetry, a narrow neck and a large acrosomal

area. Three factors were found to be independently and significantly related to natural pregnancy in a multivariate Cox regression analysis, of which %Z was the most important, followed by VSL (straight-line velocity) and female age33. More large prospective randomized studies using automated evaluations are required to establish the ‘true’ clinical value of these systems. These must be performed using standardized and controlled slide preparation and sperm cell staining methods. The appropriateness of the manually established normal sperm morphology thresholds may have to be re-examined, or new thresholds may have to be determined by regression analysis.

CONCLUSIONS Automated systems have been shown to have the potential to eliminate the biases and subjectivity plaguing the manual evaluation of sperm morphology. Although they are objective, the accuracy of the results from these systems can also be compromised by methodological errors. Variables such as sperm preparation methods, sperm cell staining methods, focus, parameter settings and the softand hardware components used can have a significant effect on the precision of evaluations. To ensure comparative and reliable results, procedures and instruments must be standardized and quality control maintained. The studies performed, at least with the use of the IVOS, have shown that its precision and the predictive value of its outcomes are at least equal to the outcomes produced by an experienced observer performing manual evaluations. The group of patients identified with < 5% normal sperm morphology, as with the manual evaluation of sperm morphology, have been shown to have a significantly depressed fertilization and pregnancy probability. Further clinical studies are needed to determine the true value of the automated systems, whereby multiple parameters, morphometric and kinematic, are measured in relation to fertility outcomes to create predictive models.

ADVANCES IN AUTOMATED SPERM MORPHOLOGY EVALUATION

REFERENCES 1. Ombelet W, et al. Sperm morphology assessment: historical review in relation to fertility. Hum Reprod Update 1995; 1: 543 2. Coetzee K, Kruger TF, Lombard CJ. Predictive value of normal sperm morphology: a structured literature review. Hum Reprod Update 1998; 4: 73 3. Eliasson R. Standards for investigation of human semen? Andrologia 1971; 3: 49 4. Williams WW, ed. Sterility, the Diagnostic Survey of the Infertile Couple. Springfield, MA: WW Williams, 1964 5. Freund M. Standards for the rating of human sperm morphology. A co-operative study. Int J Fertil 1996; 11: 97 6. David G, et al. Anomalies morphologiques du spermatozoïde humain. 1) Propositions pour un système de classification. J Gynécol Obstet Biol Reprod 1975; 4 (Suppl 1): 17 7. Fredericsson B. Morphologic evaluation of spermatozoa in different laboratories. Andrologia 1979; 11: 57 8. Hofmann N, Freundl G, Florack M. Die Formstörungen der Spermatozoen im Sperma und Zervikalschleim als Spiegel testikulärer Erkrankungen, Gynäkologe 1985; 18: 189 9. Kruger TF, et al. Sperm morphologic features as a prognostic factor in in vitro fertilization. Fertil Steril 1986; 46: 1118 10. Davis RO, Katz D. Computer-aided analysis: technology at a crossroads. Fertil Steril 1993; 59: 953 11. Coetzee K, Kruger TF. Accuracy and prognostic value of computer-assisted (IVOS) sperm morphology evaluations. Middle East Fertil Soc J 1999; 4: 222 12. Davis RO, et al. Accuracy and precision of the CellForm-Human* automated sperm morphometry instrument. Fertil Steril 1992; 58: 763 13. Calamera JC, et al. Development of an objective and manual technique to study the human sperm morphology. Andrologia 1994; 26: 331 14. World Health Organization. WHO Laboratory Manual for the Examination of Human Semen and Semen–Cervical Mucus Interaction, 3rd edn. Cambridge: Cambridge University Press, 1992 15. Goulart AR, de Alencar Hausen M, Monteiro-Leal LH. Comparison of three computer methods of sperm head analysis. Fertil Steril 2003; 80: 625 16. Moruzzi JF, et al. Quantification and classification of human sperm morphology by computer-assisted image analysis. Fertil Steril 1988; 50: 142

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17. Coetzee K, et al. Comparison of two staining and evaluation methods used for computerized human sperm morphology evaluations. Andrologia 1997; 29: 133 18. Sofikitis NV, et al. Confocal scanning laser microscopy of morphometric human sperm parameters: correlation with acrosin profiles and fertilizing capacity. Fertil Steril 1994; 62: 376 19. Kruger TF, et al. Sperm morphology: assessing the agreement between the manual method (strict criteria) and the sperm morphology analyzer IVOS. Fertil Steril 1995; 63: 134 20. El-Ghobashy AA, West CR. The human sperm head: a key for successful fertilization. J Androl 2003; 24: 232 21. Kruger TF, et al. A prospective study on the predictive value of normal sperm morphology as evaluated by computer (IVOS*). Fertil Steril 1996; 66: 285 22. Kruger TF, et al. A new computerized method of reading sperm morphology (strict criteria) is as efficient as technician reading. Fertil Steril 1993; 59: 202 23. Lacquet FA, et al. Slide preparation and staining procedures for reliable results using computerized morphology (IVOS*). Arch Androl 1996; 36: 133 24. Menkveld R, et al. Effects of different staining and washing procedures on the results of human sperm morphology evaluation by manual and computerised methods. Andrologia 1997; 29: 1 25. Davis RO, Gravance CG. Standardization of specimen preparation, staining, and sampling methods improves automated sperm-head morphometry analysis. Fertil Steril 1993; 59: 412 26. MacLeod IC, Irvine DS. The predictive value of computer-assisted semen analysis in the context of a donor insemination programme. Hum Reprod 1995; 10: 580 27. Wang C, et al. Computer-assisted assessment of human sperm morphology: comparison with visual assessment. Fertil Steril 1991; 55: 983 28. Wang C, et al. Computer-assisted assessment of human sperm morphology: usefulness in predicting fertilizing capacity of human spermatozoa. Fertil Steril 1991; 55: 989 29. Mundy AJ, Ryder TA, Edmonds DK. Morphometric characteristics of motile spermatozoa in subfertile men with an excess of non-sperm cells in the ejaculate. Hum Reprod 1994; 9: 1701 30. Garret C, Baker HWG. A new fully automated system for the morphometric analysis of human sperm heads. Fertil Steril 1995; 63: 1306 31. Coetzee K, Kruger TF. Automated sperm morphology analysis: Quo Vadis. Assist Reprod Rev 1997; 7: 109

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32. Menkveld R, et al. The evaluation of morphological characteristics of human spermatozoa according to stricter criteria. Hum Reprod 1990; 5: 586 33. Garrett C, et al. Automated semen analysis: ‘zona pellucida preferred’ sperm morphometry and straightline velocity are related to pregnancy rate in subfertile couples. Hum Reprod 2003; 18: 1643 34. Ombelet W, et al. Results of a questionnaire on sperm morphology assessment. Hum Reprod 1997; 12: 1015 35. ESHRE Andrology Special Interest Group. Guidelines on the application of CASA technology in the analysis of spermatozoa. Hum Reprod 1998; 13: 142 36. Menkveld R, et al., eds. Atlas of Human Sperm Morphology. Baltimore: Williams & Wilkins, 1991 37. Davis RO, et al. Accuracy and precision of the CellForm-Human automated sperm morphometry instrument. Fertil Steril 1992; 58: 763 38. Coetzee K, Kruger TF, Lombard CJ. Repeatability and variance analysis on multiple readings performed by a computer semen analyser (IVOS). Andrologia 1999; 3: 165

39. Cooper TG, et al. Internal quality control of semen analysis. Fertil Steril 1992; 58: 172 40. Davis RO, Rothmann SA, Overstreet JW. Accuracy and precision of computer-aided sperm analysis in multicenter studies. Fertil Steril 1992; 57: 648 41. Kruger TF, et al. Computer assisted sperm analyzing system: an analysis of intermachine morphology evaluations and intraslide evaluations using IVOS (Dimension system version 3). Presented at the American Society for Reproductive Medicine Congress, Boston, November, 1996, S223 42. Coetzee K, et al. Assessment of inter- and intralaboratory sperm morphology readings using a Hamilton Thorne Research IVOS semen analyzer. Fertil Steril 1999; 71: 80 43. World Health Organization. WHO Laboratory Manual for the Examination of Human Semen and Semen–Cervical Mucus Interaction, 2nd edn. Cambridge: Cambridge University Press, 1987 44. Coetzee K, et al. Clinical value in using an automated semen morphology analyser (IVOS). Fertil Steril 1999; 71: 222

11 Sperm morphology training and quality control programs are essential for clinically relevant results Daniel R Franken, Thinus F Kruger

INTRODUCTION

specific endocrine disruptors are present7–11. These statements were confirmed by Coetzee et al.12, who summarized all the important articles in a meta-analysis. Training of andrology technologists can be accomplished using different educational approaches, of which the one-to-one workshop is the most successful teaching method. Direct communication and input on a one-to-one basis with an experienced worker ensures that the trainee understands the basic concepts of sperm morphology. This method, however, has a disadvantage in that only a small number of trainees can be trained per session. Our experience has indicated that a maximum of ten students per teacher can be trained per session13,14. A second and also valuable teaching method is the so-called group consensus technique15. In this method, the trainer (usually an individual with ample experience in sperm morphology evaluation) uses computer or video images that are projected onto a screen during training sessions. The advantage of this method lies in the fact that large numbers of students can be trained during a single session. The disadvantage of this method lies in the mass communication style, and the individual is often lost during group discussions. A third training method is the use of an interactive CD-ROM program. Such an interactive computer program contains a variety of

Primary knowledge and understanding of the morphological appearance, and bright-field microscopic configuration, of a normal human sperm cell form the basis of the evaluation method for sperm morphology in which more strict criteria are applied. Disagreement in the results can be caused by a variety of factors, such as discrepancy in the specific techniques used during the analysis. Since more clinicians are becoming aware of the importance of training and subsequent quality control measurements, standardization in semen analysis methodologies has become mandatory. In close agreement with the present author’s beliefs, Kvist and Bjorndahl have made an important contribution towards the standardization of techniques needed to obtain a globally accepted and World Health Organization (WHO)-recognized semen analysis result1. The techniques focus mainly on assessments of sperm concentration, sperm motility, sperm morphology and sperm vitality. Several authors have stressed the value of the assessment of human sperm morphology during both in vitro2–5 and in vivo6 studies. Furthermore, assessment of human sperm morphology and sperm concentration can also serve as a variable in reproductive health studies involving endocrinology and environmental toxicology, when 181

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high-quality images of numbered spermatozoa. Advantages of this method include training at the individual’s own leisure and time, as he/she can repeat specific sections of the program where certain concepts are poorly understood. During previous studies we have presented numerous sperm morphology workshops in Africa, the Middle East and Europe. The format of these workshops consisted generally of handson, one-on-one teaching, accompanied by various sessions of consensus training, as well as the use of a CD-ROM program (Strict 1-2-3). During the group consensus training sessions, participants were requested to evaluate photomicrographic images of sperm cells projected onto a large nonreflecting screen. It is important to remember that the educational value of the training will be enhanced if trainees are exposed to all the abovedescribed methods.

SPERM MORPHOLOGY QUALITY CONTROL Sperm morphology evaluations have important clinical value only in cases where the evaluation of normal/abnormal cells is done with accuracy. In most cases, manual reading by light microscopy under high-power magnification (1000×) has been the method of evaluation. Several factors have been identified that can influence the outcome of sperm morphology readings. These factors include quality of the slide, and staining procedures. Typically, a poor slide consists of a thick semen layer with multiple sperm cells on top of one another, thus causing extensive overlapping of cell heads, tails and debris. Each andrology laboratory should therefore have an internal as well as an external quality control program. For example, the results obtained from each technician on the quality control sample are tabulated and plotted on a graph against the sample number. The mean and standard deviation of the results for each sample are computed and also plotted against the sample number. As

part of the internal quality control system, each andrology technician should be able to prepare high-quality sperm slides in order to provide repeatable and reliable morphology readings for the referring clinicians. At Tygerberg, a protocol has been developed for the preparation of sperm slides that not only are of a high quality but also fulfill the requirements of the manual reading techniques for sperm morphology16. These slides adhere to the description for the preparation of semen smears supplied by the WHO17–20.

Slide preparation For each sample, at least two smears should be prepared from a fresh sample for duplicate assessments in case of poor staining. The slide should first be cleaned, washed in 70% alcohol and dried, before a drop of semen is applied to the slide (Figure 11.1)20. To ensure optimal slide quality, the following standard protocol should be used during slide preparations: (1) frosted, precleaned glass slides with grounded edges are used at all times; (2) sperm counts are used as a guide to determine the sperm droplet size eventually used to prepare the smear (if the sperm count is > 60 × 106 cells/ml, a < 10-µl droplet is used, while if the sperm count is < 60 × 106 cells/ml, a 10–30-µl drop is used; the final number of sperm cells in both cases should

Second slide is used to make thin semen smears

Droplet volume is determined by the sperm concentration

Figure 11.1 Feathering method to prepare undiluted sperm morphology smears

SPERM MORPHOLOGY TRAINING AND QUALITY CONTROL PROGRAMS

produce 8–12 spermatozoa per high-power magnification); (3) the semen drop is typically placed in the middle of the slide at a point more or less 20% from the frosted end, using a micropipette fitted with disposable tips (Gilson P100; Lasec Laboratories, Cape Town, South Africa). The semen is gently touched at a 45° angle with the width side of a second slide; this allows the semen to spread evenly across the width of the first slide, after which the second slide is slightly pulled backwards and then pushed forwards while pushing downwards over the entire length of the first slide.

MONITORING THE TECHNICIAN’S SPERM MORPHOLOGY READING SKILLS The Tygerberg approach A typical Tygerberg sperm morphology training session consists of a multiple approach method that relies on hands-on, one-on-one individual training (experienced worker vs. inexperienced worker). We believe that this training method is imperative during the initial stages of teaching. Furthermore, we also use the consensus training and CD-ROM interactive programs. In our experience, after the training sessions, participants were enrolled on the continuous quality control (CQC) program. Participants received, on a quarterly basis, two Papanicolaou prestained sperm slides from normo-, terato- or severe teratozoospermic samples. The participant recorded the percentage of normal cells present on these slides, and the results were forwarded to the reference laboratory at Tygerberg Hospital. The ‘correct’ results according to the reference laboratory, i.e. the percentage of normal forms present on each of the slides, were subsequently supplied to the participating laboratory13,14. Due to the fact that the morphological slides used for evaluation of the standard of the trainees were random samples from different sperm donors, standardization was needed with respect

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to an index that is not dependent on the morphological level. On the assumption that the reference laboratory’s morphology reading is the gold standard, an index was calculated using the following standardized statistical score: Standard deviation (SD) score = trainee score – reference laboratory score divided by SD test slides that were shipped to trainees14

As expected, the standard deviation decreases with lower levels of morphology, i.e. < 4% normal forms. The SD score reflects the number of SD units by which the measurement of the trainee differs from the gold standard for the specific slide. Each trainee can be evaluated according to the SD score for his/her level of agreement with the gold standard. Two SD score levels were chosen in order to evaluate poor readings, and for this purpose we selected the values ± 0.5SD and ± 0.2SD. The individual SD scores obtained from the training and follow-up contacts can be plotted against time on a graph that also indicates the limits. An ongoing study at Tygerberg Hospital aims to record the value of quarterly monitoring and refresher courses on morphology reading skills of technicians over a period of 40 months. Nineteen individuals from 13 different andrology laboratories from Switzerland, Malaysia and Singapore were enrolled in a sperm morphology quality control program after initial training sessions. The mean values for the test slides (two slide sets) reported by each individual are presented in Figure 11.2. We regarded recordings outside the ± 0.2SD score as a warning (Figure 11.2), and results outside the ± 0.5SD score as an indicator for the individual to become concerned about his/her sperm morphology reading skills. Five of the 19 participants (Figure 11.2 numbers 1, 7, 8, 9 and 19) attended annual refresher courses during the period. Participants 13 and 19 did not attend any refresher training, but maintained the reading skills acquired after one-to-one training. Adequate technician training is of paramount significance to achieve consistent results

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SD scores for two test slides

1 Action

0.5

Warning

0 Warning

–0.5

Action

–1

–1.5 1

9 10 11 12 13 14 15 16 17 18 19

Participating individuals

Figure 11.2

Mean standard deviation (SD) scores reported by 19 individuals from 13 andrology laboratories for test slides 1 and 2

within a given laboratory. Even when strict criteria are utilized13,14, interlaboratory variation is probably the result of various factors, including (1) different semen and smear preparation techniques, (2) differences in interpretation and (3) technician experience15. Using specific criteria, we were able to classify the trainees according to their reported results.

Classification of reading skills Poor reading skills

If 50% of readings recorded over the 40-month period were inside the limits of error, i.e. the ± 0.5SD score, poor reading standards were assumed. Using the overall correctness of each individual, the results depicted in Figure 11.2 indicated that five (26%) participants (5, 6, 11, 17, 18) had poor reading skills during the evaluation period. Marginal reading skills

If 51–59% of readings recorded over the 40month period were within the ± 0.5SD score, marginal reading skills were assumed.

Good reading skills

If 60–69% of the readings recorded over the 40month period fell inside the limits of error, i.e. the ± 0.5SD score, good reading standards were assumed. Five (26%) individuals (9, 12, 14, 15, 16) had good reading skills. Excellent reading skills

If ≥ 70% of the readings recorded over the 40month period were within the ± 0.5SD score, excellent reading skills were assumed. Results in Figure 11.2 show that nine (47%) of the partaking individuals (1, 2, 3, 4, 7, 8, 10, 13, 19) maintained excellent reading skills. Our results clearly illustrate that an external quality control program can be successfully implemented on condition that continuous monitoring is part of the program. In general, we were satisfied with the overall reading skills of the study group, since 73% maintained sperm morphology reading skills that were classified as good or excellent. We firmly believe that the technical maintenance of morphology readings is, apart from the initial training sessions, also dependent on annual refresher courses. The five participants, namely 1, 7, 8, 9 and 19 who randomly attended refresher

SPERM MORPHOLOGY TRAINING AND QUALITY CONTROL PROGRAMS

courses were able to maintain their acquired reading skills. These individuals consistently produced reading skills that were within ± 0.2SD score limits of error (Figure 11.3). This study also highlights the feasibility of initiating a global sperm morphology quality control program. Finally, an important finding during the study was the significant role that the annual refresher courses played in the maintenance of morphology reading skills. Here, for the first time, we illustrated that those technicians who attended refresher courses were able to maintain their morphology reading skills over an extended period. In general, all participants (except refresher course attendees) showed a decline in reading at about 6–9 months after initial training. We believe that this is a tendency that will occur in any andrology unit, and laboratory directors should be aware of this phenomenon. Previous studies21–23 concluded that the only way to ensure comparable interlaboratory results is through participation in a multicenter proficiency testing program24. Similar to the present study, Keel et al.21 suggested that such a proficiency testing system should comprise an external interlaboratory quality program. During this program, simulated identical patient specimens are tested by participating individuals/ laboratories.

0.25 0.2 0.15 SD scores

0.1 0.05 0 –0.05 –0.1 –0.15 –0.2 –0.25

9 10 11 12 13 14

Slide set number

Figure 11.3 Standard deviation (SD) scores of an individual with excellent sperm morphology reading skills

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REFERENCES 1. Kvist U, Bjorndahl L. Editorial. In Kvist U, Bjorndahl L, eds. Manual on Basic Semen Analysis. Oxford: Oxford University Press, 2002: v 2. Kruger TF, et al. Sperm morphologic features as a prognostic factor in in vitro fertilization. Fertil Steril 1986; 46: 1118 3. Kruger TF, et al. Predictive value of abnormal sperm morphology in in vitro fertilization. Fertil Steril 1988; 49: 111 4. Enginsu ME, et al. Evaluation of human sperm morphology using strict criteria after Diff-Quik staining: correlation of morphology with fertilization in vitro. Hum Reprod 1991; 6: 854 5. Ombelet W, et al. Teratozoospermia and in-vitro fertilization: a randomized prospective study. Hum Reprod 1994; 9: 1479 6. Eggert-Kruse W, et al. Clinical relevance of sperm morphology assessment using strict criteria and relationship with sperm–mucus interaction in vivo and in vitro. Fertil Steril 1995; 63: 612 7. Carlsen E, et al. Evidence for decreasing quality of semen during the past 50 years. Br J Med 1992; 303: 609 8. Auger J, et al. Decline in semen quality in fertile men in Paris during the past 20 years. N Engl J Med 1995; 332: 281 9. Irvine SE, et al. Evidence of deteriorating semen quality in the United Kingdom: birth cohort study in 577 men in Scotland over 11 years. Br J Med 1996; 312: 467 10. Auger J, Jouannet P. Evidence for regional differences of semen quality among French fertile men. Hum Reprod 1997; 12: 740 11. Swan SH, Elkin EP, Fenster L. The question of declining sperm density revisited: an analysis of 101 studies published 1934–1996. Environ Health Perspect 1997; 108: 961 12. Coetzee K, Kruger TF, Lombard CJ. Predictive value of normal sperm morphology: a structured literature review. Hum Reprod Update 1998; 4: 73 13. Franken DR, Barendson R, Kruger TF. A continuous quality control (CQC) program for strict sperm morphology. Fertil Steril 2000; 74: 721 14. Franken DR, et al. The development of a continuous quality control (CQS) programme for strict sperm morphology among sub-Saharan African laboratories. Hum Reprod 2000; 15: 667 15. Eustache F, Auger J. Inter-individual variability in the morphological assessment of human sperm: effect of

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16.

17.

18. 19.

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the level of experience and the use of standard methods. Hum Reprod 2003; 18: 1018 Menkveld R, et al. Effect of different staining and washing procedures on the results of human sperm morphology evaluation by manual and computerized method. Andrologia 1997; 29: 1 Menkveld R, et al. The evaluation of morphological characteristics of human spermatozoa according to stricter criteria. Hum Reprod 1990; 5: 586 Menkveld R, et al., eds. Atlas of Human Sperm Morphology. Baltimore: Williams & Wilkins, 1991 Menkveld R, Kruger TF. Evaluation of sperm morphology by light microscopy. In Acosta AA, Kruger TF, eds. Human Spermatozoa in Assisted Reproduction. London: Parthenon Publishing, 1996: 189 World Health Organization. WHO Laboratory Manual for the Examination of Human Semen and

21.

22.

23.

24.

Semen–Cervical Mucus Interaction, 4th edn. Cambridge: Cambridge University Press, 1999: 1 Keel BA, et al. Results of the American Association of Bio-analysts national proficiency testing programme in andrology. Hum Reprod 2000; 15: 680 Cooper TG, Atkinson AD, Nieschlag E. Experience with external quality control in spermatology. Hum Reprod 1999; 14: 765 Dunphy BC, et al. Quality control during the conventional analysis of semen, an essential exercise. J Androl 1989; 10: 378 Neuwinger J, Behre, HM, Nieschlag E. External quality control: the andrology laboratory: an experimental multicenter trial. Fertil Steril 1990; 54: 308

12 Role of acrosome index in prediction of fertilization outcome Roelof Menkveld

INTRODUCTION

shown that the AI is another sensitive parameter, like normal sperm morphology evaluated according to strict criteria, in the prediction of in vitro fertilization rates of ≥ 50%, and that the AI is especially useful in the P-pattern group of patients. Since the AI can provide additional information, compared with normal sperm morphology, it can therefore be regarded as an independent parameter13 for the prediction of assisted reproductive procedure outcomes. This additional role of the AI as a prognostic factor has been discussed in only a few publications14–18, and is reviewed briefly in this chapter, with emphasis on the functional role played by the acrosome in the fertility pathway, particularly in relation to sperm binding to the zona pellucida, and its usefulness in assisted reproduction procedures, especially intracytoplasmic sperm injection (ICSI).

The clinical usefulness of the strict Tygerberg criteria for sperm morphology evaluation1 for in vitro fertilization outcome and later for in vivo pregnancies has been demonstrated by Kruger et al.2,3 and Van Zyl et al.4, respectively, and thereafter has been confirmed in many publications5. However, even in the presence of the so-called Ppattern or in the poor-prognosis group (≤ 4% morphological normal forms), some men achieve fertilization of oocytes in vitro, and in vivo pregnancies occur occasionally3,5. Therefore, efforts to develop more sensitive predictors, especially for expected in vitro fertilization rates, are continuously being put forward. Some of these predictors are based on sperm biochemical tests such as the sperm chromatin dispersion test6, the sperm chromatin structure assay7 and the ubiquitin-based sperm assay8, while others incorporate a combination of semen variables, for example the post-wash total progressively motile cell count9, or have refined certain existing semen variables such as sperm morphology by the use of more specific sperm morphology parameters, namely the sperm deformity index10 or the spontaneous acrosome reaction as seen with Spermac staining11. In this regard, Menkveld et al.12 introduced the acrosome index (AI) as an additional tool in the prediction of in vitro fertilization outcome. It was

ROLE OF THE ACROSOME IN THE FERTILITY PATHWAY The acrosome is formed by the Golgi apparatus during spermatogenesis, and can be described as a secretory granule situated at the apex of the sperm head, consisting of an inner acrosomal membrane that is closely associated with the nucleus of the sperm and which is continuous with the outer acrosomal membrane. The acrosomal matrix 187

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proper is located between the two membranes. The whole acrosome as well as the rest of the spermatozoon is covered by the plasma membrane. The acrosome contains a number of enzymes such as (pro)acrosin, which plays a vital role in the fertility pathway with regard to sperm binding to and penetration of the zona pellucida19. With contact of the spermatozoon to the zona pellucida, the acrosome undergoes the acrosome reaction, which can be described as an exocytotic event involving localized fusion between the outer acrosomal and plasma membranes, resulting in the formation of vesicles with the release of mainly the enzymes hyaluronidase and (pro)acrosin through the holes formed by the vesicles, and is one of the most important steps in sperm binding to and penetration of the zona pellucida20,21. It is now becoming increasingly evident that, for these functions to take place, especially with regard to sperm binding to the zona pellucida, normal sperm morphology and especially normal acrosomal morphology is essential22,23. Strong selection also takes place at the zona pellucida for spermatozoa with normal-sized acrosomes, as was nicely illustrated by Garrett and Baker24. Acrosomal size also plays an important role in the ability of the spermatozoon to undergo the acrosome reaction. Spermatozoa with large acrosomes were associated with a significantly higher percentage of live spontaneous acrosome-reacted spermatozoa, while spermatozoa with small acrosomes were associated with a high percentage of sperm death25. Semen samples containing a low percentage of spermatozoa with intact acrosomes were also associated with total fertilization failure11, due to the inability of these spermatozoa to bind to the zona pellucida, as the acrosome reaction must take place at the time of binding to the zona pellucida. However, the inability of zona pellucida-bound spermatozoa to undergo the zona pellucida-induced acrosome reaction may also play an important role in non-fertilization26,27. According to Benoff et al.28, the human sperm acrosome reaction occurs in vitro only in the most morphologically normal spermatozoa, and about

50% of all in vitro fertilization (IVF) failures are thought to be related to anomalies of acrosome structure and function.

MICROSCOPIC EVALUATION OF ACROSOMAL MORPHOLOGY At first, the human sperm acrosome was deemed too small to be visualized by direct microscopy, and scanning electron and transmission electron microscopy were used or advocated29. However, the ability to evaluate acrosome morphology by light microscopy is now acknowledged1,12. Visual evaluation of sperm acrosomal morphology, performed with good bright-field optics at 1000× or preferably 1250× magnification on high-quality Papanicolaou-stained smears, is based on acrosomal size, its form and the staining characteristics of the acrosome, and can be performed simultaneously with the routine sperm morphology evaluation1,12. For classification as a morphologically normal acrosome, the same principles are applicable as for the classification of morphologically normal spermatozoa according to strict criteria, except that the postacrosomal area of the sperm head can be abnormal, but no neck/midpiece and tail abnormalities or cytoplasmic residues may be present1. If the spermatozoon is classified as normal, the acrosome must always be classified as normal. This means that the acrosome index will always be equal to, but in most cases greater than, the percentage of morphologically normal spermatozoa. When the acrosome evaluation is done simultaneously with the routine morphology evaluation, two laboratory counters are needed. On the first counter the sperm morphology is scored as normal or abnormal, and the second is used to keep a record of the acrosomes considered to be normal, or the whole range of acrosomal defects can be scored1,12,30. Acrosomal defects as seen with the light microscope can be classified as specific defects or as nonspecific alterations. Specific acrosomal defects, which are mostly concerned with acrosome size,

ROLE OF ACROSOME INDEX IN PREDICTION OF FERTILIZATION OUTCOME

are genetically caused31,32, such as globozoospermia19, and the miniacrosome defect33. However, genetic sperm defects are not limited to the acrosome only, but may affect any part of the spermatozoon, the short or stump tail defect being one observable by light microscopy34. These conditions are rare, but when they occur, are easy to detect using the light microscope. However, acrosomes can also be classified as too large, an abnormality that may in some cases be associated with a higher rate of spontaneous acrosomal reactions25 and decreased in vitro fertilization rates2,12. Staining defects may include irregular acrosomes, multiple vacuoles, cysts and ‘empty’ acrosomes29. These staining defects may indicate damage of the acrosome membranes, with subsequent leaking of (pro)acrosin from the acrosomes35. Jeulin et al.29 found low fertilization rates of semen samples containing predominantly spermatozoa with acrosome staining defects, and postulated that the low in vitro fertilization rates associated with these increased acrosomal abnormalities might not be due to the presence of abnormal acrosomes per se, but due to a relationship between acrosomal abnormalities and nuclear immaturity of the spermatozoa. Sperm DNA abnormalities may be due to the production of reactive oxygen species (ROS) by the spermatozoa themselves, but mostly due to leukocytes present in semen samples36.

ROLE OF THE ACROSOME INDEX IN ASSISTED REPRODUCTION In 1986, Kruger et al.2 reported that sperm morphology evaluated according to strict criteria1 was a strong prognosticator of in vitro fertilization outcome in cases with a normal sperm concentration (≥ 20 × 106/ml) and progressive motility (≥ 30% motility). A cut-off value at 15% morphologically normal spermatozoa was found to be associated with a fertilization rate of 37% in the ≤ 14% group, but no pregnancies, and 82% in the ≥ 14% group. In 1988, Kruger et al.3 also reported

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that a drastic drop in the fertilization rate (7.6%) occurred when < 4% morphologically normal spermatozoa was observed in a semen sample, while in the 4–14% group the fertilization rate was 63.9%. In an initial investigation of the role of acrosomes and fertilization30, it was observed that two distinct morphological acrosome patterns could be observed in the < 4% normal morphology group, whereby fertilization did and did not occur. In the few men with good fertilization, it was striking to observe a pattern of slightly and moderately elongated spermatozoa, but with morphologically normal (size and form) acrosomes. In one of the cases, four of four ova were fertilized in vitro, although there were only 2% morphologically normal spermatozoa present, but a total of 17% of spermatozoa had normal acrosomes. In a typical case in the group with no fertilization, of six ova inseminated in vitro, it was observed that small and/or abnormal acrosomes were mainly present. This case presented with only 1% morphologically normal spermatozoa and with a total of only 4% normal acrosomes. In an ongoing study including 23 males, Menkveld et al.35 found that when the acrosome morphology was classified into different groups, i.e. normal, small, staining defects and amorphous, and expressed as an acrosome index (percentage normal acrosomes), no fertilization occurred when the acrosome index was ≤ 15%. Important was the fact that, once again, cases were found where normal sperm morphology was < 5% but the acrosome index was > 15%. In those cases with an AI > 15% the fertilization rate was always ≥ 50%. The relationship between AI (percentage normal acrosomes) and fertilization rate was underlined by the observation that statistically significant differences were found for the acrosome index between groups with fertility rates of < 50% and ≥ 50%, i.e. 1.5 ± 1.9% and 28.5 ± 11.6% normal acrosomes, respectively35. When a receiver operating characteristic (ROC) curve analysis was performed on the IVF results of the 23 males37, a cut-off value of ≥ 10%

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(sensitivity 100% and specificity 100%) was obtained for the prediction of a fertilization rate of ≥ 50%. This result was confirmed in a follow-up study of 33 males12. A higher correlation between AI and fertilization rate (r = 0.8631; p < 0.0001) was found, compared with the correlation between morphology and fertilization rate (r = 0.7953; p < 0.0001). This means that the AI can be regarded as a more sensitive measurement of fertilization potential than sperm morphology, especially in the < 5% morphologically normal spermatozoa group. This may be attributed to the fact that spermatozoa with normal acrosomes but classified as abnormal according to the strict Tygerberg criteria, such as slightly abnormal, or slightly and moderately elongated spermatozoa, are more likely to bind to the zona pellucida22,23 and to undergo the acrosome reaction38, compared with spermatozoa from samples with a low AI (< 10% normal acrosomes). In 1998, Menkveld et al.39 reported on the predictive role of the AI and normal sperm morphology compared with that of the teratozoospermia index (TZI) as described in the 1992 WHO manual40 in a study of 110 patients. It was found that the AI at a cut-off value of ≥ 9% had a better predictive value to predict the possibility of a > 50% in vitro fertilization rate, compared with normal sperm morphology at > 5%, and sperm morphology had a better predictability compared with the TZI at ≤ 1.46, by ROC curve analysis, with areas under the curve of 0.920, 0.739 and 0.634, respectively. In a study to define normal cut-off values based on data from fertile and subfertile populations, Menkveld et al.41 determined the AI cut-off value to be at 8% normal acrosomes. These results are in agreement with previous reports on the role of acrosomal morphology11,23,28,29. Liu and Baker23 found that in cases with < 30% morphologically normal spermatozoa (according to WHO criteria), the acrosome status (percentage normal) was an important

prognosticator of expected fertilization in vitro. Chan et al.11 reported that semen samples with a low percentage (< 40%) of spermatozoa with intact acrosomes were associated in 31% of cases with total fertilization failure (TFF). Benoff et al.28 showed that by increasing the insemination concentration of spermatozoa to at least 25 000/ml acrosomally normal spermatozoa in patients with poor acrosomal morphology, fertilization rates and pregnancy rates reached similar levels compared with couples in whom the male presented with normal acrosomal morphology. These publications confirmed the fact that a minimum number or a minimum percentage of spermatozoa with normal acrosomes are needed for normal fertilization to occur in vitro, and underline the important physiological role played by acrosomes in the fertilization pathway21. Few reports by other investigators on the role of the AI per se have been published so far. Söderlund and Lundin14 investigated fertilization of split sibling oocytes for IVF and ICSI in patients but with < 5% morphologically normal spermatozoa with ≥ 1 × 106/ml motile spermatozoa after a swim-up procedure. With the aid of ROC curve analysis for IVF rates of ≥ 50%, a cut-off value for the AI was determined at 7%. The 81 patients were divided into two groups: group A with AI < 7% included 42 patients, and group B with AI ≥ 7% included 39 patients. The in vitro fertilization rate in group A was 43.5%, which was significantly lower compared with that of group B at 71.9% (p = 0.001). The study showed that in semen samples with < 5% morphologically normal spermatozoa and an AI ≥ 7%, the mean fertilization rate was about 70%, compared with the mean fertilization rate of 40% in the < 7% AI group. Rhemrev et al.15 found no pregnancies in a group of 87 couples where the males presented with an AI < 5% and a fast total radical-trapping antioxidant procedure (TRAP) of < 1.14 mmol/l.

ROLE OF ACROSOME INDEX IN PREDICTION OF FERTILIZATION OUTCOME

THE ACROSOME INDEX AND SELECTION OF PATIENTS FOR INTRACYTOPLASMIC SPERM INJECTION With the introduction of ICSI42, new doors have been opened for couples with severe male fertility problems, as with ICSI it is possible to overcome functional deficiencies, abnormal sperm morphology or shortage of adequate numbers of motile spermatozoa by placing a spermatozoon directly into the oocyte. ICSI can be regarded as a very invasive procedure, and may also be more expensive in many centers compared with standard IVF. Furthermore, concern exists over the possible negative effects of ICSI, due to the injection of possibly genetically abnormal spermatozoa43, on their offspring with regard to genetic and congenital abnormalities44, increased spontaneous abortions, preterm deliveries and reduced birth weights45. ICSI should therefore be restricted to those couples with an unacceptably high risk of a low fertilization rate or total fertilization failure. However, a recent publication by Greco et al.46 has shown that ICSI can also have a positive side, in so far as males with DNA-damaged (fragmented) spermatozoa in their semen samples, leading to decreased implantation and pregnancy rates but normal fertilization rates47, can be successfully treated by ICSI with testicular spermatozoa. It was found that DNA fragmentation was significantly decreased in testicular spermatozoa, leading to a pregnancy rate of 44.4% and an implantation rate of 20.7%. An alternative to performing a testicular biopsy to obtain spermatozoa with low DNA damage may be to perform a selection procedure by the use of cervical mucus, as Bianchi et al.48 have shown that spermatozoa able to cross a cervical mucus barrier possessed higher levels of DNA protamination and practically no signs of endogenous nick translations. In cases of extreme oligozoospermia49, cryptozoospermia, globozoospermia50 or obstructive azoospermia, the choice of an ICSI procedure is self-evident, but problems in deciding between

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ICSI and IVF may arise when there is a chance of recovering sufficient numbers of motile spermatozoa after sperm preparation15. In the previous section dealing with in vitro fertilization, an oocyte fertilization cut-off value of ≥ 50% was used to determine the AI cut-off value12,14. However, an in vitro fertilization rate cut-off value point of 50% may be regarded as too high to decide between ICSI and IVF. A more appropriate fertilization cut-off point for ICSI may be regarded as a fertilization rate of < 37% (two standard deviations (SDs) below the normal expected fertilization rate). The only data so far available on this aspect were published by Menkveld et al. in 199951. They conducted a prospectively designed study to investigate use of the AI as an additional parameter to sperm morphology evaluated by strict criteria in the selection of patients for ICSI. In this study, 134 semen samples were examined blindly on the day of IVF oocyte recovery. Sperm morphology and sperm acrosomal morphology were visually evaluated using light microscopy and expressed as the acrosome index (percentage normal acrosomes). ROC curve analysis indicated that for in vitro fertilization rates of ≤ 37% (2SD below their normal mean fertilization rate), the normal sperm morphology cut-off value was ≤ 3% (sensitivity 51%, specificity 89%, area under the curve 0.718) and for the acrosome index ≤ 7% (sensitivity 86%, specificity 86%, area under the curve 0.929). By lowering the fertilization rate cut-off points to < 30% and < 25%, with ROC curve analysis, the AI cut-off point was lowered to ≤ 6%, while for a fertilization rate of ≤ 20% the AI cut-off point increased to ≤ 7% again. In all instances the morphology cut-off point remained at ≤ 3%. With the AI cut-off points at ≤ 6% and ≥ 7%, fertilization rates were 22.4% (35/156 ova) and 74% (365/489 ova), respectively. According to the above results, the AI cut-off point can be set at ≤ 6% and be clinically helpful in the selection of patients who would need ICSI, especially in the group of patients showing P-pattern morphology (≤ 4%).

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In the study by Söderlund and Lundin14 there was no significant difference between the two AI groups (group A with AI < 7% and group B with AI ≥ 7%) when ICSI was performed, with fertilization rates of 65.8% and 63.5%, respectively. The study showed that in semen samples with < 5% morphologically normal spermatozoa and an AI ≥ 7% the mean fertilization rate was about 70%, compared with the mean fertilization rate of 40% in the < 7% AI group. The conclusion of Söderlund and Lundin was that evaluation of the sperm morphology and AI in combination with the total number of normal spermatozoa available after sperm preparation has a better predictive value for the choice of IVF or ICSI treatment than that of the basic semen parameters alone14. Rhemrev et al. suggested that patients with an AI < 5% may benefit from ICSI to prevent total fertilization failure and/or that males with < 2% morphologically normal spermatozoa should go for ICSI, and concluded that the evaluation of sperm morphology and AI in combination with the total number of motile sperm available after sperm preparation/separation may have a better predictive value for choice of IVF or ICSI treatment than the basic semen parameters alone15.

CONCLUSIONS The AI can play an important role in the decisionmaking process of assisted reproductive treatment procedures. Over time, the AI cut-off value for expected IVF rates of ≥ 50% has been lowered from ≥ 16% normal acrosomes to ≥ 9%, and determined to be at ≥ 8% to distinguish between a fertile and a subfertile population. However, the most important aspect is to decide whether to advise couples to undergo the ICSI procedure or not, and for this purpose a cut-off AI value of ≤ 6% normal acrosomes was determined by Menkveld et al.51 as well as by Söderlund and Lundin14, and < 5% by Rhemrev et al.15. Both Söderlund and Lundin, and Rhemrev et al. suggested that the combination of AI cut-off value

and total progressively motile spermatozoa number obtained after sperm preparation (< 1.10 × 106/ml and < 1.0 × 106/ml, respectively) are strong tools in the decision whether to perform ICSI14,15.

REFERENCES 1. Menkveld R, et al. The evaluation of morphological characteristics of human spermatozoa according to stricter criteria. Hum Reprod 1990; 5: 586 2. Kruger TF, et al. Sperm morphological features as a prognostic factor in in vitro fertilization. Fertil Steril 1986; 46: 1118 3. Kruger TF, et al. Predictive value of abnormal sperm morphology in in vitro fertilization. Fertil Steril 1988; 49: 122 4. Van Zyl JA, Kotze TJvW, Menkveld R. Predictive value of spermatozoa morphology in natural fertilization. In Acosta AA, et al., eds. Human Spermatozoa in Assisted Reproduction. Baltimore: Williams & Wilkins, 1990: 319 5. Coetzee K, Kruger TF, Lombard CJ. Predictive value of normal sperm morphology: a structured literature review. Hum Reprod Update 1998; 4: 73 6. Fernández JL, et al. The sperm chromatin dispersion test: a simple method for the determination of sperm DNA fragmentation. J Androl 2003; 24: 59 7. Evenson DP, Larson KL, Jost LK. Sperm chromatin structure assay: its clinical use for detecting sperm DNA fragmentation in male infertility and comparisons with other techniques. J Androl 2002; 23: 25 8. Sutovsky P, Terada Y, Schatten G. Ubiquitin-based sperm assay for the diagnosis of male factor infertility. Hum Reprod 2001; 16: 250 9. Rhemrev JPT, et al. The postwash total progressive motile sperm cell count is a reliable predictor of total fertilization failure during in vitro fertilization treatment. Fertil Steril 2001; 76: 884 10. Aziz N, et al. The sperm deformity index: a reliable predictor of the outcome of oocyte fertilization in vitro. Fertil Steril 1996; 66: 1000 11. Chan PJ, et al. Spermac stain analysis of human sperm acrosomes. Fertil Steril 1999; 72: 124 12. Menkveld R, et al. Acrosomal morphology as a novel criterion for male fertility diagnosis: relation with acrosin activity, morphology (strict criteria), and fertilization in vitro. Fertil Steril 1996; 65: 637

ROLE OF ACROSOME INDEX IN PREDICTION OF FERTILIZATION OUTCOME

13. Jeyendran RS, Zaneveld LJD. Controversies in the development and validation of new sperm assays. Fertil Steril 1993; 59: 726 14. Söderlund B, Lundin K. Acrosome index is not an absolute predictor of the outcome following conventional in vitro fertilization and intracytoplasmic sperm injection. J Assist Reprod Genet 2001; 18: 483 15. Rhemrev JPT, et al. The acrosome index, radical buffer capacity and number of isolated progressively motile spermatozoa predict IVF results. Hum Reprod 2001; 16: 1885 16. Kruger TF, Menkveld R. Acrosome reaction, acrosin levels, and sperm morphology in assisted reproduction. Assist Reprod Rev 1996; 6: 27 17. Mortimer D, Menkveld R. Sperm morphology assessment – historical perspectives and current opinions. J Androl 2001; 22: 192 18. Menkveld R. The use of the acrosome index in assisted reproduction. In Kruger TF, Franken DR, eds. Atlas of Human Sperm Morphology Evaluation. London: Taylor & Francis, 2004: 35 19. Schill W-B. Some disturbances of acrosomal development and function in human spermatozoa. Hum Reprod 1991; 6: 969 20. Mortimer D. Practical Laboratory Andrology. Oxford: Oxford University Press, 1994. 21. Wassarman PM. Fertilization in mammals. Sci Am 1988; 256: 52 22. Menkveld R, et al. Sperm selection capacity of the human zona pellucida. Mol Reprod Develop 1991; 30: 346 23. Liu DY, Baker HWG. Morphology of spermatozoa bound to the zona pellucida of human oocytes that failed to fertilize in vitro. J Reprod Fertil 1992; 94: 71 24. Garrett G, Baker WHG. A new fully automated system for the morphometric analysis of human sperm heads. Fertil Steril 1995; 63:1306 25. Menkveld R, et al. Relationship between human sperm acrosomal morphology and acrosomal function. J Assist Reprod Genet 2003; 20: 432 26. Bastiaan HS, et al. Relationship between zona pellucida-induced acrosome reaction, sperm morphology, sperm–zona pellucida binding, and in vitro fertilization. Fertil Steril 2003; 79: 49 27. Liu DY, Baker HWG. Disordered zona pellucidainduced acrosome reaction and failure of in vitro fertilization in patients with unexplained infertility. Fertil Steril 2003; 79: 74 28. Benoff S, et al. Numerical dose-compensated in vitro fertilization inseminations yield high fertilization and pregnancy rates. Fertil Steril 1999; 71: 1019

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29. Jeulin C, et al. Sperm factors related to failure of human in-vitro fertilization. J Reprod Fertil 1986; 76: 735 30. Menkveld R, Kruger TF. Evaluation of sperm morphology by light microscopy. In Acosta AA, Kruger TF, eds. Human Spermatozoa in Assisted Reproduction. Carnforth: Parthenon Publishing, 1996: 89 31. Hofmann N, Haider SG. Neue Ergebnisse morphologisher diagnostik der Spermatogenesestörungen. Gynaköloge 1985; 18: 70 32. Baccetti B, et al. Genetic sperm defects and consanguinity. Hum Reprod 2001; 16: 1365 33. Baccetti B, et al. A ‘miniacrosome’ sperm defect causing infertility in two brothers. J Androl 1991; 12: 104 34. Favero R, et al. Embryo development, pregancy and twin delivery after microinjection of ‘stump’ spermatozoa. Andrologia 1999; 31: 335 35. Menkveld R, et al. Relationships between sperm acrosomal status, acrosin activity, morphology (strict Tygerberg criteria) and fertilization in vitro. Hum Reprod 1994; 9 (Suppl 4): 99 36. Henkel R, et al. Effect of reactive oxygen species produced by spermatozoa and leukocytes on sperm function in non-leukocytospermic patients. Fertil Steril 2005; 83: 635 37. Menkveld R, et al. Acrosomal morphology as an additional new criterion for male fertility diagnosis: relationship with sperm functional aspects. Hum Reprod 1995; 10 (Suppl 2): 100 38. Heywinkel E, Freudl G, Hofmann N. Acrosome reaction of spermatozoa with different morphology. Andrologia 1993; 25: 137 39. Menkveld R, Stander FSH, Kruger TF. Comparison between acrosome index and teratozoospermia index as additional criteria to sperm morphology in the prediction of expected in-vitro fertilisation outcome. Hum Reprod 1998; 13 (Abstr book 1): 52 40. World Health Organization. WHO Laboratory Manual for the Examination of Human Semen and Sperm–Cervical Mucus Interaction, 3rd edn. Cambridge: Cambridge University Press, 1992 41. Menkveld R, et al. Semen parameters including WHO and strict criteria morphology, in a fertile and subfertile population: an effort towards standardisation of in vivo thresholds. Hum Reprod 2001; 16: 1165 42. Palermo G, et al. Pregnancies after intracytoplasmic injection of a single spermatozoon into an oocyte. Lancet 1992; 340: 17 43. In’t Veld PA, et al. Intracytoplasmic sperm injection (ICSI) and chromosomally abnormal spermatozoa. Hum Reprod 1997; 12: 752

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44. Bonduelle M, et al. Incidence of chromosomal aberrations in children born after assisted reproduction through intracytoplasmic sperm injection. Hum Reprod 1998; 13: 781 45. Aytoz A, et al. Outcome of pregnancies after intracytoplasmic sperm injection and the effect of sperm origin and quality on this outcome. Fertil Steril 1998; 70: 500 46. Greco E, et al. Efficient treatment of infertility due to sperm DNA damage by ICSI with testicular spermatozoa. Hum Reprod 2005; 20: 266 47. Henkel R, et al. Influence of deoxyribonucleic acid damage on fertilization and pregnancy. Fertil Steril 2004; 81: 965 48. Bianchi PG, et al. Human cervical mucus can act in vitro as a selective barrier against spermatozoa

carrying fragmented DNA and chromatin structural abnormalities. J Assist Reprod Genet 2004; 21: 97 49. Strassburger D, et al. Very low counts affects the results of intracytoplasmic sperm injection. J Assist Reprod Genet 2000; 17: 431 50. Coetzee K, et al. Short communication: an intracytoplasmic sperm injection pregnancy with a globozoospermia male. J Assist Reprod Genet 2001; 18: 311 51. Menkveld R, et al. The use of the acrosome index as an additional morphology parameter in the clinical selection of patients for ICSI. Hum Reprod 1999; 14 (Abstr book 1): 156

13 Acrosome reaction: physiology and its value in clinical practice Daniel R Franken, Hadley S Bastiaan, Sergio Oehninger

INTRODUCTION

analysis because of its expense and complexity. The need thus arose for a relatively simple test by which the human acrosome reaction could be quantified at the light microscope level. In addition, the labile nature of the human sperm acrosome makes analysis of the reaction problematic. The chosen procedure must therefore be able to distinguish between normal and degenerative reactions. In addition to the above limitations, the existence of multifactorial induction and regulating systems and the individual perspectives and methods of measurement chosen by laboratories contribute to the uncertainty that still exists regarding the subject.

For successful fertilization of oocytes by spermatozoa, a set of functionally normal parameters with regard to oocyte and spermatozoon maturity is of paramount importance. In the spermatozoon, besides motility and zona binding, the occurrence of the acrosome reaction is of primary importance in the development of functional capability. However, before spermatozoa are able to undergo the acrosome reaction, essential modifications in cell physiology of the sperm, called capacitation, must occur. Numerous studies have tried to elucidate the precise biochemical and biophysical changes involved in the process determining a spermatozoon’s fertilizing capacity. The normal progress of these changes may display important biomarkers of fertilizing ability, including the ability of the sperm to penetrate the cumulus oophorus, the corona radiata, the zona pellucida (ZP) and the vitelline membrane. The ability to evaluate the human acrosome reaction is, however, restricted by a practical limitation. The loss of the human acrosome cannot be observed on living sperm by phase-contrast or differential interference-contrast microscopy, because of its relatively small size compared with that of other mammalian species. Initially, best results were obtained by electron microscopy; this method, however, is not suited for routine

THE BIOCHEMISTRY OF CAPACITATION AND THE ACROSOME REACTION The acrosome of a human spermatozoon is a membrane-bound organelle that develops during spermatogenesis as a product of the Golgi complex. It surrounds the anterior portion of the sperm nucleus and can be divided into the following components: • Plasma membrane; • Outer acrosomal membrane; 195

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• Acrosomal matrix; • Inner acrosomal membrane; • Equatorial segment. Factors inhibiting capacitation are incorporated into the membranes of sperm during maturation in the epididymis1,2. These factors include sialoglycoproteins, sulfoglycerolipids and steroid sulfates, which induce a significant increase in the net negative charge of the outer acrosomal membrane3. This state of decapacitation (stability) is maintained after ejaculation by the presence of inhibitory macromolecules in the seminal plasma4 and in the lower areas of the female reproductive tract1. A specific glycoprotein has been identified as the primary decapacitator1; it is bound to the outer acrosomal membrane and can prevent interaction with extracellular signals as well as inhibit ion channel activity and/or enzymes5. This stability is further enhanced by the incorporation of cholesterol into the acrosomal membrane complex, preferentially into the plasma membrane. Sperm acquire their fertilizing ability in vivo during their migration through the female genital tract. Capacitation can also be induced in vitro in chemically defined media6. The complete process, however, is not yet fully understood, but is thought to involve major biochemical and biophysical changes in the membrane complex, energy metabolism and ion permeability. The most significant changes are: • Modification, redistribution and/or loss of the epididymal seminal plasma and cervical decapacitation factors – by exogenous or endogenous proteases specifically activated (plasmin, kallikrein and acrosin)1,2; • Net negative charge decrease by endogenous hydrolases (sterol sulfatase)3; • Membrane fluidity increase by the efflux of cholesterol, altering the cholesterol/phospholipid ratio and the influx of unsaturated fatty acids; these changes are thought to be serum albumin-mediated3,7;

• Altered permeability allowing the increased uptake of calcium ions, glucose and oxygen, resulting in an elevated energy state, inducing hyperactivated motility and ability to undergo the acrosome reaction8,9. Notwithstanding the extent of these changes, capacitation is also thought to be a reversible event. The exact threshold of irreversibility, however, remains undefined, i.e. what constitutes the boundary between capacitation and the acrosome reaction. Many of these structural changes proposed to characterize capacitation may, however, be irreversible, which may lead to an untimely acrosome reaction. The acrosome reaction can therefore be seen as the end-point of capacitation. Signals for initiation of the acrosome reaction are most likely received by one or more receptors on the plasma membrane surface, which transmit the message across the membrane. Zaneveld et al.5 proposed a mechanism by which membrane receptor activation of guanosine triphosphate (GTP)-binding proteins stimulates secondmessenger systems which regulate ion transport. An endogenous calcium ion threshold concentration has long been thought of as the primary inducer of the acrosome reaction8,10. Yanagimachi and Usui11 showed that upon the addition of calcium, but not magnesium, guinea-pig sperm incubated for several hours in calcium-free medium underwent the acrosome reaction within 10 minutes. Since then, calcium has been implicated in many reactions leading to complete loss of the acrosome and eventually fertilization10: • Activation of many enzymes hyaluronidase, phospholipase A2);

(acrosin,

• Activation of enzyme messenger systems (adenylate cyclase); • Neutralization of the net negative charge; • Induction of hyperactivated motility3. Stock and Fraser12 examined the extracellular Ca2+ requirements for the support of capacitation and the spontaneous acrosome reaction in human

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spermatozoa, and concluded that the optimal conditions for capacitation and the acrosome reaction in human spermatozoa require extracellular Ca2+ at 1.80 mmol/l with calcium channels providing a means of calcium entry. In contrast, White et al.13 found that the acrosome reaction rates at 4 hours and 20 hours were little different in media with or without calcium, although the absence of calcium had a significant effect on the quality of motility. The human sperm acrosome reaction is an exocytotic event characterized by significant ultrastructural changes leading to the complete loss of the outer acrosomal cap, following: • Decondensation of the acrosomal matrix; • Fenestration and vesiculation of the plasma membrane and outer acrosomal membrane; • Dispersion of the vesicles; • Release of the acrosomal content. Nagae et al.14 proposed a unique morphological sequence for this acrosome reaction. Vesicles established in the intermediate stage were formed by the invagination and pinching off of the outer acrosomal membrane and the plasma membrane. Stock et al. described a similar characterization15. In contrast, Yudin et al.16 found that human sperm undergo an acrosome reaction similar to that of other mammals, in which the outer acrosomal and plasma membranes initially fuse by fenestration followed by vesiculation. The dispersion of these vesicles leaves the spermatozoon surrounded by a single, continuous membrane, i.e. the inner acrosomal membrane. In addition to these changes, the membrane proteins of the plasma membrane overlying the equatorial/ postacrosomal region of the sperm head undergo a conformational change, resulting in activation3. This activation may facilitate fusion of the sperm with the oocyte vitelline membrane. The loss of the membranes also releases or exposes activated lysins, assisting sperm penetration of the ZP2. Acrosin, a trypsin-like serine protease found in the acrosome, has been implicated

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in a number of events leading to fertilization. These include assisting sperm penetration of the ZP, triggering of the acrosome reaction2 and activation of regulatory enzymes involved in Ca2+ transport5. Studies using p-aminobenzamidine (PABA)17, an inhibitor of mouse sperm acrosin, have shown that acrosin is a necessary factor for dispersal of the acrosomal matrix, probably through the activation of proacrosin. In the presence of PABA, the membranes undergo normal vesiculation, but ZP penetration is inhibited.

MEASUREMENT OF THE ACROSOME REACTION IN HUMAN SPERMATOZOA Capacitation and the acrosome reaction can be induced chemically, providing a controlled means for the evaluation of acrosomal exocytosis. Table 13.1 presents agents and methods commonly used to monitor and trigger the acrosome reaction. The acrosome reaction can be examined during basal conditions (incubating sperm under capacitating conditions) and/or following exogenous induction with pharmacological or physiological agonists. Inducers that have been analyzed in the clinical setting include the calcium ionophore

Table 13.1 Agents and methods commonly used to monitor the spontaneous and induced acrosome reaction Inducers of the acrosome reaction Calcium ionophore Pentoxifylline Follicular fluid Progesterone Solubilized zona pellucida Methods to assess the acrosome reaction Optical microscopy: triple-staining Transmission electron microscopy Chlortetracycline fluorescent assay Fluorescent lectins Labeling with antibodies Flow cytometry

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A2318718–21, pentoxifylline22,23, steroids24–27, follicular fluid (FF)28, solubilized ZP29–31 and low temperature20. The methodologies used to examine acrosomal exocytosis have included triple staining (optic microscopy)32, transmission electron microscopy33, chlortetracycline fluorescent assay34, fluorescent lectins35,36, labeling with antibodies37 and flow cytometry38. There are, however, inherent negative sideeffects that must be taken into account, such as the possible negative effect on motility, and the fact that the means of induction overrides the normal processes involved in the acrosome reaction. For example, using a calcium ionophore, a toxic chemical substance, as the inducer, the time required for capacitation is minimized. The addition of complex biological fluids such as maternal cord serum, FF, granulosa cells, cumulus oophorus and ZP, even though uncontrolled in nature, is physiologically more correct. This is of particular relevance when future improvements in the treatment in male infertility are to be introduced into an assisted reproduction program, and also for furthering our knowledge of the in vivo regulatory system. As mentioned above, several techniques have been employed to detect the acrosome reaction, each with its own level of characterization. Aitken and Brindle39, however, showed that probes targeting different components involved in the acrosome reaction measure acrosomal loss at different rates. The labile nature of the acrosomal vesicle also requires a means of determining sperm viability and distinguishing between ‘normal’ and degenerative reactions. In the triple-stain technique according to Talbot and Chacon35, trypan blue is used. Cross et al.36 included the supravital stain Hoechst 33258, while Aitken et al.21, in their protocol for assessing the ability of viable human spermatozoa to acrosome-react in response to A23187, employed a fluorescein-conjugated lectin in concert with the hypo-osmotic swelling test. The use of these different techniques may have a significant influence on the interpretation and comparison of results. This is illustrated by the

often equivocal results obtained in acrosome reaction studies.

PHYSIOLOGICAL INDUCERS AND REGULATORS OF THE ACROSOME REACTION Stock et al.15 found that 32% of sperm coincubated with oocyte–cumulus complexes for 14–18 hours had initiated or completed the acrosome reaction. The effect of a number of female reproductive tract products on sperm fertilizing capacity was evaluated by coincubating fertile sperm samples with endometrial, oviductal, granulosa and cumulus cells, FF and maternal serum, by Bastias et al.40. Compared with control samples, endometrial and oviductal cell cultures did not alter sperm fertilizing capacity or their movement characteristics. Sperm coincubated with FF, granulosa cells or cumulus cells, however, exhibited a significantly higher ability to penetrate zona-free hamster ova. It is therefore reasonable to propose that secretions of cumulus cells could be involved in regulation of the sperm acrosome reaction. Siegel et al.41 concluded from their study that components within the FF might influence sperm physiology and enhance sperm fertilizing capacity by activating sperm proteinase systems involved in sperm reaction and interaction. An active Sephadex G-75 fraction identified in FF was found to stimulate a rapid, transient increase in the intracellular free Ca2+ in human spermatozoa42. The ability of this fraction to induce the acrosome reaction led the authors to conclude that this influx of Ca2+ is responsible for the initiation of acrosomal exocytosis. Using indirect immunofluorescence, FF was found to induce the acrosome reaction rapidly after the sperm had been incubated for at least 10 hours43. Induction of the human acrosome reaction by whole FF and/or the active Sephadex G-75 component was found to satisfy the ultrastructural criteria known for physiological reactions, as shown by transmission electron microscopy16.

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Yudin et al.16 also showed that human sperm capacitated for 6 hours at 40°C and then incubated with FF for 180 seconds resulted in 40% of the sperm reacting. Sperm incubated for 22 hours before FF treatment had their acrosome reaction rate enhanced six-fold, illustrating the potential effect of FF. An adequate preincubation period followed by FF treatment therefore seems to result in the synchronization of capacitation and facilitation of the acrosome reaction. In contrast, Stock et al.44, examining the incidence of spontaneous acrosome reactions in human spermatozoa exposed to FF, found that FF can stimulate the acrosome reaction, but only after continuous exposure (> 6 hours) to 50% FF/medium. A short exposure (1 hour), even after 24 hours of preincubation, did not induce the reaction. Recent studies have shown that the human sperm acrosome reaction-inducing activity in FF can be attributed to progesterone (P). Osman et al.24 purified an active fraction from the fluid aspirated from preovulatory human follicles and identified it as 4-pregnen-3,20-dione (progesterone) and 4-pregnen-17α-ol-3,20-dione (17-hydroxyprogesterone). This was confirmed by Blackmore et al.25, Foresta et al.45 and Baldi et al.46, who found that only P and 17-hydroxyprogesterone were able to induce a rapid, long-lasting, dosedependent increase of intracellular free calcium, with maximum effect being obtained with 1.0 µg/ml. Sueldo et al.27, however, found that 1.0 µg/ml of P enhanced the acrosome reaction only after 24 hours of incubation. Luconi et al.47 found several rapid nongenomic effects of P and estrogen (E) in human spermatozoa. They seem to be mediated by the steroids binding to specific receptors on the plasma membrane that are different from the classical ones. Progesterone, specifically, has been demonstrated to stimulate calcium influx, tyrosine phosphorylation of various sperm proteins, including extracellular signaling-regulated kinases, chloride efflux and cyclic adenosine monophosphate (cAMP) increase, finally resulting in the activation of spermatozoa through the induction

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of capacitation, hyperactivated motility and the acrosome reaction. On the other hand, E, which is present in micromolar levels in follicular fluid, seems to modulate sperm responsiveness to P. This occurs when E acts rapidly on calcium influx and on protein tyrosine phosphorylation. In general, the isolation and characterization of the putative membrane receptors for P (mPR) and E (mER) in spermatozoa are still elusive. Luconi47 obtained evidence supporting the existence and functional activity of mPR and mER in human spermatozoa. To characterize these membrane receptors, they used two antibodies directed against the ligand-binding domains of the classical receptors, namely c262 and H222 antibodies for PR and ER, respectively, hypothesizing that these regions should be conserved between nongenomic and genomic receptors. In Western blot analysis of sperm lysates, the antibodies detected a band of about 57 kDa for PR and 29 kDa for ER, excluding the presence of the classical receptors. On live human spermatozoa, both antibodies were able to block the calcium and AR response to P and E, respectively, whereas antibodies directed against different domains of the classical PR and ER were ineffective. Furthermore, c262 antibody also blocks in vitro the human sperm penetration of hamster oocytes. Taken together, all these data strongly support the existence of mPR and mER different from the classical ones, mediating rapid effects of these steroid hormones in human spermatozoa. Siegel et al.41 also found that FF obtained from different women under different stimulation regimens did not affect the fertilizing potential differently. Morales et al.48, however, found that there was a positive, highly significant (r = 0.72; p > 0.005) correlation between the acrosome reaction-inducing activity and the P level of each FF sample. Nevertheless, recent reports on the chemical nature of the acrosome reaction-inducing molecule present in FF have been contradictory. In contrast to the authors who attributed the acrosome reaction-inducing activity present in human

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FF to P, Miska et al.49 reported evidence to suggest that this substance is a protein. These authors identified a protein with a molecular mass of about 50 kDa and demonstrated the substance’s sensitivity to unspecific proteases, increased temperature and pH changes. In a further study50, the same authors identified the acrosome reactioninducing substance (ARIS) as the progesteronebinding protein corticoid-binding globulin (CBG). Using anti-human CBG antibodies and dextran-coated charcoal they showed that only P bound to CBG could induce the acrosome reaction. CBG is a member of the SERPIN (serine proteinase inhibitor) superfamily that binds P tightly. Proteolysis by serine proteinases results in the release of this steroid hormone. This mechanism is thought to be essential in the activation of neutrophils by delivering high local concentrations of corticoids in inflammatory processes. The serine proteinase involved in the spermatozoon is acrosin, localized at the plasma membrane, with inactive proacrosin located within the acrosome. During capacitation, proacrosin and acrosin are exposed at the plasma membrane. The CBG–progesterone complex, which may become bound on the plasma membrane, will therefore be proteolytically cleaved by exposed acrosin, leading to high local concentrations of P and subsequent induction of the acrosome reaction, confirming the important role of acrosin in physiological induction of the acrosome reaction. The exact mechanism underlying P-stimulated calcium entry in human sperm has, however, not been fully established. Another question to be answered concerns the source of CBG. Whether it is of liver origin, where it is known to be produced, and then is accumulated in FF, or whether the cumulus or granulosa cells can synthesize this protein, still have to be established. The importance of the ZP for induction of the acrosome reaction, however, is well recognized29–31,51–53. Spermatozoa must penetrate this last barrier in the reacted state before they can penetrate and fertilize the oocyte. In vitro studies by Saling and Storey54, using mouse sperm, were

the first to demonstrate a role for the ZP in the acrosome reaction. They incubated cumulus-free eggs with sperm suspensions in which > 50% of the population had undergone the acrosome reaction. After gradient centrifugation, only acrosome-intact sperm were detected on the ZP. They concluded that the acrosome reaction of a fertilizing mouse sperm occurs on the ZP. Saling et al.55 and Bleil and Wassarman56 maintained that, at least in the mouse, the acrosome reaction is induced by a ZP constituent, the glycoprotein ZP3. They proposed the following concept: • Sperm attachment to the ZP; • Specific and irreversible binding to the ZP; • Physiological induction of the acrosome reaction; • ZP penetration. Cross et al.36 used two approaches to test the ability of the human ZP to induce acrosomal exocytosis in human sperm. Non-viable human oocytes and acid-disaggregated zonae were used, and both the zona binding and exposure to disaggregated zona induced the acrosome reaction. Using the monoclonal antibody T-6, Coddington et al.57 found that 93% of sperm bound to bisected human ZP exhibited immunofluorescent patterns indicative of the acrosome reaction. Hoshi et al.58 observed that the acrosome reaction rate after sperm attachment to the zona for 6 hours was 35.7 ± 17.7%, which was higher than in controls (2.8 ± 1.9%). The results so far indicate that the ability of spermatozoa to migrate to the ZP is a closely regulated process, ensuring that only sperm at the correct stage attach to and penetrate the ZP. It has been shown that P exerts a priming effect on the ZP-stimulated acrosome reaction in the mouse59 and in the human60. In the former studies, treatment with P followed by ZP led to maximal breakdown of phosphatidylinositol4,5-bisphosphate (PIP2), signaling a priming role for P in the initiation of exocytosis. Cross et al.29 were the first to report that treatment of human

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spermatozoa in suspension with aciddisaggregated human ZP (2–4 zonae pellucidae (ZP)/µl) increased the incidence of acrosomereacted spermatozoa. Lee et al.30 demonstrated that pertussis toxin treatment of human spermatozoa inhibited the (solubilized) ZP-induced acrosome reaction. In contrast, acrosomal exocytosis induced by the calcium ionophore A-23187 was not inhibited by pertussis toxin pretreatment. Studies by Franken et al.31 showed a dose-dependent effect of solubilized human ZP on the acrosome reaction in the range 0.25–1 ZP/µl, and also confirmed the involvement of Gi-protein during the ZP-induced acrosome reaction of human spermatozoa. More recently, Franken et al.51 reported the validation of a new microassay using minimal volumes of solubilized, human ZP to test the physiological induction of the acrosome reaction in human spermatozoa (ZP-induced acrosome reaction test or ZIAR). In such studies, a dose-dependent effect of solubilized ZP on acrosomal exocytosis was observed, reaching maximal induction using 1.25–2.5 ZP/µl for both the microassay and the standard (macro)assay. Furthermore, the inducibility of the acrosome reaction by a calcium ionophore was similar in both assays. Differences among species may account for the disparity in the results published, but the major differences among researchers are probably caused by the different experimental conditions and the varied assessment criteria. The in vitro conditions under which the work is performed can have a dramatic effect on the normal biochemical (metabolic and acrosomal) reactions of sperm, and also on the maturity of the oocyte–cumulus complexes and the molecules trapped in the complexes.

THE ACROSOME REACTION SITE The precise site of the acrosome reaction still remains clouded by controversy. Three possible sites have been proposed3:

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• The oviductal fluid of the ampulla; • The cumulus matrix; • The surface of the zona pellucida. The majority of the initial sperm acrosome reaction site studies were performed on the cauda epididymal sperm of the golden hamster, because of its relatively large acrosomal cap. The progress of the acrosome reaction can therefore be followed by phase-contrast microscopy. Data from the oviductal studies are, however, equivocal. Cummins and Yanagimachi61 studied the ampullary contents of female hamsters by phase-contrast microscopy, 4–10 hours after insemination with golden-hamster caudal epididymal sperm, and observed that 93 of 96 sperm swimming freely had modified and swollen acrosomal caps. In an earlier study, Yanagimachi and Phillips62, also using phase-microscopy, found that only four of 14 free-swimming golden-hamster sperm had modified acrosomal caps. Evaluating the cumulus matrix as the site of acrosomal reaction, Cummins and Yanagimachi61 reported that all motile golden-hamster spermatozoa observed in the cumuli from oviducts had undergone or were undergoing the acrosome reaction. Yanagimachi and Phillips62 reported that motile sperm, within the cumulus of golden-hamster cumulus-intact complexes from the ampulla, had modified acrosomes. However, in a videotaped study, Cherr et al.63 found that only 3–6% of sperm had actually completed the acrosome reaction within the cumulus matrix, which was comparable to control levels of the acrosome reaction occurring in free-swimming sperm. Their study included both cumulus-intact and cumulusfree eggs, with a higher percentage of reacted sperm found in association with the zona pellucida of cumulus-intact eggs than with the ZP of cumulus-free eggs. Tesarik64 undertook a study to determine the site of the acrosome reaction of spermatozoa penetrating into freshly inseminated human oocytes. The inseminated oocytes were treated with an

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antiacrosin monoclonal antibody, and the bound antibody visualized at the ultrastructural level with the use of a second peroxidase-conjugated antibody. His findings indicated that the acrosome reaction of the fertilizing spermatozoon must be exactly synchronized with its penetration through the egg vestments by the action of specific acrosome reaction-promoting substances in the oocyte–cumulus complex. Quantitative analysis of the results showed that the number of spermatozoa within the ZP corresponded to the number of acrosin deposits associated with acrosomal ghosts on the ZP surface. Using the triple-stain technique, the acrosomal status of sperm outside and within the cumulus during in vitro fertilization was examined65. The percentage of sperm undergoing the acrosome reaction increased significantly (p < 0.05) from 14.5 ± 1.5 to 24.5 ± 1.9 when incubated with a cumulus mass, and further increased to 49 ± 3.3 when incubated with mature expanded cumulus tissue containing an oocyte. White et al.13 exposed prepared spermatozoa for 20–30 minutes to large pieces of human cumulus oophorus; while these spermatozoa were able to penetrate deep into the cumulus mass, none were found to have clearly undergone the acrosome reaction. From this study they also concluded that spermatozoa did not require a capacitation period for penetration. In an in vitro system, depolymerization (softening) of the cumulus matrix may occur because of the high sperm concentration used. This may allow sperm to reach the zona pellucida with intact acrosomes. Cummins and Yanagimachi61 therefore studied the ability of hamster sperm to penetrate intact cumulus matrices at low (3 : 1) sperm/egg ratios. Uncapacitated sperm were unable to penetrate the cumuli; at least 2 hours of preincubation were required. Of the 628 in vitro capacitated sperm seen in and on the cumuli, 270 could penetrate, of which only ten had intact unmodified acrosomes. They concluded that penetration of the cumulus was limited to a phase in capacitation before completion of the acrosome reaction, since sperm that had lost the acrosomal cap

penetrated poorly and showed reduced viability. Corselli and Talbot66 also developed a system in which physiological sperm numbers (1–100) were used to challenge fresh hamster oocyte–cumulus complexes in capillary tubes. Their results showed that capacitated acrosome-intact hamster sperm can penetrate the extracellular matrix between the cumulus cells and can ultimately bind to the ZP. The results obtained by these two groups indicated that uncapacitated sperm tend to adhere to the cumulus cells on the periphery and are unable to penetrate, and that sperm that have lost the acrosomal cap also penetrate poorly. The cumulus matrix may therefore be seen as a selection barrier, allowing only morphologically normal sperm that can undergo a normal acrosome reaction to penetrate the zona pellucida, and/or it may contain molecules that influence the ability of sperm to undergo the reaction. CBG and P, which are present in high concentrations in the cumulus matrix, have been proposed as the physiological stimulus for initiation of the acrosome reaction.

CLINICAL RELEVANCE OF THE ACROSOME REACTION In two independent experiments, Barros et al.67 and Singer et al.68 using golden-hamster sperm and human sperm, respectively, found that sperm became infertile with prolonged incubation, as judged by their ability to bind to and penetrate the ZP. The reason for this decline in penetration with increasing incubation time was attributed to an increase in the percentage of acrosome-reacted sperm. In contrast, an increase in the penetration of zona-free hamster eggs was seen with increasing incubation time (increase in the acrosome-reacted population). Thus, acrosome-reacted sperm are prevented from penetrating the ZP. These results indicate that the fertilizing ability of spermatozoa is a time-dependent process. Although only acrosome-reacted spermatozoa are capable of fusing with zona-free oocytes, there

ACROSOME REACTION

is no significant correlation between the proportion of acrosome-reacted cells and the levels of sperm–oocyte fusion observed. These two bioassays are thus measuring two different aspects of the sperm’s ability to acrosome-react. White et al.13 similarly concluded that there was no relationship between the acrosome reaction rate and the fertilization rate of normal human oocytes in vitro. In a study to assess whether patients who did not fertilize human oocytes in vitro could be identified by a lack of acrosomal response of their spermatozoa, Pampiglione et al.69 found that patients who fertilized oocytes responded like fertile donors. It was also calculated that an acrosome reaction rate of < 31.3% predicted fertilization failure in 100% of cases. While spontaneous reactions bore no relation to fertility, the inducibility of the acrosome reaction (i.e. the difference between spontaneous and induced acrosome reaction), which describes the ability of viable sperm to undergo the acrosome reaction, was significantly reduced or absent in subfertile men, indicating acrosomal dysfunction as a likely cause of fertilization failure19. Henkel et al.70 showed that inducibility should be at least 7.5% to be indicative of good fertilization. A > 13% level of acrosome-reacted sperm after induction of the acrosome reaction was also shown to have predictive value for fertilizing potential, because elevated levels of sperm able to lose their acrosome are necessary for successful fertilization. For diagnostic purposes, the kind of induction, be it physiological by means of the ZP glycoproteins or non-physiological by the application of a calcium ionophore or low temperature, is apparently not important. However, inducibility and appropriate timing of the acrosome reaction with the penetration of the ZP61 are prerequisites for good fertilization52,71. Liu and colleagues72,73 reported a sperm defect called disordered ZP-induced acrosome reaction (DZPIAR). This defect was the cause of failure of sperm penetration in a group of in vitro fertilization (IVF) patients with a long duration of infertility. These patients were previously diagnosed as

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having idiopathic infertility with repeated poor or no fertilization during IVF treatment. Bastiaan et al.52,71 and Esterhuizen et al.53 reported similar findings in a study in which 164 andrology referrals were divided according to the percentage of normal spermatozoa in the ejaculate, namely < 4% normal forms (n = 71), 5–14% normal forms (n = 73) and > 14% normal forms (n = 20). ZIAR data for the < 4%, 5–14% and > 14% groups were 9.6 (± 0.6)%, 13.9 (± 0.5)% and 15.0 (± 1.1)%, respectively. The ZIAR result for fertile control men was 26.6 (± 1.4)%, which differed significantly from that of the three andrology referral groups. Likewise, significant differences were recorded during the hemizona assay, namely, 38.0% (< 4% normal forms), 54.5% (5–14% normal forms) and 62.6% (> 14% normal forms). Among the group with > 14% normal (Table 13.2) forms, five cases out of 21 (23%) had impaired ZIAR outcome (< 15%). Three (14%) of these men had normal morphology and sperm–zona binding, but showed a decrease in ZIAR results. The study concluded that ZIAR testing should become part of the second level of male fertility investigations, i.e. sperm functional testing, since 14% of andrology referrals revealed an impaired acrosome reaction response to solubilized ZP.

Table 13.2 Sperm–oocyte interaction results for five cases with impaired zona pellucida-induced acrosome reaction test (ZIAR) Sperm–zona binding (HZI)

Morphology (% normal forms)

ZIAR (%)

Case

HZI, hemizona binding index

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Liu and Baker72 also studied the frequency of defective sperm–ZP interaction in oligozoospermic infertile men. Sperm–ZP binding and the ZPinduced acrosome reaction were performed in 72 infertile men with oligozoospermic semen (sperm count < 20 × 106/ml). Oocytes that failed to fertilize in clinical IVF were used for the tests. Four oocytes were incubated for 2 hours with 2 × 106/ml motile sperm selected by swim-up from each semen sample. The number of sperm bound per ZP and the ZIAR were assessed. Under this condition, an average ≤ 40 sperm bound/ZP was defined as low sperm–ZP binding and a ZIAR ≤ 15% was defined as low ZIAR. In the 72 oligozoospermic men, 28% had low sperm–ZP binding. Of those (n = 52) with normal sperm–ZP binding, 69% had low ZIAR. Overall, 78% had either low ZP binding or normal ZP binding but low ZIAR. Only 22% had both normal sperm–ZP binding and normal ZIAR. They concluded that oligozoospermic men have a very high frequency of defective sperm–ZP interaction, which may be a major cause of infertility or low fertilization rate in standard IVF. Esterhuizen et al.53 reported the ZP-induced acrosome reaction response (ZIAR) among 35 couples with normal and G-pattern (good prognosis) sperm morphology and repeated poor fertilization results during assisted reproduction treatment. Results were compared with in vitro fertilization rates of metaphase II oocytes. Interactive dot diagrams divided the patients into two groups, i.e. ZIAR < 15% and ZIAR > 15%, with mean fertilization rates of 49% and 79%, respectively. The area under the curve was 99% and the 95% confidence interval did not include 0.5, demonstrating that the ZIAR test is able to predict fertilization failure among IVF patients.

CONCLUSIONS The fertilizing spermatozoon undergoes a continuous reactionary process that is temporally and spatially regulated. Spermatozoa respond to

signals during specific transformation stages and at defined sites that will ensure the binding to and penetration of the ZP. The asynchronous nature of the reaction may result in large-scale redundancy, because only the sperm in the right place at the right time will be able to penetrate the ZP and fertilize the oocyte. The in vivo situation appears to promote the probability of fertilization by ensuring that the maximum possible numbers of functionally competent spermatozoa reach the oocyte at the correct stage of capacitation. We have shown that 14% of cases with unexplained infertility may have an impaired ZIAR, and should be treated with ICSI rather than IVF. The ZIAR53 or DZPIAR72 test has true diagnostic potential, as it can assist the clinician in identifying couples who will benefit from ICSI therapy. In the clinical management of infertility, allocation of patients between standard IVF and ICSI is mainly decided on the basis of specific sperm characteristics that play a role during fertilization. Patients with impaired sperm–zona interaction, i.e. zona pellucida binding and zona-induced acrosome reaction, have a higher success rate in the ICSI laboratory compared with IVF treatment53,73. Moreover, the implementation of these functional tests in the early stages of the work-up of men with subfertile basic sperm parameters or unexplained infertility should allow identification of those cases that ought to be directed to ICSI, avoiding loss of time secondary to the use of less successful options such as intrauterine insemination therapy.

REFERENCES 1. Oliphant G, Reynolds ALB, Thomas TS. Sperm surface components involved in the control of the acrosome reaction. Am J Anat 1985; 174: 269 2. Meizel S. Molecules that initiate or help stimulate the acrosome reaction by their interaction with mammalian sperm surface. Am J Anat 1985; 174: 285 3. Langlais J, Roberts KD. A molecular membrane model of sperm capacitation and the acrosome reaction of mammalian spermatozoa. Gamete Res 1985; 12: 183

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4. Kanwar KC, Yanagimachi R, Lopata A. Effects of human seminal plasma on fertilization capacity of human spermatozoa. Fertil Steril 1979; 31: 321 5. Zaneveld LJD, et al. Human sperm capacitation and the acrosome reaction. Hum Reprod 1991; 6: 1265 6. Yanagimachi R. In vitro sperm capacitation and fertilization of golden hamster eggs in a chemically defined medium in in vitro fertilization and embryo transfer. In Hafez ESE, Semm K, eds. In Vitro Fertilization and Embryo Transfer. New York: Alan Liss, 1982: 65 7. Fleming AD, Yanagimachi R. Effect of various lipids on the acrosome reaction and fertilizing capacity of guinea pig spermatozoa, with special reference to the possible involvement of lysophospholipids in the acrosome reaction. Gamete Res 1981; 4: 253 8. Murphy SJ, Roldan ERS, Yanagimachi R. Effects of extracellular cations and energy substrates on the acrosome reaction of precapacitated guinea pig spermatozoa. Gamete Res 1986; 14: 1 9. Katz OF, Cherr GN, Lambert H. The evolution of hamster sperm motility during capacitation and interaction with the ovum vestments in vitro. Gamete Res 1986; 14: 333 10. Yanagimachi R. Calcium requirements for sperm–egg fusion in mammals. Biol Reprod 1978; 19: 949 11. Yanagimachi R, Usui N. Calcium dependence of the acrosome reaction and activation of guinea pig spermatozoa. Exp Cell Res 1974; 89: 161 12. Stock CE, Fraser LR. Divalent cations, capacitation and the acrosome reaction in human spermatozoa. J Reprod Fertil 1989; 87: 463 13. White DR, Phillips DM, Bedford JM. Factors affecting the acrosome reaction in human spermatozoa. J Reprod Fertil 1990; 90: 71 14. Nagae T, et al. Acrosome reaction in human spermatozoa. Fertil Steril 1986; 45: 701 15. Stock CE, et al. Human oocyte–cumulus complexes stimulate the acrosome reaction. J Reprod Fertil 1989; 86: 723 16. Yudin AI, Gottlieb W, Meizel S. Ultrastructural studies of the early events of the human sperm acrosome reaction as initiated by human follicular fluid. Gamete Res 1988; 20: 11 17. Fraser L. p-Aminobenzamidine, an acrosin inhibitor, inhibits mouse sperm penetration of the zona pellucida but not the acrosome reaction. J Reprod Fertil 1982; 66: 185 18. Jamil K, White IG. Induction of acrosomal reaction in sperm with ionophore A23187 and calcium. Arch Androl 1981; 7: 283

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19. Cummins JM, et al. A test of the human sperm acrosome reaction following ionophore challenge: relationship to fertility and other seminal parameters. J Androl 1991; 12: 98 20. Sanchez R. A new method for evaluation of the acrosome reaction in viable human spermatozoa. Andrologia 1991; 23: 197 21. Aitken RJ, Buckingham DW, Fang HG. Analysis of the response of human spermatozoa to A23187 employing a novel technique for assessing the acrosome reaction. J Androl 1993; 14: 132 22. Tesarik J, Mendoza C, Carreras A. Effects of phosphodiesterase inhibitors caffeine and pentoxifylline on spontaneous and stimulus-induced acrosome reactions in human sperm. Fertil Steril 1992; 58: 1185 23. Tesarik J, Mendoza C. Sperm treatment with pentoxifylline improves the fertilizing ability in patients with acrosome reaction insufficiency. Fertil Steril 1993; 60: 141 24. Osman RA, et al. Steriod induced exocytosis: the human sperm acrosome reaction. Biochem Biophys Res Commun 1989; 160: 828 25. Blackmore PF, et al. Progesterone and 17-hydroxprogesterone: novel stimulators of calcium influx in human sperm. J Biol Chem 1990; 265: 1376 26. Calvo L, et al. Acrosome reaction inducibility predicts fertilization success at in-vitro fertilization. Hum Reprod 1994; 9: 1880 27. Sueldo CE, et al. Effect of progesterone on human zona pellucida sperm binding and oocyte penetrating capacity. Fertil Steril 1993; 60: 137 28. Suarez SS, Wolf DP, Meizel S. Induction of the acrosome reaction in human spermatozoa by a fraction of human follicular fluid. Gamete Res 1986; 14: 107 29. Cross NL, et al. Induction of acrosome reactions by the human zona pellucida. Biol Reprod 1988; 38: 235 30. Lee MA, Check LH, Kopf GA. Guanine nucleotidebinding regulatory protein in human sperm mediates acrosomal exocytosis induced by the human zona pellucida. Mol Reprod 1992; 31: 78 31. Franken DR, Morales PJ, Habenicht UF. Inhibition of G protein in human sperm and its influence on acrosome reaction and zona pellucida binding. Fertil Steril 1996; 66: 1009 32. Talbot P, Chacon RS. A new technique for evaluating normal acrosome reactions of human sperm. J Cell Biol 1979; 83: 2089 33. Kohn FM, et al. Detection of human sperm acrosome reaction: comparison between methods using double staining, Pisum sativum agglutinin, concanavalin A

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and transmission electron microscopy. Hum Reprod 1997; 12: 714 Lee MA, et al. Capacitation and acrosome reactions in human spermatozoa monitored by a chlortetracycline fluorescence assay. Fertil Steril 1987; 48: 649 Talbot P, Chacon RS. A triple-stain technique for evaluating normal acrosome reactions of human sperm. J Exp Zool 1981; 215: 201 Cross NL, et al. Two simple methods for detecting acrosome-reacted human sperm. Gamete Res 1986; 15: 213 Wolf DP, et al. Acrosomal status evaluation in human ejaculated sperm with monoclonal antibodies. Biol Reprod 1985; 32: 1157 Fenichel P, et al. Evaluation of the human sperm acrosome reaction using a monoclonal antibody, GB24, and fluorescence-activated cell sorter. J Reprod Fertil 1989; 87: 699 Aitken RJ, Brindle JP. Analysis of the ability of three probes targeting the outer acrosomal membrane or acrosomal contents to detect the acrosome reaction in human spermatozoa. Hum Reprod 1993; 8: 1663 Bastias MC, Kamijo H, Osteen KG. Assessment of human sperm functional changes after in vitro coincubation with cells retrieved from the human female reproductive tract. Hum Reprod 1993; 8: 1670 Siegel MS, Paulson RJ, Graczykowski JW. The influence of human follicular fluid on the acrosome reaction, fertilizing capacity and proteinase activity of human spermatozoa. Hum Reprod 1990; 5: 975 Thomas P, Meizel S. An influx of extracellular calcium is required for the initiation of the human sperm acrosome reaction induced by human follicular fluid. Gamete Res 1988; 20: 397 Fukuda M, et al. Correlation of the acrosomal status and sperm performance in the sperm penetration assay. Fertil Steril 1989; 52: 836 Stock CE, et al. Extended exposure to follicular fluid is required for significant stimulation of the acrosome reaction in human spermatozoa. J Reprod Fertil 1989; 86: 401 Foresta C, et al. Progesterone induces capacitation in human spermatozoa. Andrologia 1991; 24: 33 Baldi E, et al. Intracellular calcium accumulation and responsiveness to progesterone in capacitating human spermatozoa. J Androl 1991; 12: 323 Luconi M. Human spermatozoa as a model for studying membrane receptors mediating rapid nongenomic effects of progesterone and estrogens. Steroids 2004; 69: 553 Morales P, et al. The acrosome reaction-inducing activity of individual human follicular fluid samples is

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highly variable and is related to the steroid content. Hum Reprod 1992; 7: 646 Miska W, Henkel R, Fehl P. Recent studies on the structure of the acrosome reaction-inducing substance (ARIS) in human follicular fluid. Mol Reprod Dev 1995; 42: 80 Miska W, Fehl P, Henkel R. Biochemical and immunological characterization of the acrosome reaction inducing substance (ARIS) of hFF. Biochem Biophys Res Commun 1994; 199: 125 Franken DR, Bastiaan HS, Oehninger SC. A microassay for sperm acrosome reaction assessment. J Assist Reprod Genet 2000; 17: 156 Bastiaan HS. Zona pellucida induced acrosome reaction (ZIAR), sperm morphology and sperm–zona binding assessments among subfertile men. J Assist Reprod Genet 2002; 19: 329 Esterhuizen AD, et al. Clinical importance of zona pellucida induced acrosome reaction (ZIAR test) in cases of failed human fertilization. Hum Reprod 2001; 16: 138 Saling PM, Storey BT. Mouse gamete interactions during fertilization in vitro: chlortetracycline as a fluorescent probe for the mouse acrosome reaction. J Cell Biol 1979; 83: 544 Saling PM, Irons G, Waibel R. Mouse sperm antigens that participate in fertilization. I. Inhibition of sperm fusion with the egg plasma membrane using monoclonal antibodies. Biol Reprod 1985; 33: 515 Bleil DJ, Wassarman PM. Sperm–egg interactions in the mouse: sequence of events and induction of the acrosome reaction by a zona pellucida glycoprotein. Dev Biol 1983; 95: 317 Coddington CC, et al. Sperm bound to zona pellucida in hemizona assay demonstrate acrosome reaction when stained with T-6 antibody. Fertil Steril 1990; 54: 504 Hoshi KH, et al. Induction of the acrosome reaction in human spermatozoa by human pellucida and effect of cervical mucus on zona-induced acrosome reaction. Fertil Steril 1993; 60: 149 Roldan ERS, Murase T, Shi Q-X. Exocytosis in spermatozoa in response to progesterone and zona pellucida. Science 1994; 266: 1578 Schuffner AA, et al. Zona pellucida-induced acrosome reaction in human sperm: dependency on activation of pertussis toxin-sensitive G(i) protein and extracellular calcium, and priming effect of progesterone and follicular fluid. Mol Hum Reprod 2002; 8: 722 Cummins JM, Yanagimachi R. Development of the ability to penetrate the cumulus oophorus by hamster

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spermatozoa capacitated in vitro in relation to the timing of the acrosome reaction. Gamete Res 1986; 15: 187 Yanagimachi R, Phillips DM. The status of acrosomal caps of hamster spermatozoa immediately before fertilization in vivo. Gamete Res 1984; 9: 1 Cherr GN, Lambert H, Katz D. Completion of the hamster sperm acrosome reaction on the zona pellucida in vivo. J Cell Biol 1984; 99: 261 Tesarik J. Appropriate timing of the acrosome reaction is a major requirement for the fertilizing spermatozoon. Hum Reprod 1989; 4: 957 Carrell DT, et al. Role of the cumulus in the selection of morphologically normal sperm and induction of the acrosome reaction during human in vitro fertilization. Arch Androl 1993; 31: 133 Corselli J, Talbot P. An in vitro technique to study penetration of hamster oocyte–cumulus complexes by using physiological numbers of sperm. Gamete Res 1986; 13: 293 Barros C, et al. Relationship between the length of sperm preincubation and zona penetration in the golden hamster: a scanning electron microscopy study. Gamete Res 1984; 9: 31

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68. Singer SL, et al. The kinetics of human sperm binding to the human zona pellucida and zona-free hamster oocyte in vitro. Gamete Res 1985; 12: 29 69. Pampiglione JS, Tan S, Cambell S. The use of the stimulated acrosome reaction test as a test of fertilizing ability in human spermatozoa. Fertil Steril 1993; 59: 1280 70. Henkel R, et al. Determination of the acrosome reaction in human spermatozoa is predictive of fertilization in vitro. Hum Reprod 1993; 8: 2128 71. Bastiaan HS, et al. The relationship between zona pellucida induced acrosome reaction (ZIAR), sperm morphology, sperm–zona pellucida binding and in vitro fertilization. Fertil Steril 2003; 79: 49 72. Liu DY, Baker HWG. Disordered acrosome reaction of spermatozoa bound to the zona pellucida: a newly discovered sperm defect causing infertility with reduced sperm–zona penetration and reduced fertilization in vitro. Hum Reprod 1994; 9: 1694 73. Liu DY, Garrett C, Baker HWG. Clinical application of sperm–oocyte interaction tests in in vitro fertilization–embryo transfer and intracytoplasmic sperm injection programs. Fertil Steril 2004; 82: 1251

14 Sperm–zona pellucida binding assays Sergio Oehninger, Murat Arslan, Daniel R Franken

BIOLOGY OF FERTILIZATION

dependent upon interaction of complementary gamete molecules), oocyte activation, nuclear decondensation and participation in pronuclear formation leading to syngamy (reviewed in reference 2).

Obligatory requirements for the successful completion of normal fertilization include a mature, metaphase II oocyte and motile spermatozoa that have completed the process of capacitation. The newly formed zygote undergoes early cleavage divisions depending upon the oocyte’s endogenous machinery, and at the 4–8-cell stage initiates transcription of the embryonic genome1. In vivo, these processes are synchronized with the preparation of the endometrial mucosa (window of implantation), thereby ensuring an adequate milieu receptive to the blastocyst. Spermatozoa are highly differentiated cells whose main function is to activate the oocyte and deliver components, principally its DNA, leading to embryo development. In order to fertilize the oocyte successfully, the spermatozoon must be able to perform, at least, these functions: migration (allowing transport to the fertilization site through adequate motion patterns), recognition and binding to the zona pellucida (an event dependent upon specific receptor–ligand interactions), penetration of the zona pellucida (secondary to the release of enzymes following induction of the acrosome reaction by zona components), binding to the oolemma (also

Events leading to sperm–oocyte interaction Only capacitated spermatozoa demonstrate the ability to respond to the adequate physiological stimuli that result in the display of adequate motion characteristics, acrosome reaction responsiveness and competence to interact with the oocyte and its vestments. Several cellular changes that are manifested during capacitation include, among others, removal or modification of surface proteins, efflux of cholesterol from the membranes, changes in oxidative metabolism, achievement of a hyperactivated pattern of motility and an increase in the phosphotyrosine content of several proteins. In addition to tyrosine phosphorylation of specific proteins, other modifications of cellular regulators occur, such as a decrease in calmodulin binding to proteins and an increase in calcium uptake, intracellular pH and cyclic adenosine monophosphate (cAMP) concentration2,3.

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Sperm–zona pellucida interaction: recognition, binding and induction of the acrosome reaction The early events that occur during fertilization may be viewed as a special form of highly complex cell-to-cell recognition. Cell–cell recognition mechanisms in many somatic cell systems involve carbohydrate side-chains of membrane glycoproteins, and several observations indicate that similar molecules may have a role in spermatozoon– oocyte binding in mammals. Compelling evidence has now demonstrated that carbohydrate-binding proteins on the sperm surface mediate gamete recognition by binding with high affinity and specificity to complex glycoconjugates of the zona pellucida2,4–6. In the mouse, the best characterized species so far, tight binding is achieved through interaction of zona pellucida protein 3 (ZP3) and a putative complementary sperm-binding protein(s) present in the plasma membrane. ZP3 triggers the acrosome reaction that is then followed by a secondary binding process involving zona pellucida protein 2 (ZP2) and the inner acrosomal sperm membrane, leading to zona penetration7,8. Glycosylation appears to be mandatory for ZP3-ligand function. It has been demonstrated that O-glycosylation, and particularly terminal galactose residues of Olinked oligosaccharides, are essential for maintaining mouse gamete interaction. There is also some evidence that the amino sugar N-acetylglucosamine (NAG) is the key terminal monosaccharide involved in sperm–zona interaction in the mouse9,10. In contrast, the acrosome reaction-triggering activity of ZP3 seems to depend upon the integrity of the protein backbone (reviewed in references 5, 11 and 12). Peptides synthesized based upon the published DNA sequence of ZP3 proteins are able to induce acrosomal exocytosis in some species13. The molecular identity of the sperm surface receptor(s) for ZP3 has been the subject of intensive research. A number of candidate murine ZP3 receptor molecules have been proposed,

including potential carbohydrate-binding proteins such as sp56, p95, β-1-4 galactosyltransferase and a D-mannosidase9,14–18. However, there has been confirmation neither of the structure or biological role of any of these molecules nor of their complementary ligand(s). The state of knowledge as related to the human is even more enigmatic. In the mouse, ZP3-binding and ZP3-induced acrosomal exocytosis can be dissociated from each other, that is, they seem to represent two independent processes19. There are differences in the concentration-dependency of ZP3 to express sperm-binding activity and acrosome reactioninducing activity. Specifically, the concentration response curve for ZP3 acrosome reaction-inducing activity is shifted to the right of the concentration response curve for ZP3-ligand activity. A model has been proposed predicting that ZP3 is composed of multiple ‘functional ligands’, and that the interaction of these ligands with the sperm surface is responsible for both the spermbinding activity (through glycosylated epitopes) and the ability to induce a complete acrosome reaction19. Gamete recognition and adhesion probably depend upon a multivalent ligand interaction whereby the sperm protein receptor(s) bind to a number of different epitopes within the ZP3. These functional ligands do not necessarily have to be identical. The data concerning the involvement of either O- or N-linked glycosylation sites are also equivocal, particularly in the human. The lack of native human zona pellucida to perform direct carbohydrate analyses has made an unambiguous structural definition impossible so far. We have proposed the hypothesis that, in the human, tight and specific sperm binding to the zona pellucida requires a ‘selectin-like’ interaction6,20. Hapten-inhibition tests, zona pellucida lectin-binding studies and removal/modification of functional carbohydrates by chemical and enzymatic methods have provided evidence for the involvement of defined carbohydrate moieties in initial binding. Our studies suggest the existence

SPERM–ZONA PELLUCIDA BINDING ASSAYS

of distinct zona-binding proteins on human sperm that can bind to selectin ligands (reviewed in reference 21). Additionally, results suggest a possible convergence in the types of carbohydrate sequences recognized during initial human gamete binding and immune/inflammatory cell interactions (reviewed in reference 22). Full characterization of the glycoconjugates that manifest selectinligand activity on the human zona pellucida will allow a better understanding of human gamete interaction in physiological and pathological situations. Nevertheless, determination of the biochemical components and secondary structure of the human zona proteins has been hampered by the paucity of biological material. For the past two decades, investigators have sought to identify an individual protein or carbohydrate side-chain as the ‘sperm receptor’. Using ‘knock-out’ mice, in the absence of either ZP2 or ZP3 expression, a zona pellucida fails to assemble around growing oocytes and females are infertile. In the absence of ZP1 expression, a disorganized zona assembles around growing oocytes and females exhibit reduced fertility. These observations are consistent with the current model for zona pellucida structure in which ZP2 and ZP3 form long Z-filaments crosslinked by ZP1 (reviewed in reference 23). However, recent genetic data in mice appear to be more consistent with the three-dimensional structure of the zona pellucida, rather than a single protein (or carbohydrate), determining sperm binding. Collectively, the genetic data indicate that no single mouse zona-pellucida protein is obligatory for taxon-specific sperm binding, and that two human proteins are not sufficient to support human sperm binding. An observed postfertilization persistence of mouse sperm-binding to ‘humanized’ zona pellucida correlates with uncleaved ZP2. These observations are consistent with a model for sperm binding in which the supramolecular structure of the zona pellucida necessary for sperm binding is modulated by the cleavage status of ZP224–27.

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Post-zona pellucida binding events: interaction between sperm and oocyte leading to fusion, oocyte activation, pronuclear formation and paternal contribution to early embryogenesis Spermatozoa that have undergone the acrosome reaction after interaction with and penetration through the zona pellucida are able to bind to the plasma membrane of the oocyte (oolemma). This also seems to be a specific recognition event involving putative molecules located in the equatorial segment of the sperm (sperm fusion proteins) and yet-unidentified oocyte acceptors. Binding of the gametes leads to fusion of the membranes with incorporation of the entire spermatozoon into the ooplasm. Contact of the spermatozoon with the oocyte membrane triggers electrical membrane changes in the oocyte (membrane depolarization) and the release of cortical granules, which represent fast and delayed protective mechanisms against polyspermy (reviewed in reference 2). There is still controversy as to the intimate mechanism(s) through which the spermatozoon activates the oocyte. Oocyte activation occurs in association with changes in the intracellular concentration of calcium ions, possibly modulated by a factor released by the spermatozoon once inside the oocyte. Unequivocal identification of this factor in the human and other species has not yet been achieved28. Sperm–oolemma binding and fusion are followed by activation of the oocyte’s second-messenger systems (calcium, phosphatidylinositol-4,5-biphosphate (PIP2)), pH changes, protein synthesis, cyclin accumulation, DNA synthesis, nuclear envelope breakdown and the first cleavage division in some species. An increase in intracellular calcium is associated with microtubular rearrangement and pronuclear formation2. There is obviously extensive crosstalk between the spermatozoon and the oocyte. In addition to the effects secondary to membrane fusion and the

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release of oocyte activating factor(s) by the spermatozoon, the oocyte uses molecules that induce sperm head decondensation (male pronucleus growth factor) and the substitution of protamines by histones2,29. Fertilization is achieved after the oocyte completes meiosis, female and male pronuclei are formed and syngamy (pronuclei union) is accomplished.

ABNORMALITIES OF FERTILIZATION: CLINICAL LESSONS FROM IN VITRO FERTILIZATION AND INTRACYTOPLASMIC SPERM INJECTION SETTINGS It has been reported that sperm–zona pellucida binding is a crucial step and that it reflects multiple sperm functions30–32. Many patients who are unable to fertilize oocytes under in vitro fertilization (IVF) conditions have a severe impairment of this functional step. A defective capacity to undergo the acrosome reaction is probably also a significant factor in some patients33. It has been shown recently that acrosomal exocytosis can be studied in vitro using small volumes of solubilized human zonae pellucidae and that Gproteins are involved as mediators34. This confirms previous studies that demonstrated the involvement of heterotrimeric G-proteins in induction of the acrosome reaction in other species19. It has also been demonstrated that functional/biochemical/morphological sperm immaturity (e.g. high content of creatine kinase) is present in many cases of male infertility, resulting in fertilization deficiencies35. In addition to a defective sperm–zona pellucida interaction, fertilization failure can also be due to sperm–oolemma fusion defects or to abnormal communication between the penetrating spermatozoon and the oocyte (e.g. lack or deficient sperm–oocyte activating factor, male pronucleus growth factor or other). Recent evidence from the intracytoplasmic sperm injec-

tion (ICSI) setting clearly demonstrates that post-gamete fusion abnormalities may occur. Advances in fluorescent imaging by laser scanning confocal microscopy and other novel techniques permit the sophisticated high-resolution examination of gametes and embryos, including the fate of the sperm centrosome, the oocyte’s microtubule organizing center, mitochondrial distribution and the initiation of embryo cleavage36. We remain enthusiastic about ongoing studies that may help to elucidate the contribution of the gametes (functional, biochemical–molecular and genetic) to early embryogenesis, and identify specific molecules involved in fertilization disorders.

CLINICAL ASPECTS: MALE SUBFERTILITY AND SEMEN EVALUATION Men consulting for infertility which is defined as male-factor typically present abnormalities of semen analysis consistent with varying degrees of oligoasthenoteratozoospermia, alone or in combination. In addition, other structural and biochemical sperm alterations can be demonstrated. From an anatomical point of view they can be divided into: membrane alterations (that can be assessed by tests of resistance to osmotic changes, translocation of phosphatidylserine and others), nuclear aberrations (abnormal chromatin condensation, retention of histones and presence of DNA fragmentation), cytoplasmic lesions (excessive generation of reactive oxygen species, loss of mitochondrial membrane potential or retention of cytoplasm, indicative of immaturity such as high creatine kinase content or presence of caspases) and flagellar disturbances (disturbances of the microtubules and the fibrous sheath). Some of these alterations are indicative of immaturity, the presence of an apoptosis phenotype, infectionnecrosis or other unknown causes (reviewed in references 37–43). Notwithstanding their occurrence and weak correlations with clinical outcomes, it is not clear

SPERM–ZONA PELLUCIDA BINDING ASSAYS

how these abnormalities impact directly on sperm function, particularly gamete transportation, fertilization and contribution to embryogenesis. Furthermore, most such assays are still experimental, and more research is needed to validate their results in the clinical setting and to determine their true capacity to predict male fertility potential. On the other hand, there are other specific and critical sperm functional capacities that can be more reliably examined in vitro. These functions include: motility, competence to achieve capacitation, zona pellucida binding, acrosome reaction, oolemma binding, decondensation and pronuclear formation. The assessment of some of these features is what is typically considered as sperm functional testing.

VALIDITY OF SPERM FUNCTION ASSAYS: RESULTS OF A META-ANALYSIS The categories of functional assays that are usually considered include: (1) bioassays of gamete interaction (e.g. the heterologous zona-free hamsteroocyte test and homologous sperm–zona pellucida binding assays); (2) induced acrosome-reaction testing; and (3) computer-aided sperm motion analysis (CASA) for the evaluation of sperm motion characteristics33,44–50. We recently reported an objective, outcomebased examination of the validity of the currently available assays based upon the results obtained from 2906 subjects evaluated in 34 published and prospectively designed, controlled studies. The aim was carried out through a meta-analytical approach that examined the predictive value of four categories of sperm functional assays (computer-aided sperm motion analysis or CASA, induced acrosome-reaction testing, sperm penetration assay or SPA and sperm–zona pellucida binding assays) for IVF outcome51. Results of this meta-analysis demonstrated a high predictive power of the sperm–zona pellucida

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binding and induced acrosome-reaction assays for fertilization outcome under in vitro conditions51. On the other hand, the findings indicated a poor clinical value of the SPA as a predictor of fertilization, and a real need for standardization and further investigation of the potential clinical utility of CASA systems. Although this study provided objective evidence in which clinical management and future research may be directed, the analysis also pointed out limitations of the current tests, and the need for standardization of present methodologies and the development of novel technologies. It is important to note that there are no studies addressing the validity and predictive power of these assays for natural conception.

DESIGN OF IN VITRO SPERM–ZONA PELLUCIDA BINDING ASSAYS Our group has published extensively on the development and validation of an in vitro bioassay (the hemizona assay or HZA) for the assessment of tight human sperm binding to the homologous zona pellucida. The initial studies were based on the hypothesis that capacitated spermatozoa bind in a specific, tight and irreversible manner to the homologous, biologically intact zona pellucida, and undergo a physiologically induced acrosome reaction (exocytosis triggered by components of the zona pellucida). This hypothesis was tested by incubation of spermatozoa and the zona pellucida from microbisected human oocytes, followed by determination of the kinetics, sperm concentration-, sperm morphology- and time-dependency of binding, and sperm acrosomal status on tight binding. The HZA was introduced as a novel diagnostic test for the binding of human spermatozoa to the human zona pellucida to predict fertilization potential46. In the HZA, the two matched zona hemispheres created by microbisection of the human oocyte provide three main advantages: (1) the two halves (hemizonae) are functionally equal

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surfaces, allowing controlled comparison of binding and reproducible measurements of sperm binding from a single egg; (2) the limited number of available human oocytes is amplified because an internally controlled test can be performed on a single oocyte; and (3) because the oocyte is split microsurgically, use of even fresh oocytes cannot lead to inadvertent fertilization and pre-embryo formation46,52. The two most common zona binding tests currently used are the HZA46 and a zona pellucidabinding test32,47. Both bioassays have the advantage of providing a functional homologous test for sperm binding to the homologous zona pellucida, comparing populations of fertile and infertile spermatozoa in the same assay. The internal control offered by the HZA represents an advantage by decreasing the number of oocytes needed during the assay and diminishing the intra-assay variation46,52–57. Different sources of human oocytes can be used in the assay: oocytes recovered from surgically removed ovaries or postmortem ovarian tissue, and surplus oocytes from the IVF program. Since fresh oocytes are not always available for the test, various alternatives have been implemented for storage. Others have described the storage of human oocytes in dimethylsulfoxide (DMSO) at ultralow temperatures58. Additionally, Yanagimachi and colleagues showed that highly concentrated salt solutions provided effective storage of hamster and human oocytes such that the spermbinding characteristics of the zona pellucida were preserved59,60. In developing the HZA, we have examined the binding ability of fresh and DMSOand salt-stored (under controlled pH conditions) human oocytes, and have concluded that the sperm binding ability of the zona remains intact under all these conditions53,61. Subsequently, we have assessed the kinetics of sperm binding to the zona, showing maximum binding at 4–5 h of gamete coincubation, with similar binding curves for both fertile and infertile semen samples46,53. Detailed descriptions of oocyte collection, handling and micromanipulation, as well as

semen processing and sperm suspension preparations for the HZA, have been published elsewhere46,53. The assay has been standardized to a 4-h gamete coincubation, exposing each hemizona to a sperm droplet (50–100 µl of a dilution of 500 000 motile sperm/ml prepared after swimup). Human tubal fluid supplemented with synthetic serum substitute or human serum albumin is usually the medium utilized for sperm preparation and gamete coincubation. After coincubation, the hemizonae are subjected to pipetting through a glass pipette in order to dislodge loosely attached sperm. The number of tightly bound spermatozoa on the outer surface of the zona is finally counted using phase-contrast microscopy (200×). Results are expressed as the number of sperm tightly bound to the hemizona for controls and patients, and also as the hemizona index (HZI), i.e. the number of sperm tightly bound, for the control sample (×100)46. The assay has been validated by a clear-cut definition of the factors affecting data interpretation, i.e. kinetics of binding, egg variability and maturation status, intra-assay variation and influence of sperm concentration morphology, motility and acrosome reaction status53–55,57,62,63. Because of the definition of the assay’s limitations and its small intra-assay variation (less than 10%), the power of discrimination of the HZA has been maximized. Conversely, for other sperm–zona binding tests, several oocytes have to be used because of the high inter-egg variation, and in fact a high intra-assay coefficient of variation has been reported32,47. The specificity of the interaction between human spermatozoa and the human zona pellucida under HZA conditions is strengthened by the fact that the sperm tightly bound to the zona are acrosome-reacted54,62. Results of interspecies experiments performed with human, cynomolgus monkey and hamster gametes have demonstrated a high species specificity of human sperm–zona pellucida functions under HZA conditions, providing further support for the use of this bioassay in infertility and contraception testing64.

SPERM–ZONA PELLUCIDA BINDING ASSAYS

In prospective, blinded studies, we have investigated the relationship between sperm binding to the hemizona and IVF outcome31,56,65–67. Results have shown that the HZA can successfully distinguish the population of male-factor patients at risk for failed or poor fertilization (Figure 14.1). Powerful statistical results allow use of the HZA for prediction of the fertilization rate67–70. The HZA can distinguish a population of malefactor patients who will encounter low fertilization rates in IVF, and, when combined with the information provided by other sperm parameters (morphology and motion characteristics), gives reliable and useful information in the clinical arena. Of the basic sperm parameters, sperm morphology is the best predictor of the ability of spermatozoa to bind to the zona pellucida. Sperm from patients with severe teratozoospermia (‘poorprognosis’ pattern or less than 4% normal sperm scores as judged by strict criteria) have an impaired capacity to bind to the zona pellucida under HZA conditions (membrane/receptor deficiencies?). In our studies, when the HZA was removed from the regression analysis in order to identify the predictive value of other sperm parameters (sperm concentration, morphology and motion characteristics), percentage progressive motility was the second best predictor of in vitro fertilization outcome31. We speculated that the relationship between sperm morphology and IVF results depends upon an effect on zona pellucida binding. On the other hand, motility seemed to affect the rate of fertilization outside the prediction of the HZA. It would appear that, although important in achieving binding, motility may be more important for cumulus and zona pellucida penetration, factors not directly evaluated in the HZA. Logistic regression analysis provided a robust HZI range predictive of the oocyte’s potential to be fertilized. This HZI cut-off value was approximately 35%. Overall, for failed vs. successful and

poor vs. good fertilization rate, the correct predictive ability (discriminative power) of the HZA was 80% and 85%, respectively. Consequently, this information may be extremely valuable for counseling patients in the IVF setting (for example, considering a HZI below 35%, the chances of poor fertilization are 90–100%, whereas for a HZI over 35%, the chances of good fertilization are 80–85%) (Figure 14.1)31,67,68,70. The HZA has demonstrated excellent sensitivity and specificity with a low incidence of falsepositive results. For a HZI of 35%, the positive predictive value of the HZA is 79% and its negative predictive value is 100% (considering good vs. poor fertilization rates). In the HZA, false-positive results can be expected, since other functional steps follow the tight binding of sperm to the zona pellucida and are essential for fertilization and preembryo development.

100 90 Rate of oocyte fertilization in IVF (%)

PREDICTIVE VALUE OF THE HEMIZONA ASSAY FOR IN VITRO FERTILIZATION OUTCOME

215

80 70 60 50 40 Good fertilization

Poor or failed fertilization False positive results

20 10 10

100

Hemizona index (%)

Figure 14.1 Cluster analysis of hemizona index (HZI) and rate of fertilization of mature oocytes in in vitro fertilization (IVF) considering a cut-off HZI of 35%. Cluster : high HZI, successful fertilization in the range 50–100% of oocytes; cluster : low HZI, failed or poor fertilization in the range 0–50% of oocytes; cluster : false-positive results with high HZI but failed fertilization (< 15% of cases). For patients undergoing IVF treatment, and if HZI results are < 35%, the chances of poor fertilization are 90–100%, whereas for HZI results > 35%, the chances of good fertilization are 80–85%. Note the absence of false-negative results and evidence of 15% false-positive results. The cluster analysis was performed with combined data from references 56 and 66–68

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PREDICTIVE VALUE OF THE HEMIZONA ASSAY FOR PREGNANCY OUTCOME IN INTRAUTERINE INSEMINATION THERAPY The prediction of pregnancy in intrauterine insemination (IUI) cycles has been expected to be much more difficult than prediction of fertilization in IVF. This is due to the multifactorial nature of conception, as it depends upon the presence of many sperm functions and additional female parameters. For IUI therapy, the most significant female parameters are the quality or quantity of the oocyte(s) and the transportation of capacitated sperm to the fertilization site (e.g. effects of uterine and tubal environment, IUI preparation technique) (reviewed in reference 71). However, there are also other potential and more subtle female factors, such as exposure of spermatozoa to peritoneal and follicular fluid, that have been found to affect sperm binding to the zona pellucida and the ability to respond to physiological inducers of the acrosome reaction72,73. Sperm–zona pellucida binding is a crucial and common step in the journey leading to fertilization during both in vivo and in vitro models. We therefore tested the power of the HZA to predict pregnancy outcome in patients undergoing IUI therapy using the husband’s sperm. Only couples with a diagnosis of unexplained infertility and male factor infertility were asked to participate in the clinical trial. During a 3-year span, 82 couples who underwent 313 IUI treatment cycles and who were categorized into unexplained/male factor infertility agreed to participate. The male partner had a HZA within 3 months of the first IUI cycle, and couples underwent 1–6 IUI cycles within the next 12-month period. All female patients were subjected to controlled ovarian hyperstimulation using a similar gonadotropin protocol. For all patients involved in the study, the HZI results ranged between 0 and 178%. Minimum and maximum HZI values that achieved a pregnancy were 17% and 109%, respectively. When

we analyzed the data according to a 30–35% cut-off HZI range, which was proven optimum for prediction of successful fertilization in IVF66,67, the HZA had a high negative predictive value (NPV) of almost 90% (i.e. patients with a HZI < 30% had a very low chance of conception) (Table 14.1). On the other hand, results demonstrated that the positive predictive value (PPV) of the test decreased in parallel with its NPV with increasing cut-off values (r = –0.7, p < 0.05 and r = –0.8, p < 0.05 for PPV and NPV, respectively). This was reflected as increased false-positive rates with higher HZI values (Figure 14.2). This result confirmed that a variety of pre- and postsperm–zona pellucida binding factors play an active role in establishing a pregnancy: patients with high HZI values still may not be able to achieve conception. In light of these findings, we re-examined the data in the range of HZI between 0 and 60%. This approach was also used in earlier IVF studies, where it was confirmed that successful fertilization occurred in nearly all patients with a HZI > 60% under optimal IVF conditions74. With a HZI cutoff value of 30–35%, we found a relatively higher PPV of 69%, but still a high incidence of falsepositive results, with a very high negative predictive value of 93% (Table 14.1). The data were also subjected to receiver operating characteristic (ROC) analysis to assess the contributions of all male and female parameters, for the overall population and also after categorization of patients according to the subgroup of etiology (male factor or unexplained). In this model, a HZI with cut-off value of 32% demonstrated significant power for the prediction of pregnancy in the male factor infertility subgroup, with 69% PPV and 93% NPV (p = 0.005). The average path velocity was the second male-factor parameter that had significance as predictor in this subgroup (30% PPV and 95% NPV with cut-off value of 46.5 µm/s, p = 0.001). The duration of infertility was a strong predictor of pregnancy in all patients and in both subgroups. Binary logistic regression analysis applied to all male and female

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SPERM–ZONA PELLUCIDA BINDING ASSAYS

All patients

Patients with HZI value between 0 and 60%

Positive predictive value

100

Negative predictive value

80 Predictive value

Table 14.1 Predictive value of hemizona assay (HZA) for pregnancy with intrauterine insemination (IUI) therapy considering a hemizona index (HZI) cut-off range of 30–35%. Predictive values were calculated for all patients and for patients who had HZI results in the range 0–60%

60 40 20 0 0

Positive predictive value (%) False-positive rate (%) False-negative rate (%) Negative predictive value (%)

parameters also confirmed the HZI as the most powerful and single parameter predictive of conception in couples with a diagnosis of male factor infertility (–2 log likelihood of 28.778 and χ2 of 7.720, p = 0.005) (Table 14.2). Although this evidence continues to encourage use of the HZA in the screening work-up of consulting couples before starting IUI therapy, larger prospective studies are still needed to confirm these favorable initial results.

PREDICTIVE VALUE OF THE HEMIZONA ASSAY FOR NATURAL CONCEPTION It has been speculated that information from the semen analysis can be used to predict the likelihood that a couple will conceive within a period of time. This probability is influenced by a host of factors including semen quality, and studies in large groups or using simple models are required to overcome existing limitations75. The world literature has consistently used the World Health Organization (WHO) guidelines for normalcy cut-offs to address clinical situations. However, recent studies have raised doubts about such established guidelines.

100

120

Cut-off value for HZI

Figure 14.2 Relationship between different cut-off values of hemizona index (HZI) and corresponding positive and negative predictive values for conception in intrauterine insemination (IUI) therapy

Table 14.2 Results of logistic regression analysis of different sperm parameters and impact on pregnancy outcome in couples with diagnosis of male factor infertility undergoing intrauterine insemination (IUI) therapy r†

p Value

Morphology

0.19

0.07

Concentration

0.00

0.27

Motility

0.00

0.99

HZI

0.30

0.02*

†

Partial contribution; *statistically significant; HZI, hemizona index

In a prospectively designed study, Ombelet and collaborators76 compared a fertile and a subfertile population so as to define ‘normal’ values for different semen parameters. Semen analyses were performed according to WHO guidelines, except for sperm morphology (strict criteria). The authors used ROC curve analysis to determine the diagnostic potential and cut-off values for single and combined sperm parameters. Sperm morphology scored best, with a value of 78% (area under

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the ROC curve). Summary statistics showed a shift towards abnormality for most semen parameters in the subfertile population. Using the 10th centile of the fertile population as the cut-off value, the following results were obtained: 14.3 × 106/ml for sperm concentration, 28% for progressive motility and 5% for sperm morphology. Using ROC analysis, cut-off values were 34 × 106/ml, 45% and 10%, respectively. Cut-off values for normality were different from those described in the last edition of the WHO guidelines. In addition, there are well-known variations in sperm parameters among different ejaculates from the same man, and differences among groups of patients75. To the best of our knowledge, there are scant data, if any at all, on the predictive value of any sperm structural–biochemical feature or sperm function test for the outcome of natural conception. Van der Merwe et al.78 suggest that thresholds of < 5% normal sperm morphology, a concentration < 15 × 106/ml and a motility < 30% should be used to identify the subfertile male. The lower threshold for morphology also fits IVF and IUI data calculated previously. Nevertheless, thresholds for natural conception (highly predictive of pregnancy within a given time-frame) need to be determined for the basic sperm parameters as well as for HZA and other functional tests.

DISCUSSION AND CONCLUSIONS The high negative predictive value, but more important, the low false-negative rate (i.e. robust power to identify patients at high risk for fertilization failure in IVF and to fail conception in IUI), underscore the predictive ability of the HZA in the clinical setting. Liu et al.79 reported that sperm defects associated with poor sperm–zona pellucida binding or impaired zona pellucida-induced acrosome reaction and sperm–zona pellucida penetration are the major causes of failure of fertilization when all or most oocytes from a couple do not fertilize in

standard IVF. These authors further demonstrated that there is a high frequency of defective sperm–zona pellucida interaction in men with oligozoospermia (< 20 × 106/ml) and severe teratozoospermia (strict normal sperm morphology ≤ 5%). According to these authors, sperm morphology correlated with sperm–zona pellucida binding, and sperm concentration correlated with zona pellucida-induced acrosome reaction in infertile men with a sperm concentration > 20 × 106/ml. The authors suggested that a defective zona pellucida-induced acrosome reaction may cause infertility in up to 25% of men with idiopathic infertility. These patients would therefore require ICSI, despite the presence of an otherwise normal standard semen analysis79–83. The induced acrosome-reaction assays appear to be equally predictive of fertilization outcome in vitro as the sperm–zona pellucida binding tests, and are simpler in their methodologies51. Although the use of a calcium ionophore to induce the acrosome reaction is at present the most widely used methodology84,85, the assay uses non-physiological conditions that may not accurately represent fertilization potential. The recent implementation of assays using small volumes of human solubilized zonae pellucidae34,86, biologically active recombinant human ZP35,87,88 or active, synthetic ZP3 peptides13 will probably allow the design of improved, physiologically oriented acrosome reaction assays. Initially, it was believed that cloning of the human ZP3 gene would circumvent the obstacle manifested as a paucity of natural material, since a constant supply of recombinant protein would be available. However, several of the laboratories dedicated to this task have been generally unable consistently and reliably to purify a biologically active product so far (reviewed in reference 5). It seems clear that this is probably due to inadequate and heterogeneous glycosylation of the protein by the different cell lines used. Although we have been able to express and purify a human recombinant ZP3 that appears to demonstrate the full spectrum of biological activities, problems of

SPERM–ZONA PELLUCIDA BINDING ASSAYS

stable transfection, protein storage and maintenance of bioactivity have hampered progress88. Franken et al.86 devised a new microassay that is easy and rapid to perform, and facilitates the use of minimal volumes of solubilized zonae pellucidae (even a single zona) for assessment of the human acrosome reaction. The microassay has been validated against standard macroassays, and consequently offers a unique arena to test for the physiological induction of acrosomal exocytosis by the homologous zona pellucida. Moreover, initial clinical studies using the microassay have demonstrated that the zona-induced acrosome reaction (ZIAR) can predict fertilization failure in the IVF setting. The microassay ZIAR can therefore refine the therapeutic approach for male infertility prior to the onset of therapy89,90. Bastiaan et al.91,92 prospectively evaluated the relationship between sperm morphology, acrosome responsiveness to solubilized zona pellucida using the microassay, sperm–zona binding potential (HZA) and IVF outcome. ROC curve analyses indicated ZIAR to be a robust indicator for fertilization failure during IVF therapy, with a sensitivity of 81% and specificity of 75%. Sperm function tests may be of highest value in order to direct the couple to assisted reproductive technologies (ART). Assisted reproduction is usually indicated as a result of: (1) failure of urological/medical treatments of the subfertile man (if indicated); (2) the diagnosis of ‘unexplained’ infertility in the couple; (3) the presence of ‘basic’ sperm abnormalities of moderate–high degree; or (4) abnormalities of sperm function as diagnosed by predictive bioassays (such as the HZA or ZIAR). Typically, patients are selected for ICSI under the following scenarios: (1) poor sperm parameters (i.e. < 1.5 × 106 total spermatozoa with adequate progressive motility after separation, and/or severe teratozoospermia with < 4% normal forms in the presence of a borderline to low total motile fraction); (2) poor sperm–zona pellucida binding capacity with a hemizona assay index < 30%, and/or low ZIAR67,70,91; (3) failure of IUI therapy

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in cases presenting with abnormal sperm parameters, including the presence of antisperm antibodies; (4) previous failed fertilization in IVF; and (5) the presence of obstructive or non-obstructive azoospermia, where ICSI is combined with sperm extraction from the testes or the epididymis51,93–95. In the presence of severe oligoasthenoteratozoospermia, or if the outcome of sperm function testing indicates a significant impairment of fertilizing capacity, couples should be immediately directed to ICSI. This approach is probably more cost-effective and will avoid loss of valuable time, particularly in women aged > 35 years94,96. More research is needed to develop simpler assays of sperm function that can be clinically useful for the prediction of both in vivo and in vitro pregnancy outcomes. It is expected that advances in molecular biology methodologies and novel biotechnologies will help to achieve this goal.

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for binding human spermatozoa. Gamete Res 1989; 22: 15 Franken DR, et al. Defining the valid hemizona assay: accounting for binding variability within zonae pellucidae and within semen samples from fertile males. Fertil Steril 1991; 56: 1156 Franken DR, et al. Comparison of sperm binding potential of uninseminated, inseminated-unfertilized, and fertilized-noncleaved human oocytes under hemizona assay conditions. Mol Reprod Dev 1991; 30: 56 Oehninger S, Acosta AA, Veeck L. Recurrent failure of in vitro fertilization: role of the hemizona assay (HZA) in the sequential diagnosis of specific sperm/oocyte defects. Am J Obstet Gynecol 1991; 164: 1210 Oehninger S, et al. Human preovulatory oocytes have a higher sperm-binding ability than immature oocytes under hemizona assay conditions: evidence supporting the concept of ‘zona maturation’. Fertil Steril 1991; 55: 1165 Overstreet JW, Hembree WC. Penetration of the zona pellucida of nonliving human oocytes by human spermatozoa in vitro. Fertil Steril 1976; 27: 815 Yanagimachi R, et al. Retention of biologic characteristics of zona pellucida in highly concentrated salt solution: the use of salt-stored eggs for assessing the fertilizing capacity of spermatozoa. Fertil Steril 1979; 31: 562 Yoshimatsu N, Yanagimachi R. Zonae pellucidae of salt-stored hamster and human eggs: their penetrability by homologous and heterologous spermatozoa. Gamete Res 1988; 21: 115 Kruger TF, et al. Hemizona assay: use of fresh versus salt-stored human oocytes to evaluate sperm binding potential to the zona pellucida. J In Vitro Fert Embryo Transf 1991; 8: 154 Coddington CC, et al. Sperm bound to zona pellucida in hemizona assay demonstrate acrosome reaction when stained with T-6 antibody. Fertil Steril 1990; 54: 504 Coddington CC, et al. Functional aspects of human sperm binding to the zona pellucida using the hemizona assay. J Androl 1991; 12: 1 Oehninger S, et al. The specificity of human spermatozoa/zona pellucida interaction under hemizona assay conditions. Mol Reprod Dev 1993; 35: 57 Franken DR, et al. The hemizona assay (HZA): a predictor of human sperm fertilizing potential in in vitro fertilization (IVF) treatment. J In Vitro Fert Embryo Transf 1989; 6: 44

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66. Oehninger S, et al. Hemizona assay: assessment of sperm dysfunction and prediction of in vitro fertilization outcome. Fertil Steril 1989; 51: 665 67. Oehninger S, et al. Clinical significance of human sperm–zona pellucida binding. Fertil Steril 1997; 67: 1121 68. Franken D, et al. The ability of the hemizona assay to predict human fertilization in vitro in different and consecutive IVF/GIFT cycles. Hum Reprod 1993; 8: 1240 69. Gamzu R, et al. The hemizona assay is of good prognostic value for the ability of sperm to fertilize oocytes in vitro. Fertil Steril 1994; 62: 1056 70. Oehninger S, Franken D, Kruger T. Approaching the next millennium: how should we manage andrology diagnosis in the intracytoplasmic sperm injection era? Fertil Steril 1997; 67: 434 71. Duran HE, et al. Intrauterine insemination: a systematic review on determinants of success. Hum Reprod Update 2002; 8: 373 72. Marin-Briggiler CI, et al. Effect of antisperm antibodies present in human follicular fluid upon the acrosome reaction and sperm–zona pellucida interaction. Am J Reprod Immunol 2003; 50: 209 73. Munuce MJ, et al. Modulation of human sperm function by peritoneal fluid. Fertil Steril 2003; 80: 939 74. Coddington CC, et al. Hemizona index (HZI) demonstrates excellent predictability when evaluating sperm fertilizing capacity in in vitro fertilization patients. J Androl 1994; 15: 250 75. Ford WC. Prediction of fecundability from semen analysis: problems in providing an accurate prognosis. Hum Fertil 1999; 2: 25 76. Ombelet W, et al. Semen parameters in a fertile versus subfertile population: a need for change in the interpretation of semen testing. Hum Reprod 1997; 12: 987 77. World Health Organization. WHO Laboratory Manual for the Examination of Human Semen and Sperm–Cervical Mucus Interaction, 4th edn. Cambridge: Cambridge University Press, 1999: 14 78. van der Merwe FH, et al. The use of semen parameters to identify the subfertile male in the general population. Gynecol Obstet Invest 2004; 59: 86 79. Liu de Y, Garrett C, Baker HW. Clinical application of sperm–oocyte interaction tests in in vitro fertilization–embryo transfer and intracytoplasmic sperm injection programs. Fertil Steril 2004; 82: 1251 80. Liu DY, Baker HWG. Disordered acrosome reaction of spermatozoa bound to the zona pellucida: a newly discovered sperm defect causing infertility with

81.

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91.

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reduced sperm–zona penetration and reduced fertilization in vitro. Hum Reprod 1994; 9: 1694 Liu DY, Baker HW. Defective sperm–zona pellucida interaction: a major cause of failure of fertilization in clinical in-vitro fertilization. Hum Reprod 2000; 15: 702 Liu de Y, Baker HW. Frequency of defective sperm–zona pellucida interaction in severely teratozoospermic infertile men. Hum Reprod 2003; 18: 802 Liu de Y, Baker HW. High frequency of defective sperm–zona pellucida interaction in oligozoospermic infertile men. Hum Reprod 2004; 19: 228 Tesarik J. Appropriate timing of the acrosome reaction is a major requirement for the fertilizing spermatozoon Hum Reprod 1989; 4: 957 Cummins J, et al. A test of the human sperm acrosome reaction following ionophore challenge. J Androl 1991; 12: 98 Franken DR, Bastiaan HS, Oehninger S. Physiological induction of the acrosome reaction in human sperm: validation of a microassay using minimal volumes of solubilized, homologous zona pellucida. J Assist Reprod Genet 2000; 17: 374 van Duin M, Polman J, De Breet IT. Recombinant human zona pellucida protein ZP3 produced by Chinese hamster ovary cells induces the human sperm acrosome reaction and promotes sperm–egg fusion. Biol Reprod 1994; 51: 607 Dong KW, et al. Characterization of the biological activities of a recombinant human zona pellucida protein 3 (ZP3) expressed in human ovarian (PA-1) cells. Am J Obstet Gynecol 2001; 184: 835 Esterhuizen AD, et al. Clinical importance of a micro-assay for the evaluation of sperm acrosome reaction using homologous zona pellucida. Andrologia 2001; 33: 87 Esterhuizen AD, et al. Clinical importance of zona pellucida induced acrosome reaction (ZIAR test) in cases of failed human fertilization. Hum Reprod 2001; 16: 136 Bastiaan HS, et al. Zona pellucida induced acrosome reaction, sperm morphology, and sperm–zona pellucida binding assessments among subfertile men. J Assist Reprod Genet 2002; 19: 329 Bastiaan HS, et al. Relationship between zona pellucida-induced acrosome reaction, sperm morphology, sperm–zona pellucida binding, and in vitro fertilization. Fertil Steril 2003; 79: 49 Oehninger S. Clinical and laboratory management of male infertility: an opinion on its current status. J Androl 2000; 21: 814

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94. Oehninger S. Place of intracytoplasmic sperm injection in clinical management of male infertility. Lancet 2001; 357: 2068 95. Oehninger S, Gosden RG. Should ICSI be the treatment of choice for all cases of in-vitro conception?

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No, not in light of the scientific data. Hum Reprod 2002; 17: 2237 96. Monzó A, et al. Outcome of ICSI in azoospermic patients: stressing the liaison between the urologist and reproductive medicine. Urology 2001; 58: 69

15 Detection of DNA damage in sperm Ralf Henkel

INTRODUCTION

using defective spermatozoa in ICSI is much higher, which in turn increases the risk of transferring damaged DNA into oocytes. In addition, reports regarding increased chromosomal abnormalities, minor or major birth defects or childhood cancer suggest increased risks for babies born after ICSI15–20, and have led to serious concerns about this technique. For these grave reasons, various authors from different working groups have suggested that tests for DNA integrity and damage should be introduced into the routine andrological laboratory work-up21–25. Compared with other sperm parameters such as motility, Zini et al.26 regard the evaluation of sperm DNA fragmentation as a particularly reliable assay because of its low biological variability. In the past, a number of test systems have been developed to investigate sperm DNA damage at different levels and different sites. Among these, some highly sophisticated assays examine chromosomal aberrations, including multicolor fluorescence in situ hybridization (FISH), or assays that probe for structural integrity of sperm DNA such as the sperm chromatin structure assay (SCSA), using flow cytometry. Other test systems for sperm nuclear maturity and condensation such as the aniline blue stain are rather simple, and are based on the evaluation of stained sperm smears by a technologist.

It has been reported that sperm DNA damage is predictive of fertilization and pregnancy after natural conception1–3 and following the use of different techniques of assisted reproduction, namely intrauterine insemination (IUI)4, in vitro fertilization (IVF)5–8 and intracytoplasmic sperm injection (ICSI)9–12. This has important clinical implications for assisted reproduction techniques (ART), because the more invasive is the technique, the higher is the risk that a genetically damaged male genome will be transferred into the oocyte and fertilize the oocyte in vitro10,13. Normally, if the genetic damage in the male germ cell is severe, embryonic development stops at the time when the paternal genome is switched on, resulting in failed pregnancy10. However, genetic and biological protection mechanisms do not necessarily preclude further embryonic development, since Ahmadi and Ng14 have demonstrated that fertilization with damaged spermatozoa can result in live-born (mouse) pups. This study also showed that the injection of a cytosolic sperm factor into the oocyte is a key point in the activation of oocytes. Since the DNA fragmentation rate is significantly higher in patients with poor semen quality, and DNA damage cannot be recognized while selecting spermatozoa to be injected for ICSI, the probability of 225

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Since most of these assays are reportedly predictive of fertilization and pregnancy, this chapter contributes to an understanding of the currently available assays, so that the results of each can be better assessed. Moreover, in view of the varying financial capabilities of different andrology laboratories, this will then enable selection of which test systems can be employed to offer the most effective andrological diagnosis on the one hand, with optimum results following ART on the other. A summary of the test systems discussed in this chapter with a short description of the principle as well as main advantages and disadvantages is depicted in Table 15.1.

TEST SYSTEMS TO ASSESS DNA DAMAGE/FRAGMENTATION Sperm DNA damage can occur at different levels, i.e. (1) direct damage of DNA in the form of strand breakages due to apoptosis, oxidative stress or radiation, (2) during chromatin condensation and packaging, resulting in immature nuclear condensation, or (3) at the chromosomal level in the form of chromosomal aberrations or aneuploidies. As each of these levels is important for the transmission of male genetic information and chromatin condensation, they reflect sites where damage that can have serious effects on fertilization and pregnancy may occur. Therefore, methods to measure such damage have been developed. Some of these assays have been tested for their predictive value for male infertility within the scope of an ART program. Direct DNA damage can occur in the form of base modifications or single- and/or double-strand breakages. Base modifications are assessed through the measurement of 8-hydroxydeoxyguanosine, one of about 20 major biomarkers of oxidative DNA damage that has been shown to be representative, highly specific and potentially a mutagenic product. DNA strand breakages are often measured by means of the comet (single cell

gel electrophoresis) assay, TUNEL (terminal deoxynucleotide transferase-mediated dUTP nick-end labeling) assay, in situ nick translation, the metachromatic shift of acridine orange fluorescence in the acridine orange staining test or the sperm chromatin structure assay.

Measurement of 8-hydroxydeoxyguanosine 8-Hydroxydeoxyguanosine (8-OHdG) occurs as substantial oxidative modification of DNA and is present in abundance in DNA27 at levels of 2–4 per 100 000 deoxyguanosine molecules28,29 in human spermatozoa. It can be measured in genomic DNA by means of high-performance liquid chromatography (HPLC), with subsequent electrochemical or gas chromatography–mass spectrometry detection. In order to perform this test, DNA has to be extracted from human sperm, followed by enzymatic digestion and detection by means of HPLC analysis. Since the method depends on sufficient extraction of 8-OHdG, quality of DNA digestion and detection limit of the HPLC, relatively high numbers of spermatozoa are necessary30. Several studies have revealed significantly higher amounts of 8-OHdG in the spermatozoa of smokers than in non-smokers, which are due to the high oxidative DNA damage. On the other hand, the intake of antioxidants such as vitamin C and its concentration in seminal plasma provide protection again this oxidative damage29,31. High amounts of 8-OHdG have also been linked to male infertility32–34. If this DNA damage is not repaired, 8-OHdG may be mutagenic and may cause embryonic loss, malformations or childhood cancers3,28. Despite that this test shows clear clinical significance, i.e. it is highly specific, quantitative and correlated with sperm function30,35,36, the method is not commonly used because special equipment is required. Moreover, artifactual oxidation of deoxyguanosine can occur and lead to inaccurate results.

Base modifications

DNA fragmentation, single- and doublestrand breaks

DNA fragmentation, single- and doublestrand breaks Single-strand DNA breaks Differentiates between single- and doublestranded DNA and RNA

Susceptibility of nuclear DNA to denaturation

Measurement of 8hydroxydeoxy guanosine

Comet assay

TUNEL assay

In situ nick translation

Acridine orange test

Sperm chromatin structure assay (SCSA)

DNA damage

Assay principle

Assay

Type of assay

• Special equipment • Artifactual oxidation of deoxyguanosine • Large amount of sample

Clinically significant High specificity Quantitative Correlates with sperm function Simple to perform and cheap Correlates with TUNEL assay High sensitivity Observation of individual cells Small number of cells required Correlates with fertility

• • • • • • • • • •

Clinically significant High sensitivity and specificity Large number of sperm counted by flow cytometry Unbiased quantitative assessment of DNA-bound acridine orange • Correlates with sperm function and fertility

• • • •

• Special equipment • More expensive

Continued

• Distinction between differently labeled sperm not always easy • Special equipment

• Not specific to oxidative damage • Special equipment

• Correlates with TUNEL assay • Specific for single-strand DNA breaks • Specific to endogenous DNA breaks • Easy to perform • Low cost • Correlates with sperm function and fertility

• Not specific to oxidative damage • More costly • Special equipment

• Clinically significant • High sensitivity and specificity • Correlates with sperm function and fertility

• Fluorescence microscopy • Experienced observer • Not specific to oxidative damage

Main disadvantages

Main advantages

Table 15.1 Principles and main advantages and disadvantages of the most important test systems used to examine sperm DNA damage and chromosomal aberrations

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Competing for same DNA binding site with protamines Detection of chromosomal abnormalities

chromomycin A3 stain

Fluorescence in situ hybridization (FISH)

Clinically significant High specificity and sensitivity Low cost Correlates with fertility in IVF Easy to perform

• Clinically significant • High specificity and sensitivity • Correlates with fertility, pregnancy and disease of offspring

• Clinically significant • High specificity and sensitivity • Correlates with fertility in IVF and ICSI

• Correlates with acridine orange stain, TUNEL assay and aniline blue stain • Easy to perform

• • • • •

Main advantages

• Labor-intensive • Expensive • Special equipment

• Possible inconsistencies due to subjective appraisal

• Clinical relevance not yet proven • Inconsistencies due to subjective appraisal

• Correlation with other sperm parameters controversial • Inconsistencies due to subjective appraisal

Main disadvantages

TUNEL, terminal deoxynucleotide transferase-mediated dUTP nick-end labeling; IVF, in vitro fertilization; ICSI, intracytoplasmic sperm injection

Chromosomal aberration and aneuploidy

Binding to damaged dense chromatin

Staining of lysine residues of remaining histones

Assay principle

Toluidine blue stain

DNA condensation/ Aniline blue stain packaging

Assay

Continued

Type of assay

Table 15.1

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DETECTION OF DNA DAMAGE IN SPERM

Comet assay The single cell (micro)gel electrophoresis or ‘comet’ assay was developed to evaluate DNA integrity, including single- and double-strand breaks in somatic cells37. In 1988, Singh et al.38 used alkaline conditions at pH > 13, a modification of the assay, which enables the detection of DNA single-strand breaks, alkali-labile sites, DNA–DNA/DNA–protein crosslinking and single-strand breaks associated with incomplete excision repair sites, and increased the sensitivity of the test. Since that time, the range of applications and the number of users have increased. In this assay, DNA strand breaks migrate in an agarose gel, and, depending on the amount of damaged DNA, create a bigger or smaller tail, which is visualized by means of DNA-specific fluorescent dyes, while intact, supercoiled, compact DNA remains in the nucleus39. The shape resembles a comet, and hence the name of the assay. For evaluation of the comet assay, the length of the tail, the percentage of DNA in the tail (intensity of tail staining) or the product of these two parameters, the tail moment, are taken into consideration. With regard to male infertility and sperm function, several groups40–43 have shown the clinical relevance of the comet assay. Although its predictive value has also been documented for fertilization and embryo development in both IVF and ICSI7,44,45, the origin of such DNA damage remains obscure, and sources including apoptosis, improper DNA packaging and ligation during spermatogenesis or oxidative stress have been discussed. For the last, two different sources, reactive oxygen species (ROS) produced by leukocytes or by the spermatozoa themselves, seem possible (for review see reference 46). Interestingly, ROS that are normally found in the male reproductive tract can induce this DNA damage47. Moreover, DNA damage was reportedly increased after exposure to toxins (including cigarette-smoking), chemotherapy or radiation48–51. Unfortunately, to date, there is no standardized protocol to perform and evaluate this test, and it

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is therefore difficult to compare the results from different groups. While some authors calculate the percentage of comet-forming sperm, others report the average extent of the tails in a given sperm population40,52,53. On the other hand, the assay is easy to perform, is one of the most sensitive techniques available to measure DNA strand breaks39 and correlates very well with results of the TUNEL assay54.

TUNEL assay Another test specific for broken sperm DNA is the TUNEL (terminal deoxynucleotide transferasemediated dUTP nick-end labeling) assay55. The principle is based on the addition of labeled DNA precursors (dUTP: deoxyuridine triphosphate) at single- and double-strand DNA breaks by means of an enzymatically catalyzed reaction, using the template-independent terminal deoxynucleotide transferase (TdT). It incorporates biotinylated or fluorescinated dUTP to the 3′-OH ends of the DNA, which increase with the number of strand breaks. Compared with other methods to detect DNA damage, the TUNEL assay is more sophisticated, more expensive and more time-consuming. However, good-quality control parameters such as low intraobserver and interobserver variability have been demonstrated56. In addition, flow-cytometric measurement of the sperm sample analyzing a large amount of cells is possible. Due to its high specificity and reproducibility, the TUNEL assay is one of the most frequently used test systems to investigate sperm DNA fragmentation. Its relevance in respect of sperm function57–59 as well as fertilization and pregnancy has been proved repeatedly4,5,8,60. Sperm DNA fragmentation provides a clinical explanation even for early embryonic death61 and recurrent pregnancy loss62. Moreover, Shoukir et al.63 found a significantly lower blastocyst formation rate after ICSI compared with IVF, and postulated a negative paternal effect on preimplantation embryo development. The TUNEL assay evaluates DNA fragmentation, which is a rather late stage of

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apoptosis, and it cannot actually distinguish between apoptotic and necrotic cells64,65. This is even more important, as Sakkas et al.66 found that TUNEL positivity and apoptotic markers such as the asymmetric distribution of phosphatidylserine in the sperm plasma membrane do not always exist in unison.

In situ nick translation In contrast to the TUNEL assay, which detects both single- and double-strand DNA breaks, in situ nick translation detects only single-strand DNA breaks. This test quantifies the incorporation of labeled (biotinylated or fluorescinated) dUTP at the 3′-OH recessed termini of singlestranded DNA in a template-dependent enzymatic reaction by means of DNA polymerase I. Labeling with the in situ nick translation is indicative of endogenous nicks in the DNA67,68. Data obtained in human spermatozoa with both techniques, in situ nick translation and the TUNEL assay, are highly correlated69. In somatic cells, necrotic nuclei seem to be preferably stained by in situ nick translation, while the TUNEL assay appears to be rather indicative of apoptosis70. However, since spermatozoa do not show the typical morphological alterations characteristic of apoptosis in somatic cells, additional specific tests for other markers of apoptosis such as phosphatidylserine externalization, Fas expression or the presence of other active proapoptotic factors should be performed in order to distinguish clearly between apoptosis and necrosis.

Acridine orange test The acridine orange test is a slide-based version of the original human sperm chromatin heterogeneity test71 that was developed by Tejada et al.72. This test measures the susceptibility of sperm nuclear DNA to acid-induced denaturation by means of the metachromatic properties of acridine orange. This dye intercalates into the DNA as a monomer,

which fluoresces green with double-stranded DNA, and binds to single-stranded DNA or RNA as an aggregate that emits red-orange light after excitation73. Due to its simplicity, several working groups have correlated the acridine orange test with different sperm functional parameters, including normal sperm morphology, as well as with male fertility in assisted reproduction programs. While Ibrahim and Pedersen74 could not find a significant correlation between the acridine orange test and sperm motility and the penetration of zonafree hamster oocytes in the sperm penetration assay, others have demonstrated significant correlations with motility75, sperm count72, sperm– zona pellucida binding76 and fertilization in an assisted reproduction program for IVF and ICSI77–79. Additionally, a significant correlation between chromatin integrity and normal sperm morphology as one of the most predictive sperm parameters for fertilization in vitro has been shown repeatedly75,77. Despite these mainly positive reports regarding the clinical value of the acridine orange test, concern has arisen about its reliability. This is mainly based on: (1) the poor conditions for the metachromatic shift from green to red-orange as the dye adsorbs on the glass surface, and (2) the difficulty in distinguishing between normal, green, and abnormal, red-orange, sperm heads accurately, especially if a sperm head contains both single- and double-stranded DNA. Furthermore, rapid fading of the fluorescence80 and heterogeneous slide staining81 are additional problems when performing this test. Thus, Evenson et al.53,71 developed the more reliable sperm chromatin structure assay (SCSA).

Sperm chromatin structure assay The SCSA is based on the same principle of metachromatic shift of the color of acridine orange as in the acridine orange test. However, in contrast to the acridine orange test, the detection method in the SCSA is flow cytometry. This

DETECTION OF DNA DAMAGE IN SPERM

approach makes it possible to measure large amounts of spermatozoa (typically 5000–10 000) per sample, which in turn renders the technique easy and highly reproducible82. Moreover, the inter- and intra-assay variability as well as the technical problems described for the acridine orange test are overcome by this automated reading. The interassay variability of the flowcytometric detection of sperm chromatin damage has been shown to be less than 5%83. In addition to the advantages described thus far and summarized in Table 15.1, the flexibility of this assay needs to be mentioned. The test can be performed on fresh and frozen samples, which makes it easier to collect the specimen or even to ship them for evaluation22. A number of clinical studies have revealed the SCSA to be reliable and predictive for assessing the male fertility status. A percentage of chromatin-disturbed spermatozoa (red-orange stained sperm) higher than 30% is indicative of male infertility and poor fertilization in IUI, IVF and ICSI, including ongoing pregnancy81,82,84–87. Considering that sperm DNA integrity as measured by means of the SCSA is a more constant parameter over a longer period of time, compared with other sperm parameters83, this assay has also been found suitable for effective use in epidemiological studies88.

TEST SYSTEMS TO ASSESS SPERM DNA CONDENSATION/PACKAGING Apart from test systems that directly assess the quality and integrity of the DNA itself, assays have been developed that probe DNA packaging and maturity. This is of particular importance because in spermatozoa, the histones, which are the predominant nuclear proteins in any somatic cell, are replaced during spermiogenesis by protamines in a multistep process. These protamines are disulfide bridge-stabilized, highly basic proteins that fit into the minor grooves of the DNA, neutralize the negative charges of the phosphate groups and thus

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enable the DNA to form linear arrays fitting into the major groove of the neighboring strand, instead of the voluminous supercoiled ‘solenoids’ present in somatic cells. This results in a highly condensed sperm nucleus in which the DNA takes up about 90% of the total volume. In contrast, the nuclear volume of the DNA in mitotic chromosomes is about 15%, and in somatic cells about 5%89. In the case of disturbed chromatin condensation, histones persist in the sperm nucleus and cause decondensation problems in the male genome after the spermatozoon enters the oocyte. Thus, patients showing abnormalities of this essential sperm maturation process during spermiogenesis are subfertile or infertile90–92. Various methods based on different principles for evaluation of the maturity grade of sperm chromatin condensation are available, and are discussed below.

Aniline blue stain Immature, poorly chromatin-condensed sperm nuclei still contain the lysine-rich histones. In an acid–base reaction, acidic aniline blue binds to the basic lysine residues and thus discriminates between lysine-rich histones and arginine/ cysteine-rich protamines. This test provides a positive blue staining of spermatozoa with disturbed chromatin condensation, while mature spermatozoa that contain protamines will not be stained. Terquem and Dadoune93 originally described this simple and inexpensive slide-based test. However, owing to this feature, and the fact that the test is visually scored by a technologist, inconsistencies due to subjective assessment might arise, which in turn can compromise its repeatability. On the other hand, Franken et al.94 have shown a coefficient of intra-assay variability for the aniline blue stain of less than 10%, indicating that it is a repeatable technique. According to studies by Dadoune et al.95 and Auger et al.96, a normal ejaculate should contain at least 75% aniline blue-negative spermatozoa,

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which indicates normal chromatin condensation. These data were confirmed by Haidl and Schill97 and Hammadeh90, who showed that normal chromatin condensation is mandatory to induce fertilization. With regard to IVF and pregnancy, different groups97–100 have demonstrated the clinical significance of this simple test, and the supplementation of routine semen analysis with this assay during andrological work-up has been suggested100. However, the question of whether the quality of sperm chromatin condensation contributes to poor fertilization and pregnancy rates after ICSI remains debated. While studies by Van Ranst et al.101 and Hammadeh et al.102 employing the aniline blue stain failed to predict the outcome of fertilization by ICSI, Sakkas et al.103 showed, when applying the chromomycin A3 (CMA3) stain, a significantly higher percentage of spermatozoa with poorly packed chromatin in the ejaculate and only about half the fertilization rate in ICSI patients, compared with IVF patients. ICSI embryos even had a significantly lower developmental potential to reach the blastocyst stage. In a comparative study using the aniline blue and the CMA3 stain, Razavi et al.104 confirmed this result, as only the detection of sperm protamine deficiency by means of CMA3 showed a significant effect on ICSI outcome. Thus, it appears that poor sperm chromatin condensation may contribute to the failure of fertilization after ICSI.

Toluidine blue stain The toluidine blue stain is another slide-based simple and inexpensive test method to evaluate sperm DNA structure and packaging105,106, which is based on the metachromatic and orthochromatic staining abilities for chromatin. Toluidine blue is basic thiazine nuclear dye that is intensively incorporated into damaged dense chromatin. Like acridine orange, after acid treatment of somatic apoptotic cells, this dye shows a metachromatic shift of color from light blue in normal sperm heads to purple-violet in nuclei with fragmented DNA106. To differentiate spermatozoa for DNA

integrity, Erenpreisa et al.107 introduced this method. The same authors demonstrated a high correlation (r = 0.63–0.70; p < 0.01) between the toluidine blue stain, the acridine orange stain and the aniline blue stain, and concluded that the technique is sensitive enough to estimate in situ sperm DNA integrity. In addition, a significant correlation between the purple-violet staining pattern and the TUNEL assay could be revealed108. In an earlier study by Barrera et al.109, it was found that sperm from fertile donors showed mostly the orthochromatic pale-blue staining pattern, whereas in oligozoospermic patients a high percentage of spermatozoa revealed the metachromatic purple-violet staining. Unfortunately, further direct clinical significance has not yet been proved. Therefore, one can rely only on the high correlation with acridine orange and on the experience with that test.

Chromomycin A3 stain Chromomycin A3 (CMA3) is a guanine–cytosinespecific fluorochrome that competes directly with protamines for the same binding site in the DNA. Like the aniline blue stain, the CMA3 stain is a slide-based method that identifies poorly condensed DNA. Strongly stained sperm heads apparently lack protamines, whereas spermatozoa not stained by CMA3 show normal chromatin condensation. Thus, the stain is indicative of an underprotamination of spermatozoa67,68. This was confirmed by the observation by Bizzaro et al.110 that CMA3 positivity of murine and human spermatozoa decreases after in situ protamination with salmon protamines. Moreover, these authors showed that the addition of increasing amounts of salmon protamines induced distinct morphological changes, so that initially deprotaminated sperm heads, which were decondensed, regained their original condensed appearance after the treatment. With regard to other sperm parameters, the CMA3 stain has been significantly and positively correlated with normal sperm morphology and

DETECTION OF DNA DAMAGE IN SPERM

negatively correlated with sperm count but not with sperm motility94,111,112. Manicardi et al.68 revealed a significant association of CMA3 positivity with the presence of endogenous nicks in sperm DNA, which in turn is an indication of disturbed spermiogenesis in specific patients, as these nicks normally occur during late spermiogenesis and disappear once sperm chromatin packaging is completed113,114. Since the test is also highly predictive of fertilization after IVF as well as after ICSI104,112,115–117, and has been shown to be superior in predicting the outcome of ART as compared with aniline blue staining and the acridine orange test117, it is suggested that determination of sperm chromatin condensation should be performed in a sequential andrological diagnosis program prior to any kind of assisted reproduction. Reportedly, the calculated cut-off value for the prediction of fertilization is 30%117, i.e. at least 70% of the spermatozoa should be CMA3negative.

TEST SYSTEMS TO ASSESS CHROMOSOMAL ABERRATION AND ANEUPLOIDY Apart from direct damage to sperm DNA resulting in strand breakages and the abnormal packaging of the male genome, chromosomal aberrations including aneuploidy or structural chromosome reorganizations have been identified as a cause of male infertility. Previous research has revealed that elevated genetic damage in spermatozoa is significantly increased in infertile men118,119, and that aneuploidy is significantly higher in patients with recurrent pregnancy losses120. This is of particular importance, since it has been shown that chromosomal aneuploidy and diploidy in spermatozoa are negatively correlated with sperm count in the ejaculate and progressive motility121,122, and concern about miscarriages and chromosomal abnormalities in the offspring has been raised, particularly

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for ICSI43,123,124. The most frequently occurring aneuploidy syndromes are: triple X, Klinefelter’s (XXY), Turner’s (X instead of XX or XY), XYY, Patau’s (trisomy of chromosome 13), Edward’s (trisomy of chromosome 18) or Down’s (trisomy of chromosome 21). A very rare disorder is the Jacobsen syndrome, in which a terminal deletion of chromosome 11q occurs. Others are the ‘Cri du chat’ syndrome, which is caused by deletion of part of the short arm of chromosome 5, or the Wolf-Hirschhorn syndrome, which is caused by partial deletion of the short arm of chromosome 4. The method of choice to investigate these chromosomal aberrations is fluorescence in situ hybridization (FISH).

Fluorescence in situ hybridization The principle of FISH is the use of fluorochromelabeled chromosome-specific probes that recognize a large section of the chromosome (0.2–2.0 Mb). These probes are hybridized with a sample of spermatozoa, and the labeled part of the chromosome appears as a fluorescent domain within the nucleus, where it can be identified by means of fluorescence microscopy125. Meanwhile, probes for all human and many rodent chromosomes are available, and can be used to identify such chromosomal aberrations by applying socalled multicolor FISH, whereby usually three or four differently fluorescing probes are hybridized in parallel. This is because the scoring has to be performed visually, and the eye is limited in distinguishing different fluorescing colors. Although multicolor FISH has been shown to be highly specific with little or no error, and the specimen can be frozen even without cryoprotection until examination, the technique is currently highly labor-intensive and expensive. In this regard, Baumgartner et al.126 recently developed a laser-scanning cytometry method for automated sperm analysis of the X chromosome, but the technique is still expensive and requires highly skilled personnel.

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CONCLUSIONS Generally, DNA damage can occur at different levels, i.e. direct breakage of the DNA, abnormal chromosome packaging and chromosomal aberrations. DNA damage has been proved to be of importance for human fertility as well as for the health of the offspring. Several techniques have been developed to examine such damage. Based on this knowledge, an andrological investigation should not only consist of routine spermiogram analysis, which includes sperm count, motility and morphology, but also incorporate more sophisticated testing, e.g. for DNA damage, as there is compelling evidence for its importance and clinical relevance. The practical question arising at this point is which test should be applied. This certainly depends on the personnel and financial capabilities of an ART program or andrology unit. Other questions arise concerning the standardization of such tests. The latter is an important issue because this is closely connected with the predictive value of the test in question. Recently, more research has been performed, and understanding of the influence of the paternal genome on the reproductive process and methodology for examining such DNA damage have improved considerably. To date, various methods of testing sperm DNA integrity have been investigated with regard to their clinical value. Even though some of them are rather expensive and others are less reproducible, nowadays more information about male fertility status can and should be obtained, following which better strategies can be pursued to improve counseling and treatment of patients.

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3. Loft S, et al. Oxidative DNA damage in human sperm influences time to pregnancy. Hum Reprod 2003; 18: 1265 4. Duran EH, et al. Sperm DNA quality predicts intrauterine insemination outcome: a prospective cohort study. Hum Reprod 2002; 17: 3122 5. Sun JG, Jurisicova A, Casper RF. Detection of deoxyribonucleic acid fragmentation in human sperm: correlation with fertilization in vitro. Biol Reprod 1997; 56: 602 6. Host E, Lindenberg S, Smidt-Jensen S. DNA strand breaks in human spermatozoa: correlation with fertilization in vitro in oligozoospermic men and in men with unexplained infertility. Acta Obstet Gynecol Scand 2000; 79: 189 7. Morris ID, et al. The spectrum of DNA damage in human sperm assessed by single cell gel electrophoresis (Comet assay) and its relationship to fertilization and embryo development. Hum Reprod 2002; 17: 990 8. Henkel R, et al. Influence of deoxyribonucleic acid damage on fertilization and pregnancy. Fertil Steril 2004; 81: 965 9. Lopes S, et al. Sperm deoxyribonucleic acid fragmentation is increased in poor-quality semen samples and correlates with failed fertilization in intracytoplasmic sperm injection. Fertil Steril 1998; 69: 528 10. Henkel R, et al. DNA fragmentation of spermatozoa and ART. Reprod Biomed Online 2003; 7: 477 11. Lewis SEM, et al. An algorithm to predict pregnancy in assisted reproduction. Hum Reprod 2004; 19: 1385 12. Tesarik J, Greco E, Mendoza C. Late, but not early, paternal effect on human embryo development is related to sperm DNA fragmentation. Hum Reprod 2004; 19: 611 13. Twigg JP, Irvine DS, Aitken RJ. Oxidative damage of DNA in human spermatozoa does not preclude pronucleus formation at intracytoplasmic sperm injection. Hum Reprod 1998; 13: 1864 14. Ahmadi A, Ng SC. Developmental capacity of damaged spermatozoa. Hum Reprod 1999; 14: 2279 15. In’t Veld P, et al. Sex chromosomal abnormalities and intracytoplasmic sperm injection. Lancet 1995; 346: 773 16. Kurinczuk JJ, Bower C. Birth defects in infants conceived by intracytoplasmic sperm injection: an alternative interpretation. Br Med J 1997; 315: 1260 17. Ji BT, et al. Paternal cigarette smoking and the risk of childhood cancer among offspring of nonsmoking mothers. J Natl Cancer Inst 1997; 89: 238

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63. Shoukir Y, et al. Blastocyst development from supernumerary embryos after intracytoplasmic sperm injection: a paternal influence? Hum Reprod 1998; 13: 1632 64. Frankfurt OS, et al. Monoclonal antibody to singlestranded DNA is a specific and sensitive cellular marker of apoptosis. Exp Cell Res 1996; 226: 387 65. Didenko VV, Hornsby PJ. Presence of doublestrand breaks with single-base 3′ overhangs in cells undergoing apoptosis but not necrosis. J Cell Biol 1996; 135: 1369 66. Sakkas D, et al. Nature of DNA damage in ejaculated human spermatozoa and the possible involvement of apoptosis. Biol Reprod 2002; 66: 1061 67. Bianchi PG, et al. Effect of deoxyribonucleic acid protamination on fluorochrome staining and in situ nick-translation of murine and human mature spermatozoa. Biol Reprod 1993; 49: 1083 68. Manicardi GC, et al. Presence of endogenous nicks in DNA of ejaculated human spermatozoa and its relationship to chromomycin A3 accessibility. Biol Reprod 1995; 52: 864 69. Manicardi GC, et al. DNA strand breaks in ejaculated human spermatozoa: comparison of susceptibility to the nick translation and terminal transferase assays. Histochem J 1998; 30: 33 70. Gorczyca W, Gong J, Darzynkiewicz Z. Detection of DNA strand breaks in individual apoptotic cells by the in situ terminal deoxynucleotidyl transferase and nick translation assays. Cancer Res 1993; 53: 1945 71. Evenson DP, Darzynkiewicz Z, Melamed MR. Relation of mammalian sperm chromatin heterogeneity to fertility. Science 1980; 210: 1131 72. Tejada RI, et al. A test for the practical evaluation of male fertility by acridine orange (AO) fluorescence. Fertil Steril 1984; 42: 87 73. Rigler R. Microfluorometric characterization of intracellular nucleic acids and nucleoproteins by acridine orange. Acta Physiol Scand 1966; 67 (Suppl 1): 122 74. Ibrahim ME, Pedersen H. Acridine orange fluorescence as male fertility test. Arch Androl 1988; 20: 125 75. Shibahara H, et al. Clinical significance of the acridine orange test performed as a routine examination: comparison with the CASA estimates and strict criteria. Int J Androl 2003; 26: 236 76. Liu DY, Baker HWG. Sperm nuclear chromatin normality: relationship with sperm morphology, sperm–zona pellucida binding, and fertilization rates. Fertil Steril 1992; 58: 1178

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16 Chromosomal and genetic abnormalities in male infertility Pasquale Patrizio, Jose Sepúlveda, Sepideh Mehri

BACKGROUND

disorder can impair hormonal production or the stimulation of spermatogenesis (pretesticular event), or can impact upon control of the spermatogenic process itself (testicular event). In animal models and to some extent also in humans, genetic abnormalities affecting signaling cascades involved in the meiotic control of spermatogenesis are continuously being discovered and reported5,6. Other genetic/chromosomal disorders (for example cystic fibrosis and adult polycystic kidney disease) can affect sperm transport (posttesticular event). In this chapter we utilize the following scheme of classification: (1) male infertility with a gene defect and (2) male infertility with chromosomal aberrations (either numerical or structural)7.

About 15% of couples of reproductive age are affected by infertility, and in some 50% the male is the sole or main contributor1. The identification and initial classification of male infertility still rely on the results of semen analysis (i.e. azoospermia, oligozoospermia, asthenozoospermia, teratozoospermia or a combination), but this method alone is insufficient to determine a specific etiology of the disorder. A complete work-up, including detailed history and physical examination, hormonal and immunological assays, ultrasound or Doppler studies and genetic and chromosome testing is essential2. Recent advances in molecular genetics have greatly improved our understanding of many unexplained forms; however, 50% of cases still remain unclassified3. The advent of assisted reproductive techniques, namely intracytoplasmic sperm injection (ICSI), has provided the opportunity for severely infertile men to father their own offspring, but if genetic or chromosomal defects are responsible for infertility, then there is concern about transmitting genetic defects to the next generation4. There are different approaches to classifying male infertility on a genetic basis. In some textbooks the different forms are divided into pretesticular, testicular and post-testicular forms. A genetic or a chromosomal numerical or structural

MALE INFERTILITY WITH A GENE DEFECT These disorders are caused by a mutation at a single-gene locus, and either can occur de novo or are inherited as autosomal (dominant or recessive) or X-linked. It is estimated that over 10 000 human diseases are monogenic. The global prevalence of all single-gene diseases at birth is approximately 10/10008. Mendelian disorders observed in infertile men are detailed in Table 16.1. This list is by no means complete, but includes those 239

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Table 16.1

Gene defects and male infertility

Condition

Gene involved (mapping)

Incidence

Phenotype

Inherited

Hemochromatosis

HFE (6p21.3) HFE (1q21)-juvenile

1 : 500

Organ failure (liver and testis) by iron overload

Autosomal recessive

Autosomal dominant polycystic kidney disease

PKD1 (16p13.3) PKD2 (4q21–23) PKD3 (?)

1 : 1000

Multiple cysts (kidney, liver, spleen, pancreas, testis, epididymis, seminal vesicle)

Autosomal dominant

Cystic fibrosis

CFTR (7q31.2)

1 : 2500

Respiratory infections, Wolffian duct anomaly, pancreatic insufficiency

Autosomal recessive

Congenital adrenal hyperplasia

P450C21 (6p21.3) 21-hydroxylase deficiency (most common)

1 : 5000

Variable, elevated ACTH, inhibited FSH/LH secretion, azoospermia

Autosomal recessive

Myotonic dystrophy

DMPK (19q13.2–3)

1 : 8000

Muscle wasting, cataracts; atrophic testes

Autosomal dominant

Usher’s syndrome

USH1 (14q32) USH2 (1q41) USH3 (3q21–q25)

1 : 17 000

Low sperm motility, hearing loss, retinitis pigmentosa

Autosomal recessive

Prader–Willi syndrome

SNRPN (15q11q13)

1 : 20 000

Obesity, muscular hypotonia, mental retardation, hypogonadotropic hypogonadism

Autosomal dominant

Sex reversal syndrome

SRY (Yp11.3)

1 : 25 000

46,XX SRY(+) 46,XY SRY(–)

Y-linked

Kallman’s syndrome

KAL1 (Xp22.3)1 KAL2 (8p12)2 KAL3 (?)3

1 : 30 000

Hypogonadotropic hypogonadism, anosmia

1 X-linked recessive 2 Autosomal dominant 3 Autosomal recessive

Immotile cilia syndrome

DNAI1 (9p21–p13) DNAH5 (5p) 19q13.2, 16p2, 15q13

1 : 35 000

Sinusitis, bronchiectasis, immotile sperm

Autosomal recessive

Cerebellar ataxia

CLA1 (9q34–9) CLA3 (20q11–q13)

1 : 50 000

Eunuchoid phenotype, cerebellar impairment, atrophic testes

Autosomal recessive

Sickle cell anemia

HBB (11p15.5) (mutation)

1 : 58 000

RBC sickle shape, testicular microinfarctions

Autosomal recessive

Androgen insensitivity syndrome

AR (Xq11–q12)

1 : 60 000

Partial/complete testicular feminization

X-linked recessive

β-Thalassemia

HBB (11p15) (deletion)

1 : 114 000

Anemia; iron overload (pituitary and testis)

Autosomal recessive

Bardet–Biedl syndrome

BBS (11q13, 16q21, 3p12–q13, 15q22.3, 2q31, 20p12, 4q27, 14q32.11)

1 : 160 000

Retinal degeneration, obesity, cognitive impairment, GU malformations, polydactyly, hypogonadism

Autosomal recessive

Continued

CHROMOSOMAL AND GENETIC ABNORMALITIES IN MALE INFERTILITY

Table 16.1

241

Continued

Condition

Gene involved (mapping)

Incidence

Phenotype

Inherited

Mixed gonadal dysgenesis??

WT1 (11p13) DAX1 (Xp21.3) testatin (20p11.2)

Rare

Unilateral testis (most common with SCO) and contralateral streak gonad, ambiguous external genitalia

Autosomal dominant Xlinked recessive cytogenetic

Persistent Müllerian duct syndrome

AMH (19p13.3–p13.2) AMHR (12q13)

< 200 cases reported

Incomplete involution of Müllerian structures

Autosomal? X-linked

LH/FSH hormone and receptor mutations

LHβ (19q13.32) FSHβ (11p13)

Few male cases reported

Delayed puberty, arrested spermatogenesis

Autosomal recessive?

5α-Reductase deficiency

SRD5A1 (5p15) SRD5A2 (2p23)

Unknown

Male pseudohermaphroditism, severe hypospadias

Autosomal recessive

LH, luteinizing hormone; FSH, follicle stimulating hormone; ACTH, adrenocorticotropic hormone; RBC, red blood cell; GU, genitourinary; SCO, Sertoli cell-only syndrome

genetic conditions with the potential for clinical relevance.

Kallman’s syndrome Kallman’s syndrome (KS) consists of congenital hypogonadotropic hypogonadism and anosmia. The gene responsible for the X-linked form of KS, KAL, encodes a protein, anosmin-1, that plays a key role in the migration of GnRH neurons and olfactory nerves to the hypothalamus. As a consequence of failed neuronal migration, the hypothalamus and anterior pituitary are unable to stimulate the testis. The hallmark of KS is delayed puberty and atrophic testes (< 2 cm). Clinical manifestations depend on the degree of hypogonadism, and in some cases the syndrome may present only with subfertility. Testicular biopsies display a wide range of findings from germ-cell aplasia to focal areas of complete spermatogenesis. In addition to X-linked pedigrees, autosomal dominant and recessive kindred with KS have also been reported9.

Autosomal dominant KAL2 in 8p12 (FGFR1, fibroblast growth factor receptor-1) and autosomal recessive KAL3 are associated with nonreproductive features, including cleft palate, mirror movements and dental agenesis10. Recent studies have confirmed that mutations in the coding sequence of the KAL1 gene occur in the minority of KS cases, while the majority of familial (and presumably sporadic) cases are caused by defects in at least two autosomal genes11.

Congenital adrenal hyperplasia Congenital adrenal hyperplasia (CAH) results from inherited defects in one of the five enzymatic steps required for the biosynthesis of cortisol from cholesterol. The most common form of CAH (95%) involves a deficiency of 21-hydroxylase located on 6p21.312. Mutations in the cytochrome P450 21hydroxylase gene (CPY21) tend to be transmitted in an autosomal recessive pattern. Deficiency of

242

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21-hydroxylase occurs in three forms: (1) simple virilizing, (2) salt-wasting and (3) non-classical. The simple virilizing and salt-wasting forms of 21-hydroxylase deficiencies are characterized by excess adrenal androgen biosynthesis in utero. This disorder in males is not recognized at birth; they have normal genitalia and are not diagnosed until later, often with a salt-wasting crisis. Cortisol and aldosterone production is low, but testosterone production is normal (peripheral conversion of androstenedione). Elevated adrenal androgen secretion (due to elevated adrenocorticotropic hormone, ACTH) in male CAH patients may suppress both follicle stimulating hormone (FSH) and luteinizing hormone (LH) secretion with resultant small testes, decreased spermatogenesis and testicular androgen production13,14.

Prader–Willi syndrome Prader–Willi syndrome (PWS) was the first human disorder attributed to genomic imprinting, whereby genes are expressed differentially based upon the parent of origin. PWS results from the loss of imprinted gene SNRPN on the paternal 15q11.2–13 locus with an autosomal dominant pattern. The loss of maternal genomic material at the same locus results in another imprinted disorder (Angelman’s syndrome)15. Characteristics of this disorder include neonatal hypotonia, childhood-onset hyperphagia, obesity, mental retardation and short stature. A deficiency of GnRH is the postulated reason for the hypogonadism16.

Bardet–Biedl syndrome Bardet–Biedl syndrome (BBS) is a genetically heterogeneous disorder with linkage to eight loci17,18 (Table 16.2). Although BBS was originally thought to be a recessive disorder19, controversy exists about the presence of a recessive pattern ‘with variable penetrance’20. Cardinal features include obesity, retinitis pigmentosa, polydactyly, hypogonadotropic hypogonadism, renal cystic dysplasia and developmental

Table 16.2 Chromosome localization of genes involved in Bardet–Biedl syndrome (BBS) Gene involved

Mapping

BBS 1

11q13

BBS 2

16q21

BBS 3

3p12–q13

BBS 4

15q22.3

BBS 5

2q31

BBS 6

20p12

BBS 7

4q27

BBS 8

14q32.11

delay. Other associated clinical findings in BBS patients include diabetes, hypertension and congenital heart defects. The clinical diagnosis is based on the presence of at least four of these symptoms21. Some of the BBS genes are also involved in the function of the cilia and the formation of flagella, which can impair sperm motility and cause infertility22.

Hemochromatosis Hereditary hemochromatosis (HH) is an autosomal disorder characterized by excessive absorption of dietary iron, which may result in parenchymal iron overload and subsequent tissue damage23. Hypogonadotropic hypogonadism is the most frequent endocrinopathy associated with HH, secondary to iron deposition in the pituitary gonadotrophs, leading to loss of libido, impotence and body hair loss24. There are four types of HH, summarized in Table 16.325. Type 1 is the most common; the other types of HH are considered to be rare and have been studied in only a small number of families26.

Cerebellar ataxia and hypogonadism Cerebellar ataxia and hypogonadism is a rare autosomal recessive condition most commonly

CHROMOSOMAL AND GENETIC ABNORMALITIES IN MALE INFERTILITY

Table 16.3

243

Classification of hereditary hemochromatosis

Hereditary hemochromatosis

Locus

Inherited

Onset

Type 1 (classical)

6p21

Autosomal recessive

> 30 years

Type 2 (juvenile)

1q21

Autosomal recessive

< 30 years

Type 3

7q22

Autosomal recessive

4th–5th decade of life

Type 4

2q32

Autosomal dominant

> 60 years

observed in consanguineous unions, with onset at 20 years old. Clinical features include cerebellar impairment (speech and gait abnormalities), and eunuchoid phenotype with atrophic testis and low libido. Infertility is secondary to hypothalamic– pituitary dysfunction, possibly because of brain atrophy or hypoplasia. Genes involved are CLA1 (9q34–9) for the most common adult-onset type27, and CLA3 (20q11–q13) for infant onset28.

Other idiopathic hypogonadotropic hypogonadism Some other forms of hypogonadotropic hypogonadism previously classified as idiopathic (IHH) have recently been associated with genetic mutations. They include the DAX1 gene, which encodes a nuclear transcription factor, leading to X-linked IHH associated with congenital adrenal hypoplasia (CAH)11. Another mutation in the prohormone convertase gene (PC1) has been linked to hypogonadotropic hypogonadism, in addition to extreme obesity, hypocortisolemia and deficient conversion of proinsulin to insulin29. Homozygous mutations in GPR54, a gene encoding G-protein-coupled receptor-54, have lately been reported as another cause of hypogonadotropic hypogonadism30.

Immotile cilia syndrome The immotile cilia syndrome (ICS) is a group of heterogeneous diseases with impaired or absent ciliary motility, and the most common is

Kartagener’s syndrome. Abnormalities in the motor apparatus or axoneme, due to either missing or very short dynein arms, cause a deficit in sperm motility. Clinical manifestations include chronic cough, sinus infection, nasal polyposis, bronchiectasis and infertility with asthenozoospermia31. While infertility is universal in patients with ICS, there is another condition known as fibrous sheath dysplasia, where teratozoospermia (short tails and thick flagella) is the cardinal feature and ejaculated sperm can be motile (more than 1000 polypeptides have been identified in the constitution of the cilium), and sperm concentrations can be normal or even high32,33. Although no specific genes have been linked to this disease, the inheritance pattern in family pedigrees suggests that it is likely to be autosomal recessive. ICS is caused by mutations on genes which encode dynein axoneme chains (DNAI). The ICS that maps 9p21–p13 (CILD1) is caused by a mutation in DNAI1. Another form (CILD2) is caused by mutation on 19q13.2–qter. Other loci for the disorder have been mapped to 5p (CILD3, DNAH5 gene), 16p12 and 15q13. Because the gene defect is usually recessive, offspring are likely to be normal; still, genetic counseling is recommended when assisted reproductive techniques are used33.

Autosomal dominant polycystic kidney disease Numerous large cysts of the kidneys, liver, pancreas and spleen, and a 10–40% chance of

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MALE INFERTILITY

developing berry aneurysms in the brain, characterize this disorder. Because the syndrome is often asymptomatic until adulthood, affected men may initially present with infertility. Cysts in the epididymis and seminal vesicles or ejaculatory ducts can obstruct the ductal system and cause infertility. Three separate genetic loci have been associated with autosomal dominant polycystic kidney disease (ADPK). PKD1 accounts for 85% of the disease and has been mapped onto chromosome 16p13.3, where it encodes a receptor-like integral membrane protein involved in cell–cell and cell–matrix interaction. A mild form (PKD2) has been mapped to chromosome 4q21–23, and it encodes a non-specific calcium-permeable channel; another variant, PKD3, is currently unmapped34. An association between men with ICS and with ADPK disease has recently been observed. Electron microscopy studies have revealed abnormalities on both the flagellar dynein arms and the cilium of the kidney epithelium33.

Cystic fibrosis transmembrane regulator mutations Cystic fibrosis (CF) is the most common fatal autosomal-recessive disease in Caucasians, with an incidence of 1 : 2500 births and a carrier frequency of 1 : 25. Clinical features of CF include chronic pulmonary obstruction and infection, exocrine pancreatic insufficiency, neonatal meconium ileus and male infertility35. The CF gene, cystic fibrosis transmembrane regulator (CFTR; 7q31.2), encodes a protein that regulates the cyclic adenosine monophosphate chloride channel that controls the transport of electrolytes in many secretory epithelia. More than 1000 mutations have been identified in the CFTR gene36, encompassing about 90% of cases of CF. The CFTR gene also influences the formation of the seminal vesicles, the vas deferens and the distal two-thirds of the epididymis37. More than 95% of men with CF have abnormalities in Wolffian duct-derived structures, manifesting most

commonly as congenital bilateral absence of the vas deferens (CBAVD).

Congenital bilateral absence of the vas deferens This condition occurs in 1–2% of infertile men38, and is considered a genital form of cystic fibrosis39. These patients exhibit the same spectrum of Wolffian duct defects as seen in those with fullblown cystic fibrosis, but generally lack the severe pulmonary, pancreatic and intestinal problems. Spermatogenesis is normal in approximately 90% of men with CBAVD40. Anatomically, the body and tail of the epididymis, the vas and the seminal vesicles may be absent, but the efferent ducts and the caput epididymis are almost always present41. It is thought that CBAVD is based on allelic patterns (homozygous and compound heterozygous) similar to typical CF but with less severe mutations42. The combination of the 5T (thymidines) allele in one copy of the CFTR gene (lack of exon 9), and a CF mutation (most commonly ∆F508) in the other copy, is peculiar for men with CBAVD. Therefore, it is important to include the 5T variants (intron 8) in the genetic screening for CF in patients and their partners before using assisted reproductive technologies (ART).

Congenital unilateral absence of the vas deferens Another male infertility phenotype (possibly associated with CFTR mutations but still controversial) affects 0.5% of the general population, and only rarely presents with infertility43. Almost 40% of patients with congenital unilateral absence of the vas deferens (CUAVD) have been reported to have at least one mutation in CFTR. CUAVD is more frequent on the left side (70%), and may be associated with contralateral renal agenesis (75%). However, if CUAVD is associated with renal agenesis, the possibility of finding a CFTR mutation is lower (31%)44.

CHROMOSOMAL AND GENETIC ABNORMALITIES IN MALE INFERTILITY

Table 16.4

245

Syndromes associated with androgen receptor gene mutations

Complete androgen insensitivity syndrome

Partial androgen insensitivity syndrome

Testicular feminization syndrome (Morris’s syndrome)

Male pseudohermaphroditism Lub’s syndrome Reifenstein’s syndrome Gilbert–Dreyfus syndrome

Table 16.5

Exons of the androgen receptor gene (AR) involved in androgen sensitivity

Exon 1

Transactivation domain function (TAD) modulates transcriptional activity of AR downstream genes

Exons 2 and 3

Encode a peptide domain responsible for DNA-binding domain

Exons 4 and 8

Encode C-terminal peptide domain responsible for androgen binding

Androgen receptor gene mutations The androgen receptor (AR) is a large steroid receptor whose gene is located on the X chromosome (Xq11–q12), and is essential for masculinization (fetal life) and virilization. AR mutations result in absent or structurally altered AR (functional impairment), causing partial or complete resistance to androgens (Table 16.4). The phenotype is variable, ranging from complete insensitivity (female phenotype) to normally virilized but infertile males. Clinical features include ambiguous genitalia, testicular atrophy, micropenis and hypospadias45. Over 300 distinct mutations have been reported in the AR. Mutations in exon 1 cause complete androgen insensitivity, while some mutations in the C-terminal ligand-binding domain (LBD) cause partial insensitivity45. Due to variable phenotypes, it has been proposed that as many as 40% of men with partial or totally impaired spermatogenesis may have subtle androgen insensitivity as an underlying cause46. A recent report found that only 2% of males with idiopathic infertility carried a significant variation within the AR gene47. The AR gene

includes eight exons (three domains) (Table 16.5), and has a critical region on exon 1 of cytosineadenosine-guanine (CAG) nucleotide repeats, formerly called the transactivation domain (TAD), usually between 15 and 30 repeats in number. Variation in length of this domain (> 40) results in severe spinal–bulbar muscular atrophy (Kennedy’s disease)48. This debilitating, late-onset (after 30 years of age) disorder consists of progressive degeneration of the anterior motor neurons and muscular weakness, as well as infertility due to testicular atrophy49. Although still controversial, some men may have oligozoospermia and intermediate lengths of CAG repeats (i.e. > 30 but fewer than 40). In these instances, with the phenomenon of genetic anticipation, offspring may inherit a larger number of CAG repeats than those of their parent, and when they reproduce (second generation) may have a child with Kennedy’s disease50,51.

Myotonic dystrophy Myotonic dystrophy (MD) is the most common cause of adult-onset muscular dystrophy, and usually presents with cataracts, muscle weakness and

246

MALE INFERTILITY

wasting, hypogonadism, electrocardiogram changes, diabetes (5% of cases) and cholelithiasis (25%). Symptoms usually become evident in the adult as early as in the second decade. The gene involved is located on the long arm of chromosome 19, region q13.2–3 (DMPK gene), and encodes the serine/threonine protein kinase family (myotonin-1). In MD there is an expansion (more than 35 repeat motifs) of the CTG sequence in the 3′-untranslated region of exon 5. Since reduced gene function correlates with the degree of repeat expansion, the severity of the condition varies with the number of repeats: normal individuals have between 5 and 35 CTG copies, mildly affected persons have between 50 and 80 copies and severely affected patients can have 2000 or more copies52. Like Kennedy’s disease, this disorder is characterized by anticipation, in which amplification (anticipation) of the disease is observed in parent-to-child transmission, especially from mother to offspring53. Male infertility is observed in about 30% of subjects, whilst some degree of testicular atrophy occurs in at least 80% of males suffering from this disorder (seminiferous tubules are more involved (75%) than Leydig cells). FSH and LH levels are elevated, with normal testosterone levels. Despite these findings, 66% of married men with MD can conceive naturally. A recent report described an association between MD and defective sperm capacitation and the acrosome reaction54.

Usher’s syndrome This is the most common cause of deafness–blindness in humans. This autosomal-recessive defect maps onto three chromosomes and results in three different phenotypes (US1 (14q32), US2 (1q41), US3 (3q21–q25)). Recently an association between Usher’s syndrome and infertility has been reported54. The common denominator for these associations is an abnormality in the ciliary structure of the sperm and the photoreceptor cells, since they share docosahexaenoic acid (DHA). DHA blood levels are less than normal in patients

with retinitis pigmentosa (RP), and sperm of patients with RP have reduced motility and abnormal morphology. Patients with Usher’s syndrome type II have the most pronounced reductions of DHA in the sperm55,56.

β-Thalassemia and sickle cell anemia Autosomal-dominant genomic deletions involving the β-globin gene (HBB), 11p15.4, account for approximately 10% of all β-thalassemia mutations. At least 60 different deletions have been described to date. Clinical features range from mild anemia (trait) to hemolytic anemia (transfusion-dependent) and iron overload (major thalassemia). Infertility results from the deposition of iron in the pituitary gland and testes. At the molecular level, it is hypothesized that iron overload may induce, via reactive oxygen species (ROS), sperm DNA oxidation and alter sperm membranes57. Sickle cell anemia is an autosomal-recessive genetic disease that results from the substitution of valine for glutamic acid at 11p15.5 of the HBB, responsible for a defective form of hemoglobin, hemoglobin S (HbS). Pituitary and testicular microinfarcts from sickle cell disease account for secondary hypogonadism and infertility58.

SRY gene defects SRY (sex determining region on Y chromosome) gene is located on the short arm of the Y chromosome (Yp11.3), and is important for determining ‘maleness’. The SRY gene encodes a transcription factor, a member of the HMG-box family (DNA-binding proteins) formerly called testisdetermining factor (TDF), which initiates male sex differentiation. Mutations in this gene (1 : 25 000) give rise to XY females (Xp22.11–p21.2) with gonadal dysgenesis (Swyer’s syndrome); translocation of SRY to the X chromosome causes the XX male phenotype. All 46,XX men are sterile due to absence of the long arm of the Y chromosome containing the

CHROMOSOMAL AND GENETIC ABNORMALITIES IN MALE INFERTILITY

azoospermia factor (AZF) gene, which is necessary for normal spermatogenesis, but their external genitalia and testes are developed under the influence of the Y-chromosome genetic fragment present on the X chromosome59.

α-Reductase deficiency 5α A deficiency in the 5α-reductase type-2 isozyme produces a form of male pseudohermaphroditism (autosomal recessive) due to the lack of conversion of testosterone to dihydrotestosterone (DHT). There are two genes encoding 5α-reductase: type 1 has been mapped onto chromosome 5, while type 2 has been mapped onto chromosome 2p23 (SRD5A2 gene). Mutations in isozyme 2 are associated with low DHT (important for prostate and external genitalia development) in spite of high levels of testosterone. Clinical features include normal internal genital ductal structures and testes, but incompletely virilized external genitalia. Affected individuals exhibit perineoscrotal hypospadias and often a vaginal pouch. Generally, the testes are found in the labioscrotal folds or the inguinal canal, the seminal vesicles are rudimentary and the prostate may be absent60. Infertility results from the structural abnormalities of the external genitalia. Although spermatogenesis has been described in descended testes, natural fertility has not been reported52.

247

dysgenesis may be caused by cytogenetic mosaicism or by mutations in testis-organizing genes near to the SRY region. One of these genes may be the newly cloned human testatin gene (20p11.2), a putative cathepsin inhibitor that is expressed early in testis development, just after SRY expression62. Scrotal testes may be associated with inguinal hernias, and almost uniformly reveal seminiferous tubules with Sertoli cell-only and normal Leydig cells. The dysgenetic gonad is predisposed to malignant degeneration (one-third of patients) to gonadoblastoma or dysgerminoma, typically before puberty63.

MALE INFERTILITY WITH CHROMOSOMAL ABERRATIONS Chromosomal disorders are defined as the loss, gain or abnormal arrangement of genetic material at the chromosome level. These disorders can be further divided into numerical and structural abnormalities. Structural chromosome disorders can occur in single (deletions, duplications and inversions) or multiple (translocations) chromosomes. Usually they are a consequence of breakage that occurs during meiosis, and are becoming more frequently recognized as a contributing factor to male infertility (15% of azoospermic and 5% of oligozoospermic men)64.

Mixed gonadal dysgenesis

Klinefelter’s syndrome

In males and females, mixed gonadal dysgenesis is a heterogeneous condition characterized by a unilateral testis on one side and a streak gonad on the opposite side. The phenotype ranges from normal males to patients with ambiguous external genitalia or females, depending on the amount of testosterone secreted by the testis. Genotypically, patients are usually 46,XY or 45,X/46,XY mosaicism (most common), both of which are associated with impaired gonadal development61. Since mutations in the SRY gene have not been detected (80% have normal SRY), gonadal

Klinefelter’s syndrome (1 : 1000) is the most common genetic reason for azoospermia, accounting for about 14% of cases65. It is associated with a triad of clinical findings: small, firm testes (devoid of germ cells), azoospermia and possibly gynecomastia52. The phenotype can vary from a normal, virilized man to one with stigmata of androgen deficiency. Testicular histology shows hyalinization of the seminiferous tubules with Leydig cell hyperplasia4. This syndrome may also be associated with tall stature, female hair distribution, low intelligence

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MALE INFERTILITY

quotient (IQ), lower-extremity varicosities, obesity, diabetes, increased incidence of leukemia and non-seminatous extragonadal germ-cell tumors, and breast cancer (20-fold higher than in normal males)66. About 90% of men have the classic 47,XXY genotype; the remaining (10%) are mosaic, with a combination of XXY/XY chromosomes (30 recognized mosaic patterns). Approximately 50% of XXY cases are paternally inherited, and a recent study suggested a relationship with advanced paternal age67. The extra X chromosome might originate in paternal meiosis I (nondisjunction of the XY bivalent in 50% of cases), or in maternal meiosis I or II (40% of cases), associated with maternal age68. Natural paternity with this syndrome is possible, but almost exclusively with the mosaic genotype69. Despite a uniformly abnormal somatic genotype, 75–100% of mature sperm from 47,XXY patients have a normal haploid sex chromosome complement (X or Y instead of XY or YY)70. The absence of significant gonosomal aneuploidy with somatic aneuploidy suggests that abnormal germ-cell lines are eliminated from further development at meiotic checkpoints within the testis52.

Noonan’s syndrome This syndrome is relatively common, with an estimated incidence of 1 : 1000–2500 live births. Noonan’s syndrome (NS) patients are phenotypically equivalent to those with Turner’s syndrome (XO), and share similar characteristics, i.e. webbed neck, short stature, lymphedema, low-set ears, wide-set eyes, cubitus valgus, cardiovascular disorders and pulmonary stenosis. This syndrome is inherited in an autosomal dominant pattern with karyotype 46,XY/XO mosaicism. A recently identified genetic locus at 12q24.2–q24.31 (PTPN11 candidate gene) could be involved in encoding a protein–tyrosine phosphatase that plays a role in the cellular response to extracellular signaling74. A second type of NS (type 2) appears to be transmitted in an autosomal recessive pattern. Typically, type 2 NS patients have hypertrophic obstructive cardiomyopathy, as opposed to 10–20% in the classical NS75. Fertility impairment is due to defects in spermatogenesis associated with cryptorchidism (77% at birth) and elevated FSH76.

Chromosomal translocations XYY syndrome The XYY syndrome has an incidence of 1 : 1000 live births. Fewer than 2% of men with the 47,XYY karyotype may be infertile71. The extra Y chromosome commonly (86%) originates through paternal meiotic II nondisjunction, while the remaining cases are due to postzygotic events72. The phenotype includes tall stature, aggressive and antisocial behavior and a higher risk of leukemia3. Studies that have focused on the chromosomal complements in mature sperm from XYY men show that very few sperm (< 1%) have sexchromosomal disomy (YY, XX, XY)73. This finding supports the hypothesis that the extra Y chromosome is eliminated at meiotic checkpoints during spermatogenesis, and shows that men with 47,XYY syndrome can father offspring with normal karyotypes.

Chromosomal translocations are classified as Robertsonian (incidence 1 : 900) if they involve chromosome 13, 14, 15, 21 or 22, or reciprocal (incidence 1 : 625) if any other chromosome is involved. If there is no gain or loss of chromosome material, the translocation is considered to be ‘balanced’ (unaffected phenotype). The reproductive risk with a balanced translocation is that sperm can carry an unbalanced chromosome, leading to pregnancy loss. Reciprocal translocations can lead to reduced fertility, spontaneous abortions or birth defects, depending on the chromosomes involved and the nature of the translocation76. Many translocations have been associated with male infertility. In particular, reciprocal and Robertsonian translocations (Robertsonian chromosomes are involved in as many as 15 different

CHROMOSOMAL AND GENETIC ABNORMALITIES IN MALE INFERTILITY

translocations) are at least 8.5-fold more common in infertile men than in randomly selected males. The most common Robertsonian translocation observed in infertile males is t(13q14q), where abnormal autosome rearrangement in meiosis causes spermatogenesis impairment. Carriers of another Robertsonian translocation involving chromosomes 14 and 21 (t(14;21)) are at risk for pregnancy loss and for offspring with Down’s syndrome and birth defects77.

Chromosomal inversions An inversion occurs when a chromosome breaks in two places and the material between the breakpoints rotates 180°, hence reversing the order of the chromatin (incidence 1 : 1000). Such rearrangements may either interrupt important genes at the breakpoint, or interfere with normal chromosome pairing during meiosis, because of imbalances in chromosomal mass. Autosomal inversions, particularly those involving chromosome 9, are eight-fold more likely to occur in infertile than in fertile men. These types of chromosomal derangements tend to be balanced and result in phenotypically normal males, but with severe oligoasthenoteratospermia or azoospermia1,76.

Y chromosome microdeletions Structural changes (loss or microdeletions) of various regions of the short or long arm of the Y chromosome could result in the breakdown of spermatogenesis, and are the second most frequent genetic causes of infertility. Microdeletions derive from the homologous recombination of identical segments within palindromic sequences. The spermatogenesis region on Yq11 associated with infertility is known as azoospermia factor (AZF). The AZF region is subdivided into AZFa (proximal), AZFb (central), AZFc (distal) and AZFd (actually AZFc proximal region), and the loss of any part of these regions can result in a variety of spermatogenic and infertility phenotypes78. Transcription units in these regions (Table

Table 16.6 genesis

249

Candidate genes involved in spermato-

Region

Gene involved

AZFa AZFb AZFc

USP9Y, DBY, UTY RMBY, EIF1A, CDY DAZ

AZF, azoospermia factor

16.6) encode proteins (mostly RNA-binding proteins) involved in the regulation of spermatogenesis via translational control. More than 30 Y-chromosome genes and gene families have been identified, although their function in spermatogenesis has not been completely detailed. Moreover, in the region of AZFc, the presence of partial deletions can also be observed in normal males79. Deletions are more frequent in the AZFc region (50–60%), involving the DAZ gene (deleted in azoospermia). In almost 50% of patients with DAZ deletions (AZFc) it is possible to find sperm in the ejaculate. For azoospermic patients, sperm can be retrieved by testicular biopsy (testicular sperm extraction or TESE)80. Incomplete spermatogenesis with no evidence of elongated spermatids or sperm in TESE has been reported in patients with a complete AZFb deletion (frequency 15%)81. Deletions in the AZFa region (frequency of 2–5%) are mostly associated with Sertoli cell-only (SCO) syndrome (75%), and overall, about 9% of men with SCO have a complete AZFa deletion82. Infertile men with non-obstructive azoospermia and those with sperm concentrations below 5 million/ml (severe oligozoospermia) should be offered testing for Y chromosome microdeletions. Overall, severe oligozoospermic patients have about a 4–6% risk of Y microdeletions83, while patients with non-obstructive azoospermia have a 14% risk of Y microdeletions84,85. Y chromosome microdeletions may be passed on to a male

250

MALE INFERTILITY

offspring through ICSI86; thus, genetic counseling is recommended. Some infertile men may actually be genetic mosaics and harbor DAZ deletions only in germ line (gamete) tissue and not in somatic cells87, and thus many escape recognition with the common practice of DNA analysis from peripheral leukocytes.

may be the ‘presenting symptoms’ or phenotype of a variety of pathologies that can affect non-reproductive organs. Examples are men with congenital absence of the vas deferens whose etiology has been linked to cystic fibrosis; men with the immotile cilia syndrome and some of its variants (such as sperm fibrous sheath dysplasia), where the presenting symptoms can be chronic sinusitis or bronchiectasis; or male infertility associated with polycystic kidney disease or the rare spinobulbar muscular atrophy. Many more forms of male infertility with a possible genetic etiology are still unrecognized. The time has come to associate phenotype with genotype in a more detailed and comprehensive manner. This requires the availability of modern molecular genetic testing and collaboration between andrologists/urologists, reproductive endocrinologists and genetic counselors. Notwithstanding the current limitations to identifying genetic ‘syndromes’ associated with male

Summary The current genetic screening offered before ICSI reveals that 35% of men with non-obstructive azoospermia (20% abnormal karyotype and 15% genetic or Y deletions), and about 10% of men with severe oligozoospermia (5% abnormal karyotype and 5% genetic or Y deletions), have a genetic explanation for their absent or reduced spermatogenesis. It is becoming clearer that abnormalities, both qualitative and quantitative, of spermatogenesis

Azoospermia

Obstructive

Non-obstructive

CFTR

Karyotype Y chromosome

Normal

? AR

Abnormal (35%)

Oligozoospermia (< 5 × 106 sperm/ml)

Teratozoospermia

Asthenozoospermia

Karyotype Y chromosome

Karyotype

Normal

Abnormal (10%)

Normal

FISH EM

CFTR ? AR FISH

Abnormal (5%)

Normal

Abnormal (5%)

EM FISH ICS

Genetic counseling

ICSI PGD, CVS or aminocentesis

Figure 16.1 Algorithm for genetic evaluation of the infertile male undergoing intracytoplasmic sperm injection (ICSI). EM, electron microscopy; CFTR, cystic fibrosis transmembrane conductance regulator gene; AR, androgen receptor; PGD, preimplantation genetic diagnosis; CVS, chorionic villi sampling; FISH, fluorescence in situ hybridization; ICS, (gene screening for) immotile cilia syndrome or Kartagener’s syndrome

CHROMOSOMAL AND GENETIC ABNORMALITIES IN MALE INFERTILITY

infertility, a review of the literature on the health of offspring born after ICSI (for severe male infertility) has shown that the rate of chromosomal anomalies, compared with the general neonatal population, is increased. This slight increase is seen in the de novo sex aneuploidy rate (0.6% vs. 0.2%) and in structural autosomal abnormalities (0.4% vs. 0.07%), and is believed to be linked to the very reason for infertility in the fathers. In summary, before undergoing ICSI, every male with idiopathic infertility should be fully evaluated and submitted to a minimum of genetic testing that includes karyotype, Y chromosome deletions and the androgen receptor. Additional genetic information could be gathered by using fluorescence in situ hybridization (FISH) on spermatozoa, since both azoospermic and oligozoospermic males have an increased risk of carrying a gene defect or aneuploid chromosomes. The algorithm shown in Figure 16.1 suggests a common genetic evaluation of the infertile male prior to and after ICSI.

10.

11.

12.

13.

14.

15.

16.

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24. Walsh CH. Non-diabetic endocrinopathy in hemochromatosis. In Barton JC, Edwards CQ, eds. Hemochromatosis: Genetics, Pathophysiology, Diagnosis and Treatment. Cambridge: Cambridge University Press, 2000: 278 25. Franchini M, Veneri D. Recent advances in hereditary hemochromatosis. Ann Hematol 2005; 84: 347 26. Worwood M. Inherited iron loading: genetic testing in diagnosis and management. Blood Rev 2005; 19: 69 27. Delague V, et al. Nonprogressive autosomal recessive ataxia maps to chromosome 9q34–9qter in a large consanguineous Lebanese family. Ann Neurol 2001; 50: 250 28. Tranebjaerg L, et al. Genome-wide homozygosity mapping localizes a gene for autosomal recessive nonprogressive infantile ataxia to 20q11–q13. Hum Genet 2003; 113: 293 29. Silveira LF, MacColl GS, Bouloux PM. Hypogonadotropic hypogonadism. Semin Reprod Med 2002; 20: 327 30. Semple RK, et al. Two novel missense mutations in G protein-coupled receptor 54 in a patient with hypogonadotropic hypogonadism. J Clin Endocrinol Metab 2005; 90: 1849 31. Peeraer K, et al. Pregnancy after ICSI with ejaculated immotile spermatozoa from a patient with immotile cilia syndrome: a case report and review of the literature. Reprod Biomed Online 2004; 9: 659 32. Westlander G, et al. Different fertilization rates between immotile testicular spermatozoa and immotile ejaculated spermatozoa for ICSI in men with Kartagener’s syndrome: case reports. Hum Reprod 2003; 18: 1286 33. Chemes HE. Phenotypes of sperm pathology: genetic and acquired forms in infertile men. J Androl 2000; 21: 799 34. Al-Bhalal L, Akhtar M. Molecular basis of autosomal dominant polycystic kidney disease. Adv Anat Pathol 2005; 12: 126 35. Daudin M, et al. Congenital bilateral absence of the vas deferens: clinical characteristics, biological parameters, cystic fibrosis transmembrane conductance regulator gene mutations, and implications for genetic counseling. Fertil Steril 2000; 74: 1164 36. Quinzii C, Castellani C. The cystic fibrosis transmembrane regulator gene and male infertility. J Endocrinol Invest 2000; 23: 684 37. Patrizio P, Salameh WA. Expression of the cystic fibrosis transmembrane conductance regulator (CFTR) mRNA in normal and pathological adult

38.

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50.

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human epididymis. J Reprod Fertil Suppl 1998; 53: 261 Bernardino LF, Lima CE, Zats M. Analysis of mutations in the cystic fibrosis transmembrane regulator (CFTR) gene in patients with obstructive azoospermia. Genet Mol Biol 2003; 26: 1 Patrizio P, Zielenski J. Congenital absence of the vas deferens: a mild form of cystic fibrosis. Mol Med Today 1996; 2: 24 Silber SJ, Patrizio P, Asch RH. Quantitative evaluation of spermatogenesis by testicular histology in men with congenital absence of the vas deferens undergoing epididymal sperm aspiration. Hum Reprod 1990; 5: 89 Patrizio P, et al. Correlation between epididymal length and fertilization rate in men with congenital absence of the vas deferens. Fertil Steril 1994; 61: 265 Phillipson GTM, Petrucco OM, Matthews CD. Congenital bilateral absence of the vas deferens, cystic fibrosis mutation analysis and intracytoplasmic sperm injection. Hum Reprod 2000; 15: 431 Mulhall JP, Oates RD. Vasal aplasia and cystic fibrosis. Curr Opin Urol 1995; 32: 316 Casals T, et al. Heterogeneity for mutations in the CFTR gene and clinical correlations in patients with congenital absence of the vas deferens. Hum Reprod 2000; 15: 1476 Gottlieb B, et al. The androgen receptor gene mutations database (ARDB): 2004 update. Hum Mutat 2004; 23: 527 Aiman J, et al. Androgen insensitivity as a cause of infertility in otherwise normal men. N Engl J Med 1979; 300: 223 Hiort O, et al. Significance of mutations in the androgen receptor gene in males with idiopathic infertility. J Clin Endocrinol Metab 2000; 85: 2810 La Spada AR, et al. Androgen receptor gene mutations in X-linked spinal and bulbar muscular atrophy. Nature 1991; 352: 77 Pinsky L, Beitel LK, Trifiro MA. Spinobulbar muscular atrophy. In Scriver CR, et al., eds. Metabolic and Molecular Basis of Inherited Disease. New York: McGraw-Hill, 2000: 4147 Casella R, et al. Significance of the polyglutamine tract polymorphism in the androgen receptor. Urology 2001; 58: 651 Patrizio P, et al. Larger trinucleotide repeat size in the androgen receptor gene of infertile men with extremely severe oligozoospermia. J Androl 2001; 22: 444

CHROMOSOMAL AND GENETIC ABNORMALITIES IN MALE INFERTILITY

52. Turek PJ, Pera RA. Current and future genetic screening for male infertility. Urol Clin North Am 2002; 29: 767 53. Vogt P. Molecular genetics of human male infertility: from genes to new therapeutic perspectives. Curr Pharm Des 2004; 10: 471 54. Hortas ML, Castilla JA, Gil MT. Decreased sperm function of patients with myotonic muscular dystrophy. Hum Reprod 2000; 15: 445 55. Connor WE, et al. Sperm abnormalities in retinitis pigmentosa. Invest Ophthalmol Vis Sci 1997; 38: 2619 56. Tosi MR, et al. Clinicopathologic reports, case reports, and small case series: Usher syndrome type 1 associated with primary ciliary aplasia. Arch Ophthalmol 2003; 121: 407 57. Perera D, et al. Sperm DNA damage in potentially fertile homozygous β-thalassaemia patients with iron overload. Hum Reprod 2002; 17: 1820 58. Barden EM, et al. Body composition in children with sickle cell disease. Am J Clin Nutr 2002; 76: 218 59. Layman LC. Human gene mutations causing infertility. J Med Genet 2002; 39: 153 60. Hedia S, et al. Male pseudo-hermaphroditism due partial 5 alpha-reductase deficiency, a case report. Tunis Med 2001; 79: 261 61. Alvarez-Nava F, Soto M, Borjas L. Molecular analysis of SRY gene in patients with mixed gonadal dysgenesis. Ann Genet 2001; 44:155 62. Eriksson A, et al. Isolation of the human testatin gene and analysis in patients with abnormal gonadal development. Mol Hum Reprod 2002; 8: 8 63. Chemes H, Muzulin PM, Venara MC. Early manifestations of testicular dysgenesis in children: pathological phenotypes, karyotype correlations and precursors stages of tumor development. APMIS 2003; 111: 12 64. Nudell DM, Pagani R, Lipshultz LI. Indications for genetic evaluation of men in a reproductive medicine program. Braz J Urol 2001; 27: 105 65. Rives N, et al. Assessment of sex chromosome aneuploidy in sperm nuclei from 47,XXY and 46,XY/47,XXY males: comparison with fertile and infertile males with normal karyotypes. Mol Hum Reprod 2000; 6: 107 66. Brugh VM III, Maduro MR, Lamb DJ. Genetic disorders and infertility. Urol Clin North Am 2003; 30: 143 67. Lowe X, et al. Frequency of XY sperm increases with age in fathers of boys with Klinefelter syndrome. Am J Hum Genet 2001; 69: 1046

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68. Hargreave T. Genetic basis of male infertility. Br Med Bull 2000; 3: 650 69. Poulakis V, et al. Birth of two infants with normal karyotypes after intracytoplasmic injection of sperm obtained by testicular extraction from two men with nonmosaic Klinefelter. Fertil Steril 2001; 76: 1060 70. Bergère M, et al. Biopsed testis cells of four 47,XXY patients: fluorescence in-situ hybridization and ICSI results. Hum Reprod 2002; 17: 32 71. Prabhakara MG, et al. Genetic analysis of infertile males from Bangalore, India. Presented at the 54th Annual Meeting of the American Society of Human Genetics, Toronto, Canada, 2004, 920/T-182 72. Rives N, et al. Meiotic segregation of sex chromosomes in mosaic and non-mosaic XYY males: case reports and review of the literature. Int J Androl 2003; 26: 242 73. Wang IY, et al. Fluorescence in-situ hybridization analysis of chromosomal constitution in spermatozoa from a mosaic 47,XXY/46,XY male. Mol Hum Reprod 2000; 6: 665 74. Tartaglia M, et al. Mutations in PTPN11, encoding the protein tyrosine phosphatase SHP-2, cause Noonan syndrome. Nat Genet 2001; 29: 465 75. Van der Burgt I, Brunner H. Genetic heterogeneity in Noonan syndrome: evidence for an autosomal recessive form. Am J Med Genet 2000; 94: 46 76. Griffin DK, Finch KA. The genetic and cytogenetic basis of male infertility. Hum Fertil 2005; 8: 19 77. Antonelli A, et al. Chromosomal alterations and male infertility. J Endocrinol Invest 2000; 23: 677 78. Krausz C, Forti G, McElreavey K. The Y chromosome and male fertility and infertility. Int J Androl 2003; 26: 70 79. Hucklenbroich K, et al. Partial deletions in the AZFc region of the Y chromosome occur in men with impaired as well normal spermatogenesis. Hum Reprod 2005; 20: 191 80. Mulhall JP, et al. Azoospermic men with deletion of the DAZ gene cluster are capable of completing spermatogenesis: fertilization, normal embryonic development and pregnancy occur when retrieved testicular spermatozoa are used for intracytoplasmic sperm injection. Hum Reprod 1997; 12: 503 81. Hoops CV, et al. Detection of sperm in men with Y chromosome microdeletions of the AZFa, AZFb and AZFc regions. Hum Reprod 2003; 18: 1660 82. Kamp C, et al. High deletion frequency of the complete AZFa sequence in men with Sertoli-cell-only syndrome. Mol Hum Reprod 2001; 7: 987

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83. Reijo R, et al. Severe oligozoospermia resulting from deletions of azoospermia factor gene on Y chromosome. Lancet 1996; 347: 1290 84. Reijo R, et al. Diverse spermatogenic defects in humans caused by Y chromosome deletions encompassing a novel RNA-binding protein gene. Nat Genet 1995; 10: 383 85. Foresta C, Moro E, Ferlin A. Y chromosome microdeletions and alterations of spermatogenesis. Endocr Rev 2001; 22: 226

86. Affara NA. The role of human and mouse Y chromosome genes in male infertility. J Endocrinol Invest 2000; 23: 630 87. Calogero AE, et al. Spontaneous transmission from father to his son of a Y chromosome microdeletion involving the deleted in azoospermia (DAZ) gene. J Endocrinol Invest 2002; 25: 631

17 Reactive oxygen species and their impact on fertility R John Aitken, Liga E Bennetts

INTRODUCTION

embryo as a consequence of aberrant DNA repair in the fertilized egg5,6. Thus, high rates of DNA damage in human spermatozoa have been associated with reduced rates of fertilization in vivo and in vitro, impaired preimplantation development of the embryo, increased rates of early pregnancy loss and high rates of morbidity in the offspring, including dominant genetic disease, infertility and cancer7–15. In light of these associations, attempts are now being made to define those factors responsible for the increased DNA damage and impaired functional competence seen in the spermatozoa of infertile males. As seen in the following section, of all the potential causes undergoing active consideration at the present time, oxidative stress appears to be amongst the most important. One of the first mechanisms suggested for the induction of genetic damage in defective human spermatozoa involved endonuclease-mediated cleavage of the DNA as a result of incomplete apoptosis during spermatogenesis16–18. While plausible, recent analyses of putative apoptotic markers in spermatozoa, such as the plasma membrane translocation of phosphatidylserine, have suggested that aberrant apoptosis is not highly correlated with DNA fragmentation in the male germ line19. It has also been hypothesized that the DNA damage seen in defective human spermatozoa results from defective chromatin packaging during a critical stage of spermiogenesis. This

Male infertility is a relatively common complaint that affects approximately one in 20 men in developed countries. Despite the prevalence of this condition, relatively little is known about the underlying pathophysiology. Indeed, since the advent of intracytoplasmic sperm injection (ICSI) as a therapeutic technique in 19921, the biomedical community has paid little attention to this problem. However, an appreciation of the etiology of male infertility will be essential if we are to optimize procedures for the management of this condition and contemplate strategies for its possible prevention. Unlike female infertility, the male counterpart is not, predominantly, an endocrine condition; it is a pathology affecting germ cells. Most infertile men produce spermatozoa; however, these gametes are characterized by functional deficiencies stemming from defects occurring during spermatogenesis or sperm maturation. Interest in the origins of male infertility has recently been stimulated by data indicating that spermatozoa from such patients not only suffer from an impaired capacity for fertilization but also may exhibit high rates of DNA damage to both the mitochondrial and nuclear genomes2–4. One of the consequences of such damage is a possible increase in the mutational load carried by the 255

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proposal envisages that relief of the torsional stresses associated with chromatin packaging involves the repeated transient nicking of DNA by topoisomerase. Defects in the structure of the chromatin, or the activity of the topoisomerase system itself, may lead to the generation of gametes expressing high levels of DNA fragmentation17,20. In support of this hypothesis is the observation that errors of chromatin packaging are, indeed, commonly associated with DNA damage in the germ line21. A third hypothesis is that defective sperm function and DNA damage in the male germ line are both mediated by high levels of oxidative stress. Excessive production or exposure to reactive oxygen species (ROS) has been both statistically and causally associated with defective sperm function and DNA damage in a large number of independent studies22–27. Furthermore, nuclear DNA damage in spermatozoa appears to exhibit a tighter association with markers of oxidative stress than with apoptosis19. In order to examine this association between defective sperm quality and oxidative stress in more detail, the next section introduces the fundamental chemistry of ROS and reviews the mechanisms by which they exert their pathological effects.

REACTIVE OXYGEN SPECIES AND LIPID PEROXIDATION The acronym ROS covers a wide range of metabolites derived from the reduction of molecular oxygen, including free radicals, such as the superoxide anion (O2–•), and powerful oxidants such as hydrogen peroxide (H2O2). The term also covers molecules derived from the reaction of carbon centered radicals with oxygen, including peroxyl radicals (ROO•), alkoxyl radicals (RO•) and organic hydroperoxides (ROOH). It may also refer to other powerful oxidants such as peroxynitrite (ONOO–) or hypochlorous acid (HOCl), as well as the highly biologically active free radical, nitric oxide (•NO).

The specific term ‘free radicals’ refers to any atom or molecule containing one or more unpaired electrons. As unpaired electrons are highly energetic, and seek out other electrons with which to pair, they confer upon free radicals considerable reactivity. Thus, free radicals and related ‘reactive species’ have the ability to react with, and modify the structure of, many different kinds of biomolecule, including proteins, lipids and nucleic acids. The wide range of targets that can be attacked by ROS is a critical aspect of their chemistry that contributes significantly to the pathological significance of these oxygen metabolites. The most commonly encountered oxygen free radical in biological systems is O2–•. When in aqueous solution, O2–• has a short half-life (1 ms) and is relatively inert. The radical is more stable and reactive in the hydrophobic environment provided by cellular membranes. The charge associated with O2–• means that this molecule is generally incapable of passing across biological membranes, although this molecule has been reported to exit cells using voltage-dependent anion channels. As a result of its lack of membrane permeability, O2–• may be more damaging if produced inside biological membranes than at other sites. It is also important to note that while O2–• can act as either a reducing agent or a weak oxidizing agent in aqueous solution, under the reducing conditions prevailing within cells, O2–• acts primarily as an oxidant. Since most biological molecules only have paired electrons, free radicals are also likely to be involved in chain reactions that can propagate the damage induced by ROS. A classic example of such a chain reaction is the peroxidation of lipids in biological membranes. In this process, a ROSmediated attack on unsaturated fatty acids generates peroxyl (ROO•) and alkoxyl (RO•) radicals that, in order to stabilize, abstract a hydrogen atom from an adjacent carbon, generating the corresponding acid (ROOH) or alcohol (ROH). The abstraction of a hydrogen atom from an adjacent lipid creates a carbon-centered radical that

IMPACT OF REACTIVE OXYGEN SPECIES

combines with molecular oxygen to recreate another lipid peroxide. In order to stabilize, the latter must again abstract a hydrogen atom from a nearby lipid, creating another carbon radical that combines with molecular oxygen to create yet another lipid peroxide. In this manner, a chain reaction is created that, if unchecked, would propagate the peroxidative damage throughout the plasma membrane, leading to a rapid loss of membrane-dependent functions28. The vulnerability of human spermatozoa to oxidative attack stems from the fact that these cells are particularly rich in unsaturated fatty acids29. Such an abundance of unsaturated lipids is necessary to create the membrane fluidity required by the membrane fusion events associated with fertilization, including acrosomal exocytosis and sperm–oocyte fusion. Unfortunately for spermatozoa, such unsaturated fatty acids are particularly prone to oxidative attack because the presence of a double bond weakens the C–H bonds on the adjacent carbon atoms, facilitating the hydrogen abstraction step and initiation of peroxidative damage, as indicated below: Bis-allylic methylene group Unsaturated fatty acid

R–CH=CH–CH2–CH=CH–R′ Hydrogen abstraction

Lipid radical

OH•

OOH• or

H2O

H2O2

R–CH=CH–CH–CH=CH–R′

Such lipid peroxidation chain reactions can be promoted by the presence of transition metals such as iron and copper that can vary their valency state by gaining or losing electrons. Significantly, there is sufficient free iron and copper in human seminal plasma to promote lipid peroxidation once this process has been initiated30. When iron sulfate and ascorbate (added as a reductant to maintain the iron in a reduced state) are added to suspensions of human spermatozoa, large amounts of lipid peroxide are generated. A majority of these peroxides arise from the iron-catalyzed propagation, rather than de novo initiation, of

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lipid peroxidation cascades31, according to the following equations: ROOH + Fe2+ lipid hydroperoxide

→

RO• + OH– + Fe3+ alkoxyl radical

ROOH + Fe3+ lipid hydroperoxide

→

ROO• + H+ + Fe2+ peroxyl radical

Thus, the amounts of lipid peroxide generated on the addition of transition metals, such as iron, to human sperm suspensions will reflect the amount of lipid peroxide present in these cells at the moment the catalyst was added. The lipid peroxide content of these cells will, in turn, reflect differences in the amount of oxidative stress that the spermatozoa have suffered during their life history. Differences in susceptibility arise because of interindividual variation in (1) the presence and molecular composition of unsaturated fatty acids in the sperm plasma membrane, (2) the degree to which the spermatozoa have been exposed to ROS and transition metal catalysis during their life history and (3) the level of protection afforded by free radical scavengers, chain-breaking antioxidants and ROS-metabolizing enzymes in the vicinity of the spermatozoa during their sojourn in the male reproductive tract. Monitoring the generation of lipid peroxide breakdown products such as malondialdehyde and/or 4-hydroxy alkenals in the presence of ferrous ion promoters therefore generates a significant amount of information about the sperm population under investigation32. Such measurements of the ‘lipoperoxidative potential’ of human spermatozoa have clear diagnostic value29,32. Protection against lipid peroxidation includes membrane-associated antioxidants epitomized by α-tocopherol, a hydrophobic vitamin that is capable of intercepting alkoxyl and peroxyl radicals and terminating the peroxidation chain reaction33. This vitamin is extremely effective in breaking lipid peroxidation cascades, and has been shown to improve significantly the fertility of males selected on the basis of high levels of lipid peroxidation in their spermatozoa34. Moreover,

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this vitamin has been known since the 1940s to be essential for male reproduction. Of the small-molecular-mass scavengers involved in the protection of human spermatozoa, the most important are vitamin C, uric acid, tryptophan and taurine35,36. In terms of antioxidant enzymes, spermatozoa possess both the mitochondrial and cytosolic forms of superoxide dismutase (SOD) and the enzymes of the glutathione cycle, but very little catalase. SOD catalyzes the dismutation of O2–•, a reaction in which this molecule reacts with itself to generate H2O2. Such dismutation can occur spontaneously without SOD; however, the reaction proceeds much more slowly in the absence of this enzyme. There is sufficient SOD activity in the mitochondria and cytosol of human spermatozoa to account for most, if not all, of the H2O2 produced by these cells37. Although SOD is usually thought of in antioxidant terms, this is only true if this enzyme is tightly coupled with additional enzymes that can metabolize the H2O2 generated as a consequence of O2–• dismutation. In isolation, SOD converts a short-lived, rather inert, membrane-impermeant free radical (O2–•) into a powerful, membrane-permeant oxidant, H2O2. Although the latter is not a free radical, it is, nevertheless, a potentially pernicious molecule. If not rapidly metabolized, it has the potential to initiate both lipid peroxidation in the sperm plasma membrane and trigger DNA damage to both the nuclear and mitochondrial genomes of these cells3,38. Some insight into the relative importance of O2–• and H2O2 in the initiation of peroxidative damage in human spermatozoa has come from studies employing xanthine oxidase to generate an extracellular mixture of ROS in vitro39. In the presence of this ROS-generating system, the spermatozoa rapidly lose their motility as a consequence of the initiation and propagation of peroxidative damage. If SOD is added to the medium to remove O2–•, motility loss still occurs. However, if catalase is added to the incubation mixture to remove H2O2, then lipid peroxidation is

suppressed and sperm motility is fully maintained. The implication of these studies, that H2O2 is the major cytotoxic species of ROS as far as spermatozoa are concerned, has been confirmed by experiments in which the direct addition of this oxidant has been shown to influence both the movement of human spermatozoa and their competence for oocyte fusion38. Given the damaging nature of H2O2 it is obviously important that this oxidant is rapidly removed from spermatozoa before it can initiate lipid peroxidation or DNA damage. The enzymes of the glutathione cycle (glutathione peroxidase and reductase) are responsible for peroxide metabolism in these cells. Under normal circumstances, sufficient NADPH (reduced nicotinamide– adenine dinucleotide phosphate) is generated by the oxidation of glucose through the hexose monophosphate shunt to fuel glutathione reductase and maintain an adequate pool of reduced glutathione (GSH) to counteract the H2O2 and lipid peroxides generated as a consequence of sperm metabolism40. These reactions can be summarized as follows: Glutathione reductase GSSG + NADPH + H+

→

2GSH + NADP+

Glutathione peroxidase → GSSG + 2H2O 2GSH + H2O2

where GSSG is glutathione disulfide. It should also be noted that the detoxification of lipid peroxides by glutathione peroxidase requires the concerted action of an additional enzyme in the form of phospholipase A2. This enzyme is required to cleave the lipid peroxide away from the parent phospholipid so that it becomes available for the detoxifying action of glutathione peroxidase. In addition to these intracellular antioxidants, spermatozoa are also protected by highly specialized extracellular antioxidant enzymes secreted by the male reproductive tract. These enzymes include glutathione peroxidase 5 (GPX5)41 as well as the extremely large amounts of extracellular SOD present in epididymal and seminal plasma29.

IMPACT OF REACTIVE OXYGEN SPECIES

(a) 110

Motility (%)

Indeed, seminal plasma contains more SOD than any other fluid in biology. The world record is held by donkey semen, which contains more than 3000 units of enzyme activity per milliliter42. As seen later in this chapter, the antioxidants present in seminal plasma (SOD, albumin, uric acid and vitamin C) become extremely important in protecting spermatozoa from ROS generated by activated leukocytes entering the reproductive tract at points distal to the epididymis, such as the urethra, prostate and seminal vesicles.

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10 0

EVIDENCE FOR OXIDATIVE STRESS

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Given the potential that ROS have for causing cellular damage, it is not surprising that they have been implicated in the etiology of male infertility22,26. The evidence for an association between oxidative stress and defective sperm function comes from three major sources. First, there is evidence that many aspects of sperm function including motility and sperm–oocyte fusion are negatively correlated with the lipoperoxidative potential of these cells. This was first suggested in the pioneering studies of Thaddeus Man and colleagues at the University of Cambridge. These authors observed that human spermatozoa were extremely susceptible to the cytotoxic effects of lipid peroxidation, and that severe sperm motility loss was associated with high levels of lipid peroxide generation in the presence of transition metals29,43. These studies have subsequently been confirmed and extended in larger cohorts of patients. Thus, the lipoperoxidative potential of freshly prepared spermatozoa (i.e. their capacity to generate lipid peroxides in the presence of a ferrous ion promoter) was found to be highly predictive of their capacity for movement and their ability to exhibit sperm–oocyte fusion32,44. Indeed, the tightness of the correlations with sperm movement has suggested that peroxidative damage is one of the major causes of impaired motility32 (Figure 17.1). Moreover, the lipoperoxidative potential of washed, leukocyte-free sperm

0.5

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2.5

MA + 4HA (µmol)

Figure 17.1 Relationship between motility loss observed in populations of human spermatozoa and generation of MA + 4HA in the presence of promoter. (a) Oxidative stress induced by the incubation of spermatozoa for 15 h at 37°C. (b) Oxidative stress induced using a xanthine oxidase free radicalgenerating system. MA + 4HA represents µmol of malondialdehyde and 4-hydroxy alkenals generated by 2 × 107 spermatozoa during a 2-h incubation with promoter32

suspensions was found to be reflective of the quality of sperm movement in the original ejaculate (Figure 17.2). Such findings reinforce the notion that the diagnostic value of lipoperoxidative potential measurements lies in the fact that

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(a)

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40 VCL (µm/s)

VAP (µm/s)

60 35 30

50 40

25 30

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20 15

30 20

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Figure 17.2 Relationships between lipoperoxidation potential of purified sperm suspensions and sperm movement in the original semen samples. (a) VAP (average path velocity); (b) VCL (curvilinear velocity); (c) percentage progressive; and (d) percentage rapid (> 25 µm/s). MA + 4HA represents µmol of malondialdehyde and 4-hydroxy alkenals generated by 2 × 107 spermatozoa during a 2-h incubation with promoter

they give an accurate picture of the accumulated degree of oxidative stress suffered by spermatozoa during their life history32,45. Additional evidence for oxidative stress in defective sperm populations comes from the elevated levels of oxidative DNA damage observed in

the spermatozoa of infertile men compared with fertile controls2,27,46. Positive correlations between sperm DNA damage and the intensity of signals generated in the presence of redox-active probes (luminol and lucigenin) tend to support this view19,46. Studies in which defective sperm

IMPACT OF REACTIVE OXYGEN SPECIES

function has been correlated with the chemiluminescence generated in the presence of such probes also add weight to this argument22,23,26,47. In studies involving clinically characterized samples, elevated chemiluminescence signals have been observed in particular groups of patients including those exhibiting oligozoospermia48, spinal cord injury49 and varicocele50. Significantly elevated chemiluminescence signals have also been observed in patients exhibiting unexplained infertility22,51. Of particular clinical importance is a prospective study in which the chemiluminescence signals generated in the presence of luminol were found to correlate with the incidence of spontaneous pregnancy in a large cohort of untreated patients followed up for a maximum of 4 years52. Moreover, within this data set there were no significant correlations between fertility and the conventional criteria of semen quality. Thus, such chemiluminescence measurements of redox activity in human sperm suspensions are clearly able to add value to the traditional semen analysis. The importance of such assays has also been emphasized in studies reporting significant inverse correlations between sperm chemiluminescence and the fertilizing potential of these cells in assisted conception cycles53. Although these data are suggestive, there are two notes of caution that should be raised in evaluating these associations between chemiluminescence and fertility. First, the biochemical basis of the activities being measured by luminol- or lucigenin-dependent chemiluminescence is still the subject of debate. In the case of lucigenin, a commonly used experimental paradigm is to trigger chemiluminescence in populations of spermatozoa through the addition of an exogenous electron source in the form of NAD(P)H54. Assays performed in this manner generate intense chemiluminescence signals with human spermatozoa that are inversely correlated with the functional competence of these cells47,55. The chemistry of lucigenin chemiluminescence is complex, but a key event in the biochemical cascade leading to light

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generation is the activation of the probe by a oneelectron reduction reaction. Such activation can be achieved enzymatically by cytochrome P450 or cytochrome b5 reductase56. Once activated, the probe is then thought to react with O2–• to create an unstable dioxetane that decomposes with the generation of light. However, it has also been proposed that reduced lucigenin can itself effect the one-electron reduction of ground-state oxygen to produce O2–• and regenerate the parent lucigenin molecule. If the concentration of NAD(P)H and lucigenin in the reaction mixture is sufficiently high, such redox cycling behavior has the potential to generate a large amount of O2–• as a consequence, rather than a cause, of probe activation. Doubts have been cast on the validity of this reaction scheme57 and, as a result, we cannot be certain what proportion of the chemiluminescent signal generated in the presence of lucigenin and NAD(P)H can be accounted for by the primary production of O2–• or the secondary production of this metabolite via the redox cycling of the probe. If the latter explanation is correct, it would suggest the presence of abnormally high levels of reductase activity in the spermatozoa of infertile men58. In the case of luminol, the probe must undergo a one-electron oxidation in order to become activated. In many ways, luminol is a more reliable probe than lucigenin, and has been effectively used to record the ROS generated in human semen samples as a consequence of leukocyte contamination59,60. However, herein lies the second point of contention with chemiluminescence data generated using human semen: the extent to which the results have been influenced by the presence of contaminating leukocytes.

SOURCES OF OXIDATIVE STRESS Although most studies in this area have been careful to exclude leukocytospermic specimens containing large numbers of leukocytes (typically > 1 × 106/ml), this does not necessarily mean that

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the data have not been obfuscated as a result of leukocyte contamination. On a cell-for-cell basis, the most common type of leukocyte found in human semen samples, the neutrophil, is 1000fold more active in generating ROS than a spermatozoon. Concentrations of leukocytes well below the threshold for leukocytospermia exhibit highly significant correlations with ROS generation by washed sperm suspensions, giving r values in the order of 0.861. Despite the highly significant nature of this correlation, it does not mean that spermatozoa are incapable of generating ROS. Although various publications have variously asserted that the chemiluminescent signals generated by washed human sperm suspensions emanate exclusively from the spermatozoa62 or contaminating leukocytes63, the truth is that both sources of ROS are active. Plots of leukocyte numbers against PMA-induced chemiluminescence activity (Figure 17.3) reveal that redox activity can vary over several log orders of magnitude in the absence of detectable leukocyte contamination. However, when leukocytes are present, the chemiluminescence activity is invariably high. In order

to resolve the spermatozoa’s contribution to oxidative stress in the ejaculate, it is essential that all traces of leukocyte contamination are removed from the sperm suspension. Protocols have been described for both the efficient detection of leukocyte contamination and the selective removal of these cells using paramagnetic particles coated with anti-CD45, the common leukocyte antigen64–66. However, there are very few studies in which these stringent conditions have been met. Where this has been achieved, the results unequivocally identify defective spermatozoa as a source of redox activity49. In a recent study, leukocyte-free sperm suspensions were exposed to the powerful protein kinase C agonist, 12-myristate, 13-acetate phorbol ester (PMA). The results revealed powerful inverse correlations between the chemiluminescence activity recorded and the quality of spermatozoa, particularly their motility32. Even more important, such measurements showed very tight correlations with the fundamental quality of the original semen sample in terms of sperm morphology, count and motility (Figure 17.4)32. In other words, the measurement

6.5

Log chemiluminescence

6 5.5 5 4.5 4 3.5 3 2.5 2 –0.25

0.25

0.5

0.75

1.25

1.5

1.75

2.25

Log 1 + 80% leukocyte concentration

Figure 17.3 Plot of leukocyte concentration against 12-myristate, 13-acetate phorbol ester (PMA)-induced, luminol peroxidasemediated chemiluminescence. Note the chemiluminescence signal generated by these samples varies over log orders of magnitude in the absence of leukocyte contamination

IMPACT OF REACTIVE OXYGEN SPECIES

(a)

263

(b) 100 200 Sperm concentration (106/ml)

Motility in semen (%)

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Log PMA-based chemiluminescence (counts/min)

3.5

4.5

5.5

6.5

Log PMA-based chemiluminescence (counts/min)

Figure 17.4 Relationships between intensity of the chemiluminescence signal generated by purified leukocyte-free samples in response to 12-myristate, 13-acetate phorbol ester (PMA) in the presence of luminol peroxidase and quality of the original semen samples as reflected by (a) the percentage of motile cells in semen and (b) sperm count in semen32

of ROS generation by spermatozoa not only reflects the quality of these cells but also the quality of the underlying spermatogenic process. Why spermatozoa should vary in their capacity for ROS generation is unknown at the present time. One possibility is that the oxidative stress is being generated by virtue of defects in the sperm mitochondria. Mitochondria are extremely active organelles that are constantly mediating electron transfer reactions through the ETC (electron transport chain) in order to fuel the generation of adenosine triphosphate (ATP). One of the inherent problems with such electron transport activity is that it is leaky, and electrons have a tendency to spill out of the ETC and combine with oxygen to generate O2–•. Aberrant production of ROS by mitochondria is therefore a possible source of oxidative stress in the spermatozoa of infertile men. However, early attempts to address this question failed to find any effect of ETC inhibitors on the chemiluminescence signals generated by suspensions of defective spermatozoa22.

The caveat with these experiments is that they did not exclude the possibility that the ROS being detected were generated by contaminating leukocytes. Thus, a possible contribution of sperm mitochondria to the generation of ROS by purified human sperm suspensions still requires careful examination. Another possibility, for which there is considerable evidence, is that the spermatozoa generating high levels of ROS have experienced defective spermiogenesis resulting in morphological defects, particularly in the midpiece region of the cell. During normal spermiogenesis, Sertoli cells actively remove the sperm cytoplasm, just before these cells are released from the germinal epithelium. In most mammals, any residual cytoplasm that remains after spermiogenesis is remodeled into a discrete, spherical, cytoplasmic droplet that slowly migrates down the sperm tail during epididymal transit, prior to its release into the extracellular space. Intriguingly, human spermatozoa have lost this ability to create and shed a

MALE INFERTILITY

cytoplasmic droplet. In these cells, any residual cytoplasm left after spermiation snaps back into the neck region of the spermatozoa and remains there as a ragged appendage that bears witness to the defective testicular origins of the cell. The presence of such excess residual cytoplasm has been correlated with ROS production by several independent groups67–70. One suggested mechanism by which such residual cytoplasm might induce ROS production is through the provision of excess substrate to a putative NADPH oxidase on the sperm surface. ROS production by purified sperm suspensions is highly correlated with the cellular content of cytoplasmic enzymes such as SOD, creatine kinase and glucose-6-phosphate dehydrogenase. Most of these enzymes are simply passengers, confirming the presence of excess residual cytoplasm in sperm populations generating high levels of ROS67,68. However, it has been hypothesized that in terms of pathology, the key enzyme is glucose6-phosphate dehydrogenase5,67,71. This enzyme controls the rate of glucose oxidation through the hexose monophosphate shunt, and the latter, in turn, generates the NADPH needed to fuel ROS production by a putative NADPH oxidase enzyme such as Nox 5, a free radical-generating oxidase recently detected in the male germ line72. This link between NADPH and ROS generation is reflected in the strong correlation that exists between the glucose-6-phosphate content of purified human sperm suspensions and their capacity to generate a chemiluminescence response to PMA (Figure 17.5). By removing most of the sperm cytoplasm during spermiogenesis, the testes ensure that these cells are only able to generate a limited supply of NADPH, just enough to meet the needs of the protective glutathione cycle and support the ROS-dependent elements of sperm capacitation73–76. However, if excess residual cytoplasm is retained because of mistakes during spermiogenesis (Figure 17.6), then there is the potential to generate additional ROS that will, in turn, damage the functional competence of these cells.

–1.5

Log G6PDH activity (/108 sperm)

264

–2.0

–2.5

–3.0

–3.5 3.5

4.5

5.5

6.5

Log PMA (counts/5 min)

Figure 17.5 Cellular content of glucose-6-phosphate dehydrogenase (G6PDH) and chemiluminescence. The retention of excess residual cytoplasm increases the cellular content of cytoplasmic enzymes such as G6PDH, the presence of which correlates closely with the redox activity exhibited by human spermatozoa in response to 12-myristate, 13-acetate phorbol ester (PMA) provocation in the presence of luminol and peroxidase

Figure 17.6 Individual spermatozoa exhibit considerable variation in the amount of residual cytoplasm retained following spermiation. Cytoplasm revealed by staining for diaphorase activity67

IMPACT OF REACTIVE OXYGEN SPECIES

CONSEQUENCES OF OXIDATIVE STRESS In light of the above, we must conclude that there are two sources of oxidative stress within the ejaculate: leukocytes and defective spermatozoa. The impact of seminal leukocytes will depend on the types of white cell present, their site of entry into the male reproductive tract and their state of activation. All of the information currently available indicates that the major leukocyte species is the neutrophil, and these cells are present in the ejaculate in an activated state61,62. Where these cells enter the male reproductive tract is generally unresolved, but has a direct bearing on the pathological consequences of leukocytic infiltration. If the leukocytes gain entry at points distal to the origin of the vas deferens, as a consequence of secondary sexual gland infection for example, then their direct impact on sperm function may be limited, because at the moment of ejaculation the spermatozoa will be protected by the powerful antioxidants in seminal plasma61. Conversely, if the neutrophils entered the male reproductive tract at the level of the rete testes or epididymis, then there would be every opportunity for these cells to induce oxidative damage in the spermatozoa. Free radical-generating leukocytes also have ample opportunity to attack spermatozoa in washed preparations, where the gametes are deprived of the protective effects of seminal plasma. Indeed, apart from albumin and possibly phenol red, most in vitro fertilization (IVF) media are devoid of protective antioxidants. Some media are even supplemented with transition metals such as iron and copper, and, in this way, may actually stimulate peroxidative damage in spermatozoa77. Whenever activated leukocytes are present in washed sperm suspensions, the fertilizing capacity of the spermatozoa is suppressed62. These results have clear implications for the practice of IVF therapy, and it comes as no surprise that negative associations have been observed between leukocyte contamination of washed sperm preparations and fertilization rates in assisted conception cycles65,66.

265

The second source(s) of ROS in human ejaculates are the spermatozoa themselves49,68,69. Such intracellular free radical generation is associated with the disruption of all aspects of sperm function, including their motility, their capacity for acrosomal exocytosis, their ability to fuse with the vitelline membrane of the oocyte and the integrity of their DNA6,27. As indicated above, excess free radical generation is normally associated with defects in spermiogenesis, leading to the retention of excess residual cytoplasm in the midpiece of these cells. It is also possible that excess ROS generation by spermatozoa is driven by the redox cycling of xenobiotics present in the environment, or deficiencies in the mitochondrial ETC6. Whether such ROS-generating spermatozoa can also damage the functional competence of other spermatozoa in the immediate vicinity is still an open question. If defective spermatozoa actively generate free radicals from the moment they leave the testes, then the opportunities for collateral damage to other cells in the same sperm population might be considerable.

CONCLUSIONS In summary, oxidative stress is one of the major causes of defective sperm function. Free radical attacks on these cells damage the DNA in the sperm nucleus and induce lipid peroxidation in the sperm plasma membrane. As a consequence of these changes, the spermatozoa lose their capacity for fertilization and their ability to support normal embryonic development6. The origins of oxidative stress include leukocytic infiltration, excess free radical generation by the spermatozoa and defects in the antioxidant protection provided to these cells during their sojourn in the male reproductive tract. Further research in this area should help to advance our understanding of the origins of oxidative stress in the male reproductive tract, and assist in the development of rational approaches towards the prevention and treatment of this condition.

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REFERENCES 1. Palermo G, et al. Pregnancies after intracytoplasmic injection of single spermatozoon into an oocyte. Lancet 1992; 340: 17 2. Irvine DS, et al. DNA Integrity in human spermatozoa: relationships with semen quality. J Androl 2000; 21: 33 3. Sawyer DE, et al. Analysis of gene-specific DNA damage and single-strand DNA breaks induced by pro-oxidant treatment of human spermatozoa in vitro. Mutat Res 2003; 529: 21 4. Lewis SEM, Aitken RJ. Sperm DNA damage, fertilization and pregnancy. Cell Tissue Res 2005; in press 5. Aitken RJ. The human spermatozoon – a cell in crisis? The Amoroso Lecture. J Reprod Fertil 1999; 115: 1 6. Aitken RJ. Founders’ Lecture. Human spermatozoa: fruits of creation, seeds of doubt. Reprod Fertil Dev 2004; 16: 655 7. Aitken RJ, Krausz CG. Oxidative stress, DNA damage and the Y chromosome. Reproduction 2001; 122: 497 8. Sakkas D, et al. Sperm nuclear DNA damage and altered chromatin structure: effect on fertilization and embryo development. Hum Reprod 1998; 13: 11 9. Duran EH, et al. Sperm DNA quality predicts intrauterine insemination outcome: a prospective cohort study. Hum Reprod 2002; 17: 3122 10. Morris ID, et al. The spectrum of DNA damage in human sperm assessed by single cell gel electrophoresis (comet assay) and its relationship to fertilization and embryo development. Hum Reprod 2002; 17: 990 11. Carrell DT, et al. Elevated sperm chromosome aneuploidy and apoptosis in patients with unexplained recurrent pregnancy loss. Obstet Gynecol 2003; 101: 1229 12. Loft S, et al. Oxidative DNA damage in human sperm influences time to pregnancy. Hum Reprod 2003; 18: 1265 13. Saleh RA, et al. Negative effects of increased sperm DNA damage in relation to seminal oxidative stress in men with idiopathic and male factor infertility. Fertil Steril 2003; 79 (Suppl 3): 1597 14. Bungum M, et al. The predictive value of sperm chromatin structure assay (SCSA) parameters for the outcome of intrauterine insemination, IVF and ICSI. Hum Reprod 2004; 19: 1401 15. Virro MR, Larson-Cook KL, Evenson DP. Sperm chromatin structure assay parameters are related to fertilization, blastocyst development, and ongoing

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31. Aitken RJ, Harkiss D, Buckingham DW. Analysis of lipid peroxidation mechanisms in human spermatozoa. Mol Reprod Dev 1993; 35: 302 32. Gomez E, Irvine DS, Aitken RJ. Evaluation of a spectrophotometric assay for the measurement of malondialdehyde and 4-hydroxyalkenals in human spermatozoa: relationships with semen quality and sperm function. Int J Androl 1998; 21: 81 33. Aitken RJ, Clarkson JS, Fishel S. Generation of reactive oxygen species, lipid peroxidation and human sperm function. Biol Reprod 1989; 40: 183 34. Suleiman SA, et al. Lipid peroxidation and human sperm motility: protective role of vitamin E. J Androl 1996; 17: 530 35. Rhemrev JP, et al. Quantification of the nonenzymatic fast and slow TRAP in a postaddition assay in human seminal plasma and the antioxidant contributions of various seminal compounds. J Androl 2000; 21: 913 36. van Overveld FW, et al. Tyrosine as important contributor to the antioxidant capacity of seminal plasma. Chem Biol Interact 2000; 127: 151 37. Alvarez JG, et al. Spontaneous lipid peroxidation and production of hydrogen peroxide and superoxide in human spermatozoa. J Androl 1987; 8: 338 38. Aitken RJ, et al. Relative impact of oxidative stress on the functional competence and genomic integrity of human spermatozoa. Biol Reprod 1998; 59: 1037 39. Aitken RJ, Buckingham D, Harkiss D. Use of a xanthine oxidase oxidant generating system to investigate the cytotoxic effects of reactive oxygen species on human spermatozoa. J Reprod Fertil 1992; 97: 441 40. Storey BT, Alvarez JG, Thompson KA. Human sperm glutathione reductase activity in situ reveals limitation in the glutathione antioxidant defense system due to supply of NADPH. Mol Reprod Dev 1998; 49: 400 41. Vernet P, et al. In vitro expression of a mouse tissue specific glutathione-peroxidase-like protein lacking the selenocysteine can protect stably transfected mammalian cells against oxidative damage. Biochem Cell Biol 1996; 74: 125 42. Mennella MRF, Jones R. Properties of spermatozoal superoxide dismutase and lack of involvement of superoxides in metal-ion-catalysed lipid-peroxidation reactions in semen. Biochem J 1980; 191: 289 43. Jones R, Mann T, Sherins RJ. Adverse effects of peroxidized lipid on human spermatozoa. Proc R Soc Lond B 1978; 201: 413 44. Aitken RJ, Harkiss D, Buckingham D. Relationship between iron-catalysed lipid peroxidation potential

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and human sperm function. J Reprod Fertil 1993; 98: 257 Kodama H, et al. Increased deoxyribonucleic acid damage in the spermatozoa of infertile male patients. Fertil Steril 1997; 65: 519 Henkel R, et al. DNA fragmentation of spermatozoa and assisted reproduction technology. Reprod Biomed Online 2003; 7: 477 Aitken RJ, et al. Multiple forms of redox activity in populations of human spermatozoa. Mol Hum Reprod 2003; 9: 645 Aitken RJ, et al. Differential contribution of leucocytes and spermatozoa to the high levels of reactive oxygen species recorded in the ejaculates of oligozoospermic patients. J Reprod Fertil 1992; 94: 451 De Lamirande E, et al. Increased reactive oxygen species formation in semen of patients with spinal cord injury. Fertil Steril 1995; 63: 637 Hendin BN, et al. Varicocele is associated with elevated spermatozoal reactive oxygen species production and diminished seminal plasma antioxidant capacity. J Urol 1999; 161: 1831 Pasqualotto FF, et al. Oxidative stress in normospermic men undergoing infertility evaluation. J Androl 2001; 22: 316 Aitken RJ, Irvine DS, Wu FC. Prospective analysis of sperm–oocyte fusion and reactive oxygen species generation as criteria for the diagnosis of infertility. Am J Obstet Gynecol 1991; 164: 542 Zorn B, Vidmar G, Meden-Vrtovec H. Seminal reactive oxygen species as predictors of fertilization, embryo quality and pregnancy rates after conventional in vitro fertilization and intracytoplasmic sperm injection. Int J Androl 2003; 26: 279 Aitken RJ, et al. Reactive oxygen species generation by human spermatozoa is induced by exogenous NADPH and inhibited by the flavoprotein inhibitors diphenylene iodonium and quinacrine. Mol Reprod Dev 1997; 47: 468 Said TM, et al. Impact of sperm morphology on DNA damage caused by oxidative stress induced by beta nicotinamide adenine dinucleotide phosphate. Fertil Steril 2005; 83: 95 Baker MA, et al. Identification of cytochrome P450reductase as the enzyme responsible for NADPHdependent lucigenin and tetrazolium salt reduction in rat epididymal sperm preparations. Biol Reprod 2004; 71: 307 Afanasíev IB, Ostrachovich EA, Korkina LG. Lucigenin is a mediator of cytochrome C reduction but

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not of superoxide anion production. Arch Biochem Biophys 1999; 366: 267 Aitken RJ, Baker MA, O’Bryan MK. Shedding light on chemiluminescence: the application of chemiluminescence in diagnostic andrology. J Androl 2004; 25: 455 Aitken RJ, West K. Relationship between reactive oxygen species generation and leucocyte infiltration in fractions isolated from the human ejaculate on Percoll gradients. Int J Androl 1990; 13: 433 Aitken RJ, et al. Analysis of sperm movement in relation to the oxidative stress created by leukocytes in washed sperm preparations and seminal plasma. Hum Reprod 1995; 10: 2061 Aitken RJ, West K, Buckingham D. Leukocyte infiltration into the human ejaculate and its association with semen quality, oxidative stress and sperm function. J Androl 1994; 15: 343 Allamaneni SS, et al. Characterization of oxidative stress status by evaluation of reactive oxygen species levels in whole semen and isolated spermatozoa. Fertil Steril 2005; 83: 800 Williams AC, Ford WC. Relationship between reactive oxygen species production and lipid peroxidation in human sperm suspensions and their association with sperm function. Fertil Steril 2005; 83: 929 Aitken RJ, et al. On the use of paramagnetic beads and ferrofluids to assess and eliminate the leukocytic contribution to oxygen radical generation by human sperm suspensions. Am J Reprod Immunol 1996; 35: 541 Krausz C, et al. Stimulation of oxidant generation by human sperm suspensions using phorbol esters and formyl peptides: relationships with motility and fertilization in vitro. Fertil Steril 1994; 62: 599 Krausz C, et al. Analysis of the interaction between N-formylmethionyl-leucyl phenylalanine and human sperm suspensions, development of a technique for monitoring the contamination of human semen samples with leucocytes. Fertil Steril 1992; 57: 1317

67. Gomez E, et al. Development of an image analysis system to monitor the retention of residual cytoplasm by human spermatozoa: correlation with biochemical markers of the cytoplasmic space, oxidative stress and sperm function. J Androl 1996; 17: 276 68. Gil-Guzman E, et al. Differential production of reactive oxygen species by subsets of human spermatozoa at different stages of maturation. Hum Reprod 2001; 16: 1922 69. Ollero M, et al. Characterization of subsets of human spermatozoa at different stages of maturation: implications in the diagnosis and treatment of male infertility. Hum Reprod 2001; 16: 1912 70. Zini A, et al. Human sperm NADH and NADPH diaphorase cytochemistry: correlation with sperm motility. Urology 1998; 51: 464 71. Aitken RJ. A free radical theory of male infertility. Reprod Fertil Dev 1994; 6: 19 72. Banfi B, et al. A Ca(2+)-activated NADPH oxidase in testis, spleen, and lymph nodes. J Biol Chem 2001; 276: 37594 73. de Lamirande E, Gagnon C. Human sperm hyperactivation and capacitation as parts of an oxidative process. Free Radic Biol Med 1993; 14: 157 74. de Lamirande E, Gagnon C. Capacitation-associated production of superoxide anion by human spermatozoa. Free Radic Biol Med 1995; 18: 487 75. Aitken RJ, et al. A novel signal transduction cascade in capacitating human spermatozoa characterised by a redox-regulated, cAMP-mediated induction of tyrosine phosphorylation. J Cell Sci 1998; 111: 645 76. Aitken RJ, et al. Redox regulation of tyrosine phosphorylation in human spermatozoa and its role in the control of human sperm function. J Cell Sci 1995; 108: 2017 77. Gomez E, Aitken J. Impact of in vitro fertilization culture media on peroxidative damage to human spermatozoa. Fertil Steril 1992; 65: 880

18 How do we define male subfertility and what is the prevalence in the general population? T Igno Siebert, F Haynes van der Merwe, Thinus F Kruger, Willem Ombelet

INTRODUCTION

Much less has been published on the use of this criterion regarding in vivo fertility. In this chapter, we evaluate the classification systems for semen parameters after review of the literature published in English on semen parameters and in vivo fertility potential. We also use data from the literature to establish fertility/subfertility thresholds for semen parameters according to the WHO 1999 guidelines3–6. These thresholds should be of clinical value and useful when assessing male fertility potential for in vivo conditions, in order to identify those males with a significantly reduced chance of achieving success under these conditions.

Several semen parameters are used to discriminate the fertile male from the subfertile male. The most widely used parameters are sperm concentration, motility, progressive motility and sperm morphology. Of these parameters, sperm morphology is the single indicator most widely debated in the literature. A large number of classification systems have been used to describe the factors that constitute a morphologically normal/abnormal spermatozoon. The most widely accepted classification systems for sperm morphology are the World Health Organization (WHO) criteria of 1987 and 19921,2 and the Tygerberg strict criteria, now also used by the WHO since 19993–6. Although there is a positive correlation between normal semen parameters and male fertility potential, the threshold values for fertility/subfertility according to WHO criteria1,2 are of little clinical value in discriminating between the fertile and the subfertile male7–11. If these criteria were to be applied, a great number of fertile males (partners having had pregnancies shortly before, after or at the time of a spermiogram) would be classified as subfertile. The predictive values of sperm morphology using strict criteria in in vitro fertilization (IVF) and intrauterine insemination (IUI) have been reviewed recently and proved to be useful12,13.

WHO CRITERIA OF 1987 AND 1992 AND MALE FERTILITY POTENTIAL The semen analysis is used in clinical practice to assess male fertility potential. To be of clinical value, the methods used should be standardized, and threshold values for fertility/subfertility should be calculated for the different parameters used in the standard semen analysis. Because there are so many different methods for semen evaluation, it would be difficult to standardize the methods used in its analysis. This applies especially to the assessment of sperm 269

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morphology. The two classification systems most widely accepted are the WHO1,2 and the Tygerberg strict criteria3–6. Various methodological problems concerning sperm morphology have been identified. The variants among different methods of morphology assessment have been reported by Ombelet et al.14–16 and others17,18, and they recommend standardization of semen analysis methodologies. Some authors recommend that laboratories should adopt the accepted standards, such as those proposed by the WHO17,18. Another problem identified is the variation in intra- and interindividual and interlaboratory sperm morphology assessment18,19. This problem can be addressed by using the Tygerberg strict criteria, as Menkveld et al. showed that comparable and reliable results between and within observers could be obtained when using this method19. Franken et al. delivered dedicated work on continuous qualitycontrol programs for strict sperm morphology assessment, and demonstrated that consistent readings could be achieved; they hence stressed the need for global quality-control measurements in andrology laboratories20,21. Cooper et al.18 also urged the standardization of such quality-control programs and that quality control centers should reach agreement with each other. Previous WHO thresholds of 50% and 30% for sperm morphology were empirical values and not based on any clinical data. Several authors found these values to be of little or no clinical value7,9,10. These studies did, however, find a positive correlation between a high proportion of morphologically normal sperm and an increased likelihood of fertility and/or pregnancy. Other studies have confirmed this correlation22–25. Van Zyl et al.25 were the first to show a faster than linear decline in fertilization rate when the proportion of normal forms dropped to less than 4%. Eggert-Kruse et al.23 found a higher in vivo pregnancy rate for higher percentage normal forms at thresholds of 4, 7 and 14% using strict criteria for morphology assessment. Zinaman et

al.26 confirmed the value of sperm morphology (strict criteria) by demonstrating a definite decline in pregnancy rate in vivo when the normal morphology dropped below 8% and sperm concentration below 30 × 106/ml. In a study performed by Slama et al.27, measuring the association between time to pregnancy and semen parameters, it was found that the proportion of morphologically normal sperm influenced the time to pregnancy up to a threshold value of 19%. This value is somewhat higher than that calculated in other studies.

THE USE OF SEMEN PARAMETERS IN IVF AND IUI PROGRAMS The percentage of normal sperm morphology (strict criteria) has a positive predictive value in IVF and IUI programs. Normal sperm morphology thresholds produced positive predictive values for IVF success when using the 5% and 14% thresholds, respectively, with the overall fertilization rate and overall pregnancy rate significantly higher in the group with normal morphology ≥ 5% as compared with the < 5% group12. A metaanalysis of data from IUI programs showed a higher pregnancy rate per cycle in the group with normal sperm morphology ≥ 5%. In the group with normal sperm morphology < 5%, other semen parameters predicted IUI success13. In the IUI meta-analysis, motility28, total motile sperm count29 and concentration30 also played a role in some of the studies evaluated, while others31 stated that sperm morphology alone was enough to predict the prognosis. Because of the high cost of assisted reproduction, males with good or reasonable fertility potential under in vivo conditions should be identified on the basis of semen quality. Conversely, males with a poor fertility potential should be identified, and introduced to assisted reproduction programs.

HOW DO WE DEFINE MALE SUBFERTILITY?

FERTILITY/SUBFERTILITY THRESHOLDS FOR SPERM MORPHOLOGY USING TYGERBERG STRICT CRITERIA, SPERM CONCENTRATION AND SPERM MOTILITY/PROGRESSIVE MOTILITY In an effort to establish fertility/subfertility thresholds for the aforementioned parameters, we identified four articles in the published literature. It is our opinion that these articles constitute a representative sample of published studies of the predictive value of sperm morphology, sperm concentration and motility/progressive motility for in vivo fertility/subfertility. These articles compared the different semen parameters of a fertile and a subfertile group. They used either classification and regression tree (CART) analysis or receiver operating characteristic (ROC) curve analysis to estimate thresholds for the various semen parameters. The ROC curve was also used to assess the diagnostic accuracy of the different parameters

Table 18.1

271

and their ability to classify subjects into fertile and subfertile groups. Using ROC curve analysis, Ombelet et al.32 calculated the following thresholds: proportion normal morphology 10%, proportion normal motility 45% and normal sperm concentration 34 × 106/ml. Sperm morphology was shown to be the parameter with the highest prediction power (area under the curve (AUC) 78%). Much lower thresholds were calculated using the 10th centile of the fertile population, these thresholds being 5% for normal morphology, 28% for motility and 14.3 × 106/ml for sperm concentration (Tables 18.1 and 18.2)32. Günalp et al.33 also calculated thresholds using ROC curve analysis. These thresholds were: proportion normal morphology 10%, proportion normal motility 52%, proportion progressive motility 42% and sperm concentration 34 × 106/ml. The two parameters that performed best were progressive motility (AUC 70.7%) and

Thresholds: fertile vs. subfertile populations studied Normal morphology (%)

Motility (%)

Guzick et al.35 (2001)

—

Menkveld et al.34 (2001)

—

Günalp et al.33 (2001)

Ombelet et al.32 (1997)

—

Authors

Progressive motility (%)

Concentration × 106/ml) (× 13.5

Table 18.2 Possible lower thresholds for the general population to distinguish between subfertile and fertile men based on the assumed incidences of subfertile males in their populations Normal morphology (%)

Motility (%)

Progressive motility (%)

Menkveld et al.34 (2001)

—

Günalp et al.33 (2001)

—

14.3

Authors

Ombelet et al.

(1997)

Concentration × 106/ml) (×

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morphology (AUC 69.7%). Assuming 50% prevalence of subfertility in the population, the authors used the positive predictive value as an indicator to calculate a lower threshold for each parameter. Values of 5% for proportion normal morphology, 30% for proportion normal motility, 14% for proportion progressive motility and 9 × 106/ml for sperm concentration were calculated (Tables 18.1 and 18.2)33. In the most recent article of the four, Menkveld et al.34 found much lower thresholds than the others. Using ROC curve analysis, the following thresholds were calculated: 4% for normal morphology and 45% for normal motility. Again, morphology showed good predictive value with an AUC of 78.2%. Although a threshold for sperm concentration was not calculated (a sperm concentration less than 20 × 106/ml was used as inclusion criterion), the authors proposed that the cut-off value of 20 × 106/ml could be used with confidence, based on the resultant lower 10th centile of the fertile population. Adjusted cut-off points calculated on the assumption of 50% prevalence of male subfertility were as follows: 3% for proportion normal morphology and 20% for proportion normal motility (Tables 18.1 and 18.2)34. In the fourth article by Guzick et al.35, the authors used CART analysis and calculated two thresholds for each semen parameter which allowed designation into three groups, namely normal (fertile), borderline and abnormal (subfertile). The normal (fertile) group had values greater than 12% for morphology, greater than 63% for motility and higher than 48 × 106/ml for sperm concentration. The abnormal (subfertile) group had values lower than 9% for morphology, lower than 32% for motility and lower than 13.5 × 106/ml for sperm concentration. In these four articles, the predictive power of the different parameters was calculated as the AUC, using the ROC curve. The AUC for sperm morphology ranged from 66 to 78.2%, confirming the high predictive power of this parameter. In fact, it had the best performance among the

different semen parameters in two articles32,35. The thresholds calculated in these two articles were 10% and 9%, respectively, while Günalp et al.33 calculated a threshold of 12% using sensitivity and specificity to analyze their data, and the fourth study calculated a 4% predictive cut-off value. Although sensitivity and specificity for the values are relatively high, the positive predictive values are not. This will therefore result in classifying fertile males as subfertile, probably leading to a degree of anxiety as well as unnecessary and costly infertility treatment. A second and much lower threshold was calculated in three of the four articles. Ombelet et al.32 calculated this much lower threshold by using the 10th centile of the fertile population, while Günalp et al.33 screened the population with the positive predictive value as indicator, and Menkveld et al.34 assumed a 50% prevalence of subfertility in their study population. The lower threshold ranged from 3 to 5% (Table 18.2). These lower thresholds have a much higher positive predictive value than the higher thresholds, with a negative predictive value not much lower. We suggest that the lower threshold should be used to identify males with the lowest potential for a pregnancy under in vivo conditions. Values above the lower threshold should be regarded as normal. These findings are in keeping with previous publications by Coetzee et al.12 (IVF data) and Van Waart et al.13 (IUI data), which reported a significantly lower chance of successful pregnancy in males with normal morphology below their calculated thresholds. The higher threshold values for percentage motile sperm as calculated in the four articles (using ROC curve or CART analysis) ranged from 32 to 52%, while the lower threshold values ranged from 20 to 30%. Motility also had a high predictive power, with an AUC of between 59 and 79.1%. Günalp et al.33 calculated thresholds for progressive motility: a higher threshold of 42%, using the ROC curve, and a lower threshold of 14%, with the positive predictive value as indicator. In this study, progressive motility

HOW DO WE DEFINE MALE SUBFERTILITY?

proved to be a marginally better predictor of subfertility than sperm morphology, with AUC values of 70.7 and 69.7%, respectively33. Montanaro Gauci et al.28 found percentage motility to be a significant predictor of IUI outcome. The pregnancy rate was almost three times higher in the group with motility > 50% as compared with the group with motility < 50%. The higher threshold values for sperm concentrations calculated by Ombelet et al.32, Günalp et al.33 and Guzick et al.35 ranged from 13.5 to 34 × 106/ml, while the lower threshold values ranged from 9 to 14.3 × 106/ml. An AUC value of between 55.5 and 69.4% served as confirmation of the predictive power of this parameter. Although Menkveld et al.34 did not calculate a threshold value for sperm concentration (because values of less than 20 × 106/ml served as inclusion criteria in their study), they suggested a threshold value of 20 × 106/ml to be used with confidence, because it did not influence the results from their fertile population. The clinical value of motility and sperm concentration serves as confirmation of findings reported in numerous other publications7,8,11,22–24. Although the various parameters had good predictive power, independent of each other, the clinical value of semen analysis was increased when the parameters were used in combination. Ombelet et al.32 found that differences between the fertile and subfertile populations only became significant when two or all three semen parameters were combined. Bartoov et al.36 concluded that fertility potential is dependent on a combination of different semen characteristics. EggertKruse et al.23 found a significant correlation between the three parameters reviewed in their study. Although the different semen parameters demonstrate good individual predictive power, the clinical value of the semen analysis increases when the parameters are used in combination. We therefore suggest that no parameter should be used in isolation when assessing male fertility potential. The lower thresholds as discussed in this chapter have a much higher positive predictive value and a

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high negative predictive value. Therefore, we suggest that these lower thresholds should be used in identifying the subfertile male. As suggested by the WHO in 1999, each group should develop their own thresholds, based on the population they are working in. It seems as if the sperm morphology threshold of 0–4% normal forms indicates a higher risk group for subfertility, and fits the IVF and IUI data calculated previously12,13. The four articles discussed above32–35 showed the same trends, and can serve as guidelines to distinguish fertile from subfertile males. As far as concentration and motility are concerned, the thresholds are not clear, but a concentration lower than 106/ml and a motility lower than 30% seem to fit the general data32–35. However, more, preferably multicenter, studies are needed to set definitive thresholds.

SEMEN PROFILE OF THE GENERAL POPULATION: PARTNERS OF WOMEN WITH CHRONIC ANOVULATION In general, there is quite a poor level of understanding and evidence regarding the semen analysis profile of the general population. Many male populations have been proposed to mirror the general population in terms of semen analysis. Using donors in a semen-donation program for normality is certainly not the best option, since this population is positively biased for fertility. Army recruits are biased by age. Husbands of tubal-factor patients can be biased by a positive history of infection (tubal factor due to pelvic infection) or a good fertility history (women with tubal sterilization). Therefore, we believe that possibly the best reference group for studying the semen profile in a general population includes partners of women who have been diagnosed with chronic anovulation/PCOS (polycystic ovarian syndrome) (maximum of three menstrual periods per year). We would thus like to propose employing the lower thresholds to indicate patients with subfertility, and, by using the cohort of

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anovulatory women, we obtain a reflection of the semen profile in a general population. Two different studies, one retrospective and one prospective, evaluating the semen analysis of partners of women presenting with anovulation were selected.

Retrospective study of partners of women presenting with chronic anovulation (> 35 days) at Tygerberg Fertility Clinic Included in this study were all male partners of patients diagnosed as anovulatory at the Tygerberg Fertility Clinic. Methods used to examine the semen were according to WHO guidelines6, and for sperm morphology Tygerberg strict criteria were used3,4,6. The laboratory personnel initially evaluated all slides, and each slide was then evaluated by one observer (TFK) according to strict criteria. Sixty-two samples were eventually selected and included in the study (Table 18.3).

Prospective study of partners of women presenting with PCOS at Tygerberg Fertility Clinic Tygerberg Fertility Clinic conducted a study in patients with PCOS. The patients were diagnosed with PCOS according to the recent Rotterdam consensus statement37. The aim of this study was to establish factors influencing ovulation induction in this group. The semen of the partners of all these women was examined. Methods used to examine the semen were according to WHO guidelines6, and for sperm morphology Tygerberg strict criteria were used3,4,6. The laboratory personnel initially evaluated all slides, and all P-pattern morphology slides were re-evaluated by one observer (TFK) (Table 18.4). The thresholds used for subfertility were those suggested by Van der Merwe et al.38 in their recent review: 0–4% normal forms, < 30% motility, < 106/ml, outlined in the first section of this chapter.

Table 18.3 Retrospective study of partners of women presenting with chronic anovulation (> 35 days) at Tygerberg Fertility Clinic (< 106/ml cut-off) Patients n

Normozoospermia

46.7

Single-parameter defect azoospermia oligozoospermia (O) asthenozoospermia (A) teratozoospermia (T) polyzoospermia (P) immunological factor (I)

3 3 — 16 2 1

4.8 4.8 0 25.8 3.2 1.6

Double-parameter defect OA OT AT TP TI

— 4 — 1 1

0 6.5 0 1.6 1.6

3.2

Sperm abnormality

Triple-parameter defect OAT

Threshold values used: concentration < 106/ml, motility < 30%, morphology < 4% normal forms

DISCUSSION In the two studies (Table 18.3, retrospective; Table 18.4, prospective) ± 50% of patients had a normal semen analysis. The most common single abnormality was that of teratozoospermia (25.8% retrospective, 27.8% prospective). Azoospermia occurred in 1.4–4.8% of patients, with tripleparameter defects found in only 1.4–3.2% of cases (Tables 18.3 and 18.4). The thresholds as calculated above were used in a group of anovulatory women. These thresholds reflect the prevalence of male factor infertility in the general population. It is interesting to note that in both the retrospective and prospective studies, the prevalence of teratozoospermia (< 4%

HOW DO WE DEFINE MALE SUBFERTILITY?

Table 18.4 Prospective study of partners of women presenting with polycystic ovarian syndrome (PCOS) at Tygerberg Fertility Clinic (< 106/ml cut-off) Patients n

Normozoospermia

56.9

Single-parameter defect azoospermia oligozoospermia (O) asthenozoospermia (A) teratozoospermia (T) polyzoospermia (P) immunological factor (I)

1 1 — 20 3 —

1.4 1.4 0 27.8 4.2 0

Double-parameter defect OA OT AT TP TI OP

— 1 — 3 1 —

0 1.4 0 4.2 1.4 0

1.4

Sperm abnormality

Triple-parameter defect OAT

normal morphology) was 25.8–27.8%, making it the most common defect in this group. About 50% of all male patients had normal semen parameters in these two studies using the suggested thresholds as calculated based on the four articles discussed32–35,38. It is important to note that in PCOS patients the clinician needs to take into consideration that not only anovulation, but also, in up to 50% of these patients, the male factor needs attention, to assist in achieving a successful outcome in these couples. These lower thresholds are not absolute, but provide a continuum guiding the clinician to respond to the semen analysis. The golden rule is to repeat a semen analysis 4 weeks after the first (abnormal) evaluation to ensure that the correct approach will be followed. If the result is again abnormal, a thorough physical examination

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should be performed and the necessary treatment offered. In the case of PCOS, the female factor (anovulation) should obviously be corrected, starting, as first-line approach, with weight loss in women with a body mass index > 25. Although 50% of these patients had a male factor according to the definition used, it is also important to note that only ± 5% of these factors were serious (azoospermia and the triple-parameter defects), with 7–9.7% with a double defect. To our knowledge, this is the first attempt to use the specific suggested lower thresholds to define prevalence of the subfertile male in the general population by using an anovulatory group of women. These thresholds will guide the clinician towards a more directive management where indicated.

REFERENCES 1. World Health Organization. WHO Laboratory Manual for the Examination of Human Semen and Sperm–Cervical Mucus Interaction, 2nd edn. Cambridge: Cambridge University Press, 1987 2. World Health Organization. WHO Laboratory Manual for the Examination of Human Semen and Sperm–Cervical Mucus Interaction, 3rd edn. Cambridge: Cambridge University Press, 1992 3. Kruger TF, et al. Predictive value of abnormal sperm morphology in in vitro fertilization. Fertil Steril 1988; 49: 112 4. Kruger TF, et al. Sperm morphologic features as a prognostic factor in in vitro fertilization. Fertil Steril 1986; 46: 1118 5. Menkveld R, et al. The evaluation of morphological characteristics of human spermatozoa according to stricter criteria. Hum Reprod 1990; 5: 586 6. World Health Organization. WHO Laboratory Manual for the Examination of Human Semen and Sperm–Cervical Mucus Interaction, 4th edn. Cambridge: Cambridge University Press, 1999 7. Barratt CL, et al. Clinical value of sperm morphology for in-vivo fertility: comparison between World Health Organization criteria of 1987 and 1992. Hum Reprod 1995; 10: 587 8. Ayala C, Steinberger E, Smith DP. The influence of semen analysis parameters on the fertility potential of infertile couples. J Androl 1996; 17: 718

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9. Blonde JP, et al. Relation between semen quality and fertility: a population-based study of 430 first-pregnancy planners. Lancet 1998; 352: 1172 10. Chia SE, Tay SK, Lim ST. What constitutes a normal seminal analysis? Semen parameters of 243 fertile men. Hum Reprod 1998; 13: 3394 11. C

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