Infertility in the Male [5 ed.] 1108838057, 9781108838054

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Infertility in the Male Fifth Edition

Published online by Cambridge University Press

Published online by Cambridge University Press

Infertility in the Male Fifth Edition

Edited by

Larry I. Lipshultz Baylor College of Medicine

Stuart S. Howards University of Virginia

Craig S. Niederberger University of Illinois at Chicago

Dolores J. Lamb Weill Cornell Medical College

Published online by Cambridge University Press

University Printing House, Cambridge CB2 8BS, United Kingdom One Liberty Plaza, 20th Floor, New York, NY 10006, USA 477 Williamstown Road, Port Melbourne, VIC 3207, Australia 314–321, 3rd Floor, Plot 3, Splendor Forum, Jasola District Centre, New Delhi – 110025, India 103 Penang Road, #05–06/07, Visioncrest Commercial, Singapore 238467 Cambridge University Press is part of the University of Cambridge. It furthers the University’s mission by disseminating knowledge in the pursuit of education, learning, and research at the highest international levels of excellence. www.cambridge.org Information on this title: www.cambridge.org/9781108838054 DOI: 10.1017/9781108937054 © Cambridge University Press 2023 This publication is in copyright. Subject to statutory exception and to the provisions of relevant collective licensing agreements, no reproduction of any part may take place without the written permission of Cambridge University Press. First published Printed in Singapore by Markono Print Media Pte Ltd A catalogue record for this publication is available from the British Library. Library of Congress Cataloging-in-Publication Data Names: Lipshultz, Larry I., 1942- editor. | Howards, Stuart S., 1937- editor. | Niederberger, Craig S., editor. | Lamb, Dolores, editor. Title: Infertility in the male / edited by Larry Lipshultz, Stuart Howards, Craig Niederberger, Dolores Lamb. Description: 5th edition. | Cambridge, United Kingdom; New York, NY: Cambridge University Press, 2022. | Includes bibliographical references and index. Identifiers: LCCN 2021057951 (print) | LCCN 2021057952 (ebook) | ISBN 9781108838054 (hardback) | ISBN 9781108937054 (epub) Subjects: MESH: Infertility, Male Classification: LCC RC889 (print) | LCC RC889 (ebook) | NLM WJ 709 | DDC 616.6/92–dc23/eng/20211227 LC record available at https://lccn.loc.gov/2021057951 LC ebook record available at https://lccn.loc.gov/2021057952 ISBN 978-1-108-83805-4 Hardback Cambridge University Press has no responsibility for the persistence or accuracy of URLs for external or third-party internet websites referred to in this publication and does not guarantee that any content on such websites is, or will remain, accurate or appropriate.

........................................................................... Every effort has been made in preparing this book to provide accurate and up-to-date information that is in accord with accepted standards and practice at the time of publication. Although case histories are drawn from actual cases, every effort has been made to disguise the identities of the individuals involved. Nevertheless, the authors, editors, and publishers can make no warranties that the information contained herein is totally free from error, not least because clinical standards are constantly changing through research and regulation. The authors, editors, and publishers therefore disclaim all liability for direct or consequential damages resulting from the use of material contained in this book. Readers are strongly advised to pay careful attention to information provided by the manufacturer of any drugs or equipment that they plan to use.

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Contents List of Contributors Foreword xi Abbreviations xiii

0.

vii

9.

Introduction 1 Craig S. Niederberger, Dolores J. Lamb, Larry I. Lipshultz, and Stuart S. Howards

10. Imaging the Male Reproductive System Roger K. Khouri Jr and Tolulope Bakare

Section 1: Scientific Foundations of Male Infertility 1.

2.

Anatomy and Embryology of the Male Reproductive Tract and Gonadal Development, the Epididymis, and Accessory Sex Organs 5 Danielle Velez and Craig S. Niederberger Cellular Architecture and Function of the Testis 17 Siwen Wu, Lingling Wang, and C. Yan Cheng

3.

Maturation and Function of Sperm 39 Caroline Kang, Nahid Punjani, and Dolores J. Lamb

4.

The Male Reproductive Endocrine System Ettore Caroppo

5.

Erection, Emission, and Ejaculation 77 Ramy Abou Ghayda and Martin N. Kathrins

6.

Genomics, Epigenetics, and Male Reproduction 94 Millissia Ben Maamar and Michael Skinner

62

146

Evaluation of the Infertile Male’s Partner Sahar Wertheimer, Jessica L. Chan, and Margareta D. Pisarska

165

11. Effects of Environmental Chemicals on Male Reproduction 182 Rebecca Z. Sokol 12. Endocrine Causes of Male Infertility – Diagnosis and Treatment 197 Fiona Yuen, Ronald S. Swerdloff, and Christina C. L. Wang 13. Spermatogenesis – Diagnosis of Normal and Abnormal States 218 Mahmoud Mima and Richard A. Schoor 14. Inheritance and Male Fertility 237 Cigdem Tanrikut and Robert D. Oates 15. The Varicocele – Approaches to Diagnosis and Management 253 Sarah C. Krzastek, Ryan P. Smith, and Stuart S. Howards 16. Infection, Inflammation, and Immunological Causes of Male Infertility 277 Joshua A. Halpern, Caleb A. Cooper, Sanjay S. Kasturi, and Robert E. Brannigan

Section 2: Clinical Evaluation of the Infertile Male 7.

Infertility as a Metric of Men’s Health 107 Jeremy T. Choy and Michael L. Eisenberg

8.

Office Evaluation of the Subfertile Male Gabriella Avellino and Mark Sigman

113

Section 3: Laboratory Diagnosis of Male Infertility 17. Laboratory Evaluation of the Infertile Male J. Scott Gabrielsen

329

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Contents

26. Male Contraception 477 Darius Devlin, Martin M. Matzuk, Kelly Walker, and Jay I. Sandlow

18. Advanced Diagnostic Approaches to Male Infertility 348 Dolores J. Lamb 19. Evaluating Defects in Sperm Function Christopher J. De Jonge

363

20. Cryopreservation of Sperm – History and Current Practice 381 Cappy M. Rothman and Mitchel C. Schiewe

Section 4: Treatment of Male Infertility 21. Medical Treatment of Male Infertility 399 Craig S. Niederberger, Rodrigo Lessi Pagani, and Samuel J. Ohlander 22. Surgery to Improve Sperm Delivery 413 Saneal Rajanahally, Larry I. Lipshultz, Alexander W. Pastuszak, Danielle Velez, and Craig S. Niederberger 23. Sperm Retrieval Surgery Peter N. Schlegel

437

24. The Use of Sperm in Medically Assisted Reproduction 446 Susan Talamini and Gail S. Prins

27. Future Directions in Male Infertility Premal Patel and Ranjith Ramasamy

Section 5: Health Care Systems and Culture 28. Mental Health and Male Reproduction William D. Petok 29. Legal Issues and Male Reproduction Heather E. Ross

31. Global and Cultural Aspects of Male Reproductive Care 534 William J. Huang

Index

541

The videos can be found in the resources tab at www.cambridge.org/9781108838054

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503 517

30. Male Reproduction in the Transgender Patient 525 Brooke A. Harnisch and Stanton C. Honig

25. Male Oncofertility – Considerations for Fertility Preservation and Restoration 461 Darshan P. Patel, Alexander W. Pastuszak, and James M. Hotaling

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495

Contributors

Gabriella Avellino MD Brown Alpert Medical School. RI, USA Tolulope Bakare MD University of Texas Southwestern Medical Center, Department of Urology, Dallas, TX, USA Robert E. Brannigan MD Northwestern University Feinberg School of Medicine, Chicago, IL, USA Ettore Caroppo MD ASL Bari, U.O. Fisiopatologia della Riproduzione Umana e PMA, PTA “F Jaia,” Conversano, Bari, Italy Jessica L. Chan MD Division of Reproductive Endocrinology and Infertility, Center for Fertility and Reproductive Medicine, Division of REI, Department of Ob/Gyn, Cedars-Sinai Medical Center, Los Angeles, CA, USA C. Yan Cheng MD PhD The Mary M. Wohlford Laboratory for Male Contraceptive Research, Center for Biomedical Research, Population Council, New York, NY, USA Jeremy T. Choy MD Division of Endocrinology, Gerontology, and Metabolism, Stanford University School of Medicine, Stanford, CA, USA Caleb A. Cooper MD University of Chicago, Chicago, IL, USA Christopher J. De Jonge PhD HCLD (ABB) M Health Fairview, University of Minnesota, Minneapolis, MN, USA Darius J. Devlin, Ph.D. Regulatory Scientist at Biopharma Global

Michael L. Eisenberg MD Department of Urology, Stanford University School of Medicine, Stanford, CA, USA J. Scott Gabrielsen MD PhD Department of Urology, University of Rochester, Rochester, NY, USA Ramy Abou Ghayda MD MPH Division of Urology, Brigham and Women’s Hospital, Boston, MA, USA Joshua A. Halpern MD MS Northwestern University Feinberg School of Medicine, Chicago, IL, USA Brooke A. Harnisch MD Division of Urology, UCONN Health, Farmington, CT, USA Stanton C. Honig MD Department of Urology, Yale School of Medicine, New Haven, CT, USA James M. Hotaling MD, MS, FECSM Division of Urology, Department of Surgery, University of Utah Health, Salt Lake City, UT, USA Stuart S. Howards MD University of Virginia, Department of Urology, Charlottesville, VA, USA William J. Huang MD, PhD Department of Urology, Taipei Veterans General Hospital, School of Medicine, College of Medicine, National Yang Ming Chiao Tung University, Taipei, Taiwan Caroline Kang MD PhD James Buchanan Brady Foundation Institute of Urology, Weill Cornell Medical College, New York, NY, USA

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List of Contributors

Sanjay S. Kasturi MD New Jersey Urology, Vineland, NJ, USA Martin N. Kathrins MD Division of Urology, Brigham and Women’s Hospital, Boston, MA, USA Roger K. Khouri Jr MD University of Texas Southwestern Medical Center, Department of Urology, Dallas, TX, USA Sarah C. Krzastek MD Virginia Commonwealth University, Division of Urology, & McGuire VA Medical Center, Division of Urology, Richmond, VA, USA; University of Virginia, Department of Urology; Charlottesville, VA, USA Dolores J. Lamb HCLD PhD Brady Foundation Department of Urology, Center for Reproductive Genomics, Center for Reproductive Medicine, Englander Institute for Precision Medicine, Weill Cornell Medical College, New York, NY, USA Larry I. Lipshultz MD Scott Department of Urology, Baylor College of Medicine, Houston, TX, USA Millissia Ben Maamar PhD Center for Reproductive Biology, School of Biological Sciences, Washington State University, Pullman, WA, USA Martin M. Matzuk MD Department of Pathology & Immunology and Center for Drug Discovery, Texas, USA Mahmoud Mima MD University of Illinois at Chicago, Department of Urology, Chicago, IL, USA Craig S. Niederberger MD FACS Department of Urology, UIC College of Medicine, & Department of Bioengineering, UIC College of Engineering, University of Illinois at Chicago, Chicago, IL, USA

Rodrigo Lessi Pagani MD Department of Urology, University of Illinois at Chicago College of Medicine, Chicago, IL, USA Alexander W. Pastuszak MD PhD Division of Urology, Department of Surgery, University of Utah Health, Salt Lake City, UT, USA Darshan P. Patel MD Division of Urology, Department of Surgery, University of Utah Health, Salt Lake City, UT, USA Premal Patel MD FRCSC Section of Urology, Department of Surgery, University of Manitoba, Winnipeg, MB, Canada William D. Petok PhD Department of Obstetrics and Gynecology, Sidney Kimmel Medical College, Thomas Jefferson University, Philadelphia, PA, USA Margareta D. Pisarska MD Division of Reproductive Endocrinology and Infertility, Center for Fertility and Reproductive Medicine, Division of REI, Department of Ob/Gyn, Cedars-Sinai Medical Center, Los Angeles, CA, USA Gail S. Prins PhD Department of Urology, University of Illinois at Chicago, Chicago, IL, USA Nahid Punjani MD MPH Englander Institute for Precision Medicine, Weill Cornell Medical College, New York, NY, USA Saneal Rajanahally MD UGA, Stockbridge, Spivey Station, Sandy Springs, & Atlanta GA, USA Ranjith Ramasamy MD FACS Department of Urology, University of Miami Miller School of Medicine, Miami, FL, USA

Robert D. Oates MD FACS Boston University School of Medicine, Department of Urology, Boston Medical Center, Boston, MA, USA

Heather E. Ross Esq Ross & Zuckerman, LLP, Northbrook, IL, USA

Samuel J. Ohlander MD Department of Urology, University of Illinois at Chicago College of Medicine, Chicago, IL, USA

Cappy M. Rothman MD Center for Male Reproductive Medicine, Century City, CA, USA

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List of Contributors

Jay I. Sandlow MD Professor and Chair Director, Male Infertility and Andrology Fellowship Department of Urology Medical College of Wisconsin Milwaukee, USA Mitchel C. Schiewe MS PhD Ovation Fertility, Newport Beach, CA, USA Peter N. Schlegel MD Department of Urology, Weill Cornell Medicine, New York, NY, USA Richard A Schoor MD FACS Private Practice Urology, Smithtown, NY, USA Mark Sigman MD Alpert Medical School of Brown University, Providence, Rhode Island, USA Cigdem Tanrikut MD FACS Shady Grove Fertility, Department of Urology, Georgetown University School of Medicine, Washington, DC, USA Michael Skinner PhD Center for Reproductive Biology, School of Biological Sciences, Washington State University, Pullman, WA, USA

Ronald S. Swerdloff MD Division of Endocrinology, Department of Medicine, The Lundquist Institute and Harbor-UCLA Medical Center, Torrance, CA, USA Susan Talamini MD Department of Urology, University of Illinois at Chicago, Chicago, IL, USA Danielle Velez MD Department of Urology, University of Illinois at Chicago, Chicago, IL, USA Kelly Walker, MD, MBA Medical Director Posterity Health Christina C. L. Wang MD Division of Endocrinology, Department of Medicine, The Lundquist Institute and Harbor-UCLA Medical Center, Torrance, CA, USA Lingling Wang BSc MSc Center for Biomedical Research, Population Council, New York, NY, USA Sahar Wertheimer MD Southern California Reproductive Center, CA, USA

Ryan P. Smith MD University of Virginia, Department of Urology, Charlottesville, VA, USA

Siwen Wu MD Center for Biomedical Research, Population Council, New York, NY, USA

Rebecca Z. Sokol MD, MPH Medicine and Obstetrics and Gynecology, Keck School of Medicine, University of Southern California, Los Angeles, CA, USA

Fiona Yuen MD Division of Endocrinology, Department of Medicine, The Lundquist Institute and Harbor-UCLA Medical Center, Torrance, CA, USA

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Foreword

The fifth edition of Infertility in the Male continues to be the gold standard in the field of infertility urology. The editors represent three generations of pioneers and leaders in the field. Within the text, each has brought their own editorial skills and writing acumen. This classic is up-todate, as in the past. This is a rapidly changing field stimulated by the introduction to society of in vitro fertilization (IVF), intracytoplasmic sperm injection (ICSI), and microsurgical epididymal sperm aspiration (MESA). Forty years ago, infertility in the male was a sleepy, not very interesting pursuit. Today it is exploding with new information, cures, and insights. As in many fields, genetics has had a profound effect. Surgical procedures, as well as basic science research, has changed hyperbolically. Assembled authors is the who’s who in the field of urology. It includes basic scientists, translational scientists, and super clinicians. The 31 chapters cover all imaginable topics. The book serves as a manual for the novice and a reference source for the experienced practitioner, as well as a well-organized convenient source of information when information desired on a single topic is wanted. The opening shot over the bow, by the editors, entitled “why do we care for the male” gives an overview philosophical approach, as well as a historical perspective for treating male infertility. The text is interestingly divided into anatomy, both microscopic and gross, physiology, diagnosis and workup, including the female, and therapeutic modalities. Highlights include genomics and epigenomics of male reproduction, the environment and male infertility, and

cryopreservation of sperm, including in prepubescent males. Practical contributions include surgical sperm extraction, oncofertility, and contraception. Examples of how this text has grown and kept up with the times include chapters on inheritance of male infertility, advanced diagnostic approaches to male infertility, and future directions. The book is beautifully illustrated and there is a uniformity in the style of writing that makes it easy to read and comprehend the content. I quote from the fourth edition’s Foreword: “This fourth edition of Infertility in the Male certainly disproves the call to arms of the reproductive medicine community: when, in 1992, ICSI (intracytoplasmic sperm injection) appeared in the armamentarium of the infertility physicians it was claimed that urologists no longer had a role in the management of infertile men, except for obtaining sperm. This concept is certainly refuted and defeated by this exquisite revision of a book whose first edition was published in 1983.”

The growth in each edition is immeasurable and certainly this is true of the fifth edition of Infertility in the Male. In summary, the list of contributors and editors are those who pioneered the field, illustrating the dedication and prescience in treating the infertile couple. It is clear male factor infertility has come into its own as a serious discipline. Alan H. DeCherney MD

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https://doi.org/10.1017/9781108937054.001 Published online by Cambridge University Press

Abbreviations

3β-HSD 11βMNTDC 17OHD AATB ABA aCGH ACOG ActRII AFC AGD AGI AI AID AIDS AIS AJ AMA AMD AMH AO aPKC AR AR ARC ARIC ART ASA ASCO ASD ASRM ATP AUA AZF AZFa AZFb AZFc AZT BBT BCG BEB BMI BN BPA

3β-hydroxysteroid dehydrogenase 11β-methyl-nortestosteronedodecylcarbonate 17α-hydroxylase deficiency American Association of Tissue Banks American Bar Association array comparative genomic hybridization American College of Obstetricians and Gynecologists activin receptor type II antral follicle count abnormal anogenital distance anogenital index artificial intelligence artificial insemination with donor semen acquired immune deficiency syndrome androgen insensitivity syndrome adherens junction advanced maternal age adjusted mean difference anti-Müllerian hormone acridine orange atypical protein kinase C androgen receptor acrosome reaction arcuate nucleus acrosome reaction to ionophore challenge assisted reproductive technology antisperm antibodies American Society of Clinical Oncology anoscrotal distance American Society for Reproductive Medicine adenosine triphosphate American Urological Association azoospermia factor azoospermia factor a azoospermia factor b azoospermia factor c zidovudine basal body temperature bacille Calmette–Guérin blood–epididymis barrier body mass index Brown Norway bisphenol A

BPH BRDT BrdU BTB cAMP Cas9 CASA CatSper CBAVD CBP CBRC CBS CDC cDNA CDUS CF CFTR cGMP CI CL CLIA CMS CMV CNS CNV CoQ10 COSMIC COX COX-1 COX-2 CP CPA CpG CPPS Crb3 CREB CRISPR CT CUA CUAVD DAPI DAZ

benign prostate hyperplasia bromodomain testis-associated bromodeoxyuridine blood–testis barrier cyclic adenosine monophosphate CRISPR-associated protein 9 computer-assisted semen analysis cation channels of sperm congenital bilateral absence of the vas deferens chronic bacterial prostatitis cross-border reproductive care Cryo Bio System Centers for Disease Control and Prevention complementary DNA color Doppler ultrasound cystic fibrosis cystic fibrosis transmembrane conductance regulator cyclic guanosine monophosphate confidence interval chemiluminescence Clinical Laboratory Improvement Amendment Centers for Medicare and Medicaid Services cytomegalovirus central nervous system copy number variation coenzyme Q10 Catalogue of Somatic Mutations in Cancer cyclooxygenase cyclooxygenase 1 cyclooxygenase 2 chronic prostatitis cyproterone acetate cytosine phosphate guanine chronic pelvic pain syndrome Crumbs homolog-3 cAMP response element binding protein clustered regularly interspaced short palindromic repeats computed tomography Canadian Urological Association congenital unilateral absence of the vas deferens 40 ,6-diamidino-2-phenylindole Deleted in Azoospermia

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List of Abbreviations

DBCP DBD DBD-FISH DC DDT DEHP DES DFI DFS DHEA DHEAS DHT DI Dlg1 DMAU DMSO DNMT DOR dpp DSB DSD dsDNA DSM-5 dUTP Dvl3 EAU EB EBV ED EDC EDO EEJ EGR1 ELISA EOP EPA EPPIN EPS ER ERKO ES ESHRE ESR ESURSPIQG EV EST FDA FHA FIGO FISH FLCIVF Fmi

1,2-dibromo-3-chloropropane DNA-binding domain DNA breakage detection-fluorescence in situ hybridization dendritic cell dichlorodiphenyltrichloroethane di-ethyl-hexyl phthalate diethylstilbestrol DNA fragmentation index dysplasia of the fibrous sheath dehydroepiandrosterone dehydroepiandrosterone sulfate dihydrotestosterone donor insemination discs large 1 dimethandrolone undecanoate dimethyl sulfoxide DNA methyl transferase diminished ovarian reserve days postpartum DNA strand break disorder of sex development double-stranded DNA Diagnostic and Statistical Manual of Mental Disorders, fifth edition deoxyuridine triphosphate Disheveled 3 European Association of Urology elementary body Epstein–Barr virus erectile dysfunction endocrine disrupting chemical ejaculatory duct obstruction electroejaculation early growth response 1 enzyme-linked immunosorbent assay endogenous opioid peptide Environmental Protection Agency epididymal protease inhibitor expressed prostatic secretions estrogen receptor estrogen receptor-α knockout ectoplasmic specialization European Society of Human Reproduction and Embryology estrogen receptor European Society of Urogenital Radiology Scrotal and Penile Imaging Working Group epididymovasostomy Estrogen Therapy Food and Drug Administration functional hypothalamic amenorrhea International Federation of Gynecology and Obstetrics (FIGO) fluorescence in situ hybridization Friends of the Low-Cost Ivf Foundation Flamingo

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FNA FOAD FP Fr FSH FSHR Fzd G GABA GalNAc GAPDHS GAPDS GAS GGT GlcNAc GnRH GnRHR GPI GTP GU GWAS HA HA HAART hCG HDM HEPES HEX-B HHV HIF-1α HIV hMG HOS HPA HPF HPG HPO HPT HPV HR HSA HSG HSP HSP60 HSV HTF HTLV HyCoSy HZA HZI ICSI IDO IFFS IFRR

fine needle aspiration fetal origins of adult disease fertility preservation French follicle-stimulating hormone follicle-stimulating hormone receptor Frizzled gauge gamma aminobutyric acid N-acetylgalactosamine sperm-specific glyceraldehyde-3-phosphate dehydrogenase sperm-specific glyceraldehyde-3-phosphate dehydrogenase gender-affirming surgery γ-glutamyltranspeptidase N-acetylglucosamine gonadotropin-releasing hormone gonadotropin-releasing hormone receptor glycosyl phosphatidylinositol guanosine triphosphate genitourinary genome-wide association studies hyperactivated hyaluronic acid highly active antiretroviral therapy human chorionic gonadotropin histone demethylase N-hydroxyethylpiperazineN-ethanesulfonate hexosaminidase type B human herpesvirus hypoxia-inducible factor 1 alpha human immunodeficiency virus human menopausal gonadotropin hypo-osmotic swelling hypothalamic–pituitary–adrenal high-powered field hypothalamic–pituitary–gonadal hypothalamic–pituitary–ovarian hypothalamic–pituitary–testicular human papillomavirus hazard ratio human serum albumin hysterosalpingography heat shock protein 60-kDa heat shock protein herpes simplex virus human tubal fluid human T-cell leukemia virus hysterosalpingo-contrast sonography hemizona assay hemizona index intracytoplasmic sperm injection indoleamine 2,3-dioxygenase International Federation of Fertility Societies Infertility Family Research Registry

List of Abbreviations

IGD IGF IHH IIEF IL IM IMG IMSI INSL3 IP ISBER IUD IUI IVC IVF KD KNDy KS KSper LBD LCIVF LCR Lgl2 LH LHRH lncRNA LNG LPO LPS MA mAb MACS MAIS MAOI MAP MAPK MAR MAR MCAF MER MESA MGI MHC microTESE miRNA MIV MMAF MMAS MMP2 MMP9 MPOA

isolated gonadotropin-releasing hormone deficiency insulin-like growth factor idiopathic/isolated hypogonadotropic hypogonadism International Index of Erectile Function interleukin intramuscularly inferior mesenteric ganglia intracytoplasmic morphologically selected sperm injection insulin-like factor 3 intraperitoneal International Society for Biological and Environmental Repositories intrauterine device intrauterine insemination inferior vena cava in vitro fertilization knockdown kisspeptin/neurokinin B/dynorphin Klinefelter syndrome sperm-specific potassium ligand binding domain low-cost in vitro fertilization ligase chain reaction lethal giant larvae 2 luteinizing hormone luteinizing hormone-releasing hormone long noncoding RNA levonorgestrel lipid peroxidation lipopolysaccharide maturation arrest monoclonal antibodies magnetic-activated cell sorting mild androgen insensitivity syndrome monoamine oxidase inhibitor microtubule affinity protein mitogen-activated protein kinase medically assisted reproduction mixed agglutination reaction monocyte chemotactic and activating factor monocyte-to-eosinophil ratio microsurgical epididymal sperm aspiration Mouse Genome Informatics major histocompatibility complex microscopic/microdissection testicular sperm extraction microRNA minimally invasive vasectomy multiple morphologic abnormalities of the sperm flagella Massachusetts Male Aging Study matrix metalloprotease 2 matrix metalloprotease 9 medial preoptic area

MRI MRKH MSDS MT mTESE mV Mwh NBP ncRNA NES NHE NHL NHS NIEHS NIH NIOSH NKB NLR NNRTI NO NOS NPY NSAID NSV OA OAT OI OMIM OSHA PAH PAIS PAR PAS PatJ PBZ PCB PCD PCOS PCP PCR PCT Pd PDE PDE5 PDE5-I PDGF PESA PETG PEU PGC PGCN PID piRNA

magnetic resonance imaging Mayer–Rokitansky–Küster–Haus syndrome material safety data sheet microtubule microdissection testicular sperm extraction millivolt Multiple wing hairs nonbacterial prostatitis noncoding RNA nestorone sodium–hydrogen exchanger non-Hodgkin’s lymphoma Nance–Horan syndrome National Institute of Environmental Health Sciences National Institutes of Health National Institute for Occupational Safety and Health neurokinin B neutrophil-to-lymphocyte ratio non-nucleoside reverse transcriptase inhibitor nitric oxide nitric oxide synthase neuropeptide Y nonsteroidal anti-inflammatory drug no-scalpel vasectomy obstructive azoospermia oligoasthenoteratospermia obstructive interval Online Mendelian Inheritance in Man Occupational Safety and Health Administration polycyclic aromatic hydrocarbon partial androgen insensitivity syndrome pseudoautosomal region periodic acid–Schiff Pals1-associated tight junction protein phenoxybenzamide polychlorinated biphenyl primary ciliary dyskinesia polycystic ovary syndrome planar cell polarity polymerase chain reaction postcoital test prostatodynia phosphodiesterase phosphodiesterase type 5 phosphodiesterase type 5 isoform inhibitor platelet-derived growth factor percutaneous epididymal sperm aspiration polyethylene terephthalate postejaculatory urinalysis primordial germ cell paragigantocellular nucleus pelvic inflammatory disease Piwi-interacting RNA

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List of Abbreviations

PITX1 PKA PKC PLCζ PLR POI PPV PSA PTEN PVC PVE PVN PVS PVSA RB RCT rFSH rhFSH rhLH RI RISUG RNAi RNMS RNS ROK ROS RT-PCR RXFP2 SARM SART SAS SC SCD SCI SCO SCSA® SDF SEMG1 SERM SF1 SGE SHBG SHIM SHOX shRNA siRNA SIS sncRNA sNHE SNP SNRI

paired-like homeodomain transcription factor 1 protein kinase A protein kinase C phospholipase C zeta platelet-to-lymphocyte ratio primary ovarian insufficiency positive predictive value prostate-specific antigen phosphatase and tensin homolog polyvinyl chloride prostatovesiculoepididymitis paraventricular nucleus penile vibratory stimulation postvasectomy semen analysis reticulate body randomized controlled trial recombinant FSH recombinant human FSH recombinant human LH resistive index reversible inhibition of sperm under guidance RNA interference rare nonmotile sperm reactive nitrogen species Rho-associated kinase reactive oxygen species reverse transcriptase polymerase chain reaction relaxin family peptide receptor 2 selective androgen receptor modulator Society for Assisted Reproductive Technology sympathetic–adrenal system subcutaneously sperm chromatin dispersion spinal cord injury Sertoli cell-only Sperm Chromatin Structure Assay sperm DNA fragmentation semenogelin 1 selective estrogen receptor modulator steroidogenic factor 1 spinal generator for ejaculation sex hormone-binding globulin Sexual Health Inventory for Men short homeobox gene affecting stature short hairpin RNA small interfering ribonucleic acid saline infusion sonohysterography small noncoding RNA sperm-specific sodium–hydrogen exchanger single nucleotide polymorphism serotonin norepinephrine reuptake inhibitor

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SPA spp. SRR SSC ssDNA SSRI Stan StAR STD STI stRNA STS SV SVA T1/2 TAC TCA TDF TDS TdT TE TEFNA TESA TESE TET TF TGFβ THC TIMP-2 TJ TLR TM Tmax TMSC TNFα TRH TRUS TSH TTP TU TUIED TUNEL TURED UGCG UGT ULC UPA UPD US UTI Vangl2 VDAC3 VE VEGF VEGFr

sperm penetration assay species sperm retrieval rate spermatogonial stem cell single-stranded DNA selective serotonin reuptake inhibitor starry night steroidogenic acute regulatory protein sexually transmitted disease sexually transmitted infection small temporal RNA sequence tagged site seminal vesicle seminal vesicle aspiration half-life total antioxidant capacity tricyclic antidepressant testis-determining factor testicular dysgenesis syndrome terminal deoxynucleotidyl transferase testosterone enanthate testicular fine needle aspiration testicular sperm aspiration testicular sperm extraction ten-eleven translocation tissue factor transforming growth factor beta tetrahydrocannabinol tissue inhibitor of metalloproteinase-2 tight junction Toll-like receptor testicular microlithiasis time to maximum serum concentration total motile sperm count tumor necrosis factor alpha thyrotropin-releasing hormone transrectal ultrasonography/ultrasound thyroid-stimulating hormone time to pregnancy testosterone undecanoate transurethral incision of ejaculatory ducts terminal deoxynucleotidyl transferase deoxyuridine triphosphate nick end labeling transurethral resection of the ejaculatory ducts UDP-glucose ceramide glucosyltransferase UDP-glucuronosyltransferase Uniform Law Commission Uniform Parentage Act uniparental disomy ultrasound urinary tract infection Van Gogh-like 2 voltage-dependent anion channel 3 vasoepididymostomy vascular endothelial growth factor vascular endothelial growth factor inhibitor

List of Abbreviations

VHL VR VV VVSG VZV WBC

von Hippel–Lindau vasectomy reversal vasovasostomy Vasovasostomy Study Group varicella-zoster virus white blood cell

WHO WPATH YCMD ZIKV ZPBP

World Health Organization World Professional Association for Transgender Health Y chromosome microdeletion Zika virus zona pellucida binding protein

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Introduction Craig S. Niederberger, Dolores J. Lamb, Larry I. Lipshultz, and Stuart S. Howards

The last and fourth edition of Infertility in the Male was published in 2009, and significant advances were realized in reproductive medicine and surgery in the intervening decade. In this edition, we have covered the more recent advances in the field while maintaining the core foundation of information needed for practitioners in diagnosing and treating the man seeking care for fertility. We have also endeavored to make the book more structured, and hopefully easier to use, for the student and specialist alike. For the first time, we have organized the book into sections: “Scientific Foundations of Male Infertility,” the basic biological science undergirding reproductive medicine; “Clinical Evaluation of the Infertile Male,” which covers clinical diagnosis; “Laboratory Diagnosis of Male Infertility,” detailing laboratory diagnosis of testicular dysfunction and the basics of sperm cryopreservation; “Treatment of Male Infertility,” describing the means and strategies for therapy for these diagnoses; and finally “Health Care System and Culture,” which contextualizes male fertility care in society and the world. Many of these chapters have substantial overlap, as they consider topics from more than one perspective – while the chapter “Cryopreservation of Sperm – History and Current Practice” in the “Laboratory Diagnosis of Male Infertility” section attends to the history and laboratory processes of storing sperm for future use, “Male Oncofertility – Considerations for Fertility Preservation and Restoration” in the “Treatment of Male Infertility” section describes the conditions the clinician will encounter to utilize banking; “Sperm Retrieval Surgery” details how to surgically obtain sperm, and “The Use of Sperm in Medically Assisted Reproduction” explains how to use cryopreserved sperm in medically assisted reproduction techniques such as in vitro fertilization/intracytoplasmic sperm injection. While chapters in the fourth edition included sentences in bold to draw the attention of the reader to their most pertinent parts, to facilitate the use of the book in

practice, chapters now also include Key Points in boxes to facilitate and cement understanding and real-world use. Multiple related chapters in the fourth edition were combined – thus, although there are fewer chapters in this book, compared to its predecessor, they are deeper, more interrelated, and more understandable. The section “Scientific Foundations of Male Infertility” begins, as did the previous edition, with a chapter detailing the anatomy and embryology of the male reproductive tract and gonadal development, the epididymis, and accessory sex organs, thus forming the basis of accurate anatomic diagnosis and surgery. The following chapter describes the complex interplay of cells and their communicating molecules that coordinate the production of sperm in the testis; its immediate succeeding chapter details how and what happens to sperm in the epididymis that makes them capable of fertilizing the ovum. As the male reproductive system is largely controlled by the endocrine system, a chapter follows describing the production and control of sex steroids in the male, laying the essentials for accurate endocrine therapy detailed later in the book. Once sperm is made, it must make its exit, and the chapter on erection, emission, and ejaculation then addresses these processes. Science never sleeps, and the final chapter in this section describes the enormous leaps in genomic modification and epigenetics during the last decade that are sure to be the foundation for diagnostic and therapeutic advances in the years to come. The next section “Clinical Evaluation of the Infertile Male” brings our current knowledge of male reproductive pathology and its diagnosis to the armamentarium of the male fertility specialist. It begins with one of the most rapidly evolving areas in the field, our understanding of how other diseases are related to reproductive dysfunction, a chapter on “Infertility as a Metric of Men’s Health.” This presents one of the most important reasons why we care for male infertility – it may reveal significant underlying health conditions. Following is the chapter

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Introduction

“Office Evaluation of the Subfertile Male” that gives the practitioner concrete strategies to be used in the office encounter, including questions to ask, what to look for, and clinical interpretation of the semen analysis. As the field is unusual, in that two people are required for an outcome, “Evaluation of the Infertile Male’s Partner” provides a high-level review of the diagnosis of the female. By reading it, the practitioner will have a clear understanding of the steps taken in parallel by the female fertility specialist in order to best integrate reproductive care. “Imaging the Male Reproductive System” provides the reader with when and how to use radiographic and ultrasonographic tools in the diagnosis of the infertile male and, importantly, when they are not necessary. Another area of explosive growth in the field in the past decade has been in our understanding of environmental toxicants and their effect on male reproduction, reviewed in “Effects of Environmental Chemicals on Male Reproduction.” With “Endocrine Causes of Male Infertility – Diagnosis and Treatment,” the foundation presented in the chapter detailing the male endocrine system in the prior section is carried forward into pathological endocrine states and how to diagnose them. The chapter “Spermatogenesis – Diagnosis of Normal and Abnormal States” provides an overview of spermatogenic pathology and its diagnosis, integrating the basic knowledge describing spermatogenesis in the prior section with related systems in this section, as well as providing context for treatment to be detailed more completely in a subsequent section on therapy. The chapter “Inheritance and Male Fertility” delineates genomic conditions manifesting as male reproductive dysfunction and carries forward the epigenetic background laid in the prior section into what practitioners need to consider in the clinic. Still bedeviling clinicians and patients alike, the commonly encountered varicocele is elucidated in the chapter bearing its name, including its history, pathophysiology, diagnosis, indications for treatment, and, as this chapter is targeted to a specific condition, the treatment itself. The section concludes with a chapter detailing infectious and immunological considerations in the diagnosis of male infertility, an often confounding area for those diagnosing and treating the infertile male. With a clear understanding of the material presented thus far, the practitioner is ready to diagnose any man presenting with infertility using the tools currently available in reproductive clinical science. In the time-tested process of clinical evaluation of male reproductive dysfunction, the practitioner next

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obtains laboratory testing. The next section begins with an overview of the two pillars of reproductive laboratory assessment – endocrine and sperm – in “The Laboratory Evaluation of the Infertile Male” and provides a highlevel overview of other topics such as sperm DNA fragmentation. These cutting-edge forms of assessment of male infertility are substantially expanded in the subsequent chapter “Advanced Diagnostic Approaches to Male Infertility” that details the myriad forms of sperm DNA integrity assays and which and when they are best used, and an encyclopedic list of currently known genomic defects affecting male fertility that are currently used for clinical diagnosis in some parts of the world. Sperm are dynamic cells, and the following chapter “Evaluating Defects in Sperm Function” describes the assays used in determining how well sperm swim and do their job in fertilizing an ovum. As the laboratory is critical in freezing sperm for future use, the final chapter in the section “Cryopreservation of Sperm – History and Current Practice” describes methods of preserving sperm, while contextualizing these techniques in their use both specifically as a therapy and broadly in a health care system. With an understanding of the biological science of male reproduction and how to diagnose its dysfunction in the clinic and laboratory, the reader is now prepared to treat specific conditions of male infertility in the next section. The first chapter “Medical Treatment of Male Infertility” reviews endocrine therapy, nonendocrine medicines, and nutraceuticals. Should sperm be produced in the testis but encounter barriers to traversing and exiting the male reproductive tract, “Surgery to Improve Sperm Delivery” details the procedural methods to address the various causative problems. Should the making of sperm in the testis be at fault or it not be possible to alter the reproductive tract to deliver sperm, going to the source of sperm in the male gonad is necessary and the subsequent chapter “Sperm Retrieval Surgery” describes when and how to do so. If sperm is obtained from the testis or present in low quantities in the ejaculate, medically assisted reproduction is required and the chapter “The Use of Sperm in Medically Assisted Reproduction” details those methods, providing the practitioner with an understanding of what happens to sperm in the laboratory and beyond. During the past decade, a field coined “oncofertility” has expanded into a systematic approach to fertility preservation for cancer survivors, and the chapter bearing its name describes all aspects of this in great detail. The other side of fertility is when it no longer is desired, and the chapter “Male

Introduction

Contraception” enumerates the many exciting advances that may soon lead to options beyond female oral contraceptives, barrier methods, and vasectomy. The section concludes with the thrilling advances coming in male fertility therapy in “Future Directions in Male Infertility.” The final section puts fertility care in its context in the health care system and in society. Mental health is an indispensable component of care, as patients are often devastated to learn that they have problems in the most basic of human desires, and the chapter “Mental Health and Male Reproduction” attends to this critical facet of care. Reproduction is a social construct with often legal implications and needs, and the chapter “Legal Issues and Male Reproduction” offers a review of those most pertinent. Gender affirmation procedures have become more common during the last decade, and those individuals needing these procedures should be offered the opportunity for children, with considerations prior to and after treatment; the chapter “Male Reproduction in the Transgender Patient” concretely reviews this rapidly evolving field. Finally, male reproductive care is not limited to one corner of the world, and the chapter “Global and Cultural Aspects of Male Reproductive Care” reviews various customs and practices to which practitioners must be sensitive in order to effectively and respectfully provide care. A few notes before we release you into the world of male reproductive health care. The first is one of nomenclature. In the early days of treating male infertility, there were no options for spermatogenic dysfunction such as testis sperm extraction with intracytoplasmic sperm injection. Hence, in diagnosing azoospermia, there were only two forms – one involving obstruction that may be addressed with surgical correction, and the other due to spermatogenic dysfunction where nothing was available. One was, and still is, termed “obstructive azoospermia,” an apt and accurate name for the condition. The other was called “nonobstructive azoospermia,” as nothing could be done for it. Today, that second form is very

commonly treated with medicine and surgery, and to continue to call it “nonobstructive azoospermia” does little to explain what it actually is. We have begun more commonly using the descriptive term “azoospermia due to spermatogenic dysfunction,” and you will find that terminology in this book. However, in this edition, as the older term “nonobstructive azoospermia” remains in the literature, we have left it to individual chapter authors which nomenclature they prefer for azoospermia when the pathology is spermatogenic dysfunction. You will also find disagreements and multiple viewpoints when authors cover similar topics. We heartily encouraged these, as medicine is as much art as science, and the interpretation of the medical literature is not an exercise in revealing absolute truths. We chose leading practitioners and scientists to write chapters, but every human has orientations and biases. We want you to see those for what they are and to choose your own interpretations and clinical strategies. Finally, as we conclude the compilation of this book, we are missing something extremely important – the effect of the COVID-19 pandemic on male reproduction. Early studies suggest that men convalesced from the disease do not harbor virus in semen, but these results are highly preliminary [1]. With the tragic, staggering numbers of those infected, we will ultimately have more information; however, at this time, we do not have sufficient study data to clearly discuss the effects, since much of this disease still remains a mystery. In future years, we will know more. With those notate bene, it is time for you to begin the book. Happy reading, and we wish you all the best in your care for men struggling with infertility!

References 1.

Pan F, Xiao X, Guo J, et al. No evidence of severe acute respiratory syndrome-coronavirus 2 in semen of males recovering from coronavirus disease 2019. Fertil Steril 2020;113:1135–9.

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Section 1 Chapter

1

Scientific Foundations of Male Infertility

Anatomy and Embryology of the Male Reproductive Tract and Gonadal Development, the Epididymis, and Accessory Sex Organs Danielle Velez and Craig S. Niederberger

Introduction To understand the range of male infertility conditions, knowledge of normal male anatomy and its embryology is essential. This chapter will review the major developmental events from fertilization to organization of the germ layers into organ systems. It will review in depth the formation of the male genitourinary system and external genitalia. By laying a strong foundation in embryology and anatomy, providers will have a more thorough understanding of disease states.

Fertilization to Germ Layers After fertilization of the ovum by sperm, a genetically unique zygote is formed on day two. The zygote undergoes multiple mitotic divisions to create the morula (from the Latin word for mulberry) on day three. The internal cells of the morula are surrounded by trophoblastic cells, which will later become the placenta. By day four, a fluid-filled cavity called the yolk sac, or umbilical vesicle, appears and separates the trophoblast from the developing embryo. The subsequent blastocyst then attaches to the uterine wall on day six, stimulating the trophoblast to rapidly proliferate and differentiate into its two layers: the inner cytotrophoblast and the outer syncytiotrophoblast. The syncytiotrophoblast invades the endometrium, embedding the blastocyst and establishing early uteroplacental circulation, as shown in Fig. 1.1. With implantation, the blastocyst divides into a bilaminar embryonic disc of pluripotent epiblast and hypoblast. Other structures that form during the second week include the amniotic cavity, amnion, umbilical vesicle, and chorionic sac. The syncytiotrophoblasts produce human chorionic gonadotropin (hCG), which maintains the corpus luteum to secrete estrogen and progesterone throughout pregnancy [1]. Gastrulation takes place during the third week with proliferation and movement of epiblast cells on the embryonic disc, creating the primitive streak and notochord. The bilaminar embryonic disc matures into the three germ

layers: the embryonic endoderm, mesoderm, and ectoderm. The primitive streak forms the mesoderm until the fourth week, after which it degenerates and disappears. This mesoderm further differentiates into paired structures straddling the notochord; from medial to lateral, these are the paraxial, intermediate, and lateral mesoderm (Fig. 1.2). The urinary and genital systems largely develop from the intermediate mesoderm. Around day 16, a caudal outpouching from the umbilical vesicle extends into the connecting stalk of the developing placenta. This diverticulum is known as the allantois. The proximal part of this diverticulum will eventually fibrose and become the adult urachus, or median umbilical ligament. The mesoderm of the allantois expands into blood vessels, which feed the placenta as the umbilical arteries. Late in the fourth week of development, as the embryo folds in the horizontal plane, a longitudinal elevation of the mesoderm appears on either side of the dorsal aorta, called the urogenital ridge. This urogenital ridge gives rise to the nephrogenic cord and the gonadal ridge.

Key Points • Following fertilization, the zygote undergoes multiple mitotic divisions to form the morula on day two, then implants into the endometrium as a blastocyst around day six. • By the third week of development, the embryo has divided into its three main germ layers: the endoderm, mesoderm, and ectoderm. • The majority of the urogenital system is derived from the intermediate mesoderm.

Renal Development The fetal kidney passes through three phases in establishing its permanent form: the pronephros,

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Endometrial capillary

Uterine gland

Amniotic cavity

Epiblast

Endometrial epithelium Syncytiotrophoblast

Hypoblast

Cytotrophoblast Primary umbilical vesicle Fig. 1.1 Blastocyst implantation into the endometrium.

Notochord

Neural groove Ectoderm

Endoderm Paraxial mesoderm Intermediate mesoderm

Lateral mesoderm

Fig. 1.2 Transverse section of the embryo, highlighting the relationship between the paraxial, intermediate, and lateral mesoderms prior to lateral folding.

mesonephros, and metanephros. The bilateral pronephroi appear early in the fourth week near the developing neck. The pronephric ducts run caudally and open into the cloaca (from the Latin word for sewer), which drains the allantois and hindgut. The majority of the pronephros degenerates as the mesonephros develops at the end of

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the fourth week. Fig. 1.3 shows the progression of the nephrogenic cord as it passes from the pronephros to the mesonephros and metanephros, with its insertion into the cloaca. The mesonephros are made up of primitive glomeruli and mesonephric tubules, which drain into the mesonephric ducts. These structures serve as an interim kidney for 4 weeks, until the appearance of the metanephros. Although the majority of the mesonephros also involutes by week 12, the mesonephric duct persists in the male as the appendix of the epididymis, ductus deferens, ejaculatory duct, and seminal vesicles (Table 1.1). The metanephros develops in the sacral region in the fifth week, and is functional by weeks 9–10. It consists of the ureteric bud (metanephric diverticulum) and the metanephrogenic blastema. Through reciprocal induction, the ureteric bud and metanephrogenic blastema become the collecting system and nephron, respectively. The ureteric bud begins as a diverticulum of the mesonephric duct, near its insertion into the cloaca. Influenced by the metanephrogenic blastema, the ureteric bud branches repeatedly, growing from the ureter to the renal pelvis, major and minor calyces, and collecting tubules, as shown in Fig. 1.4. Each tubule induces clusters of mesenchymal cells in the metanephrogenic blastema to differentiate into parts

Chapter 1: Anatomy and Embryology

Degenerating part of pronephric duct

Cervical somites

Pronephros

Pronephros Nephrogenic cord Allantois Liver primordium Omphalomesenteric duct

Mesonephros

Mesonephros

Mesonephric tubules

Metanephros

Allantois

Cloaca

Cloaca

Ureteric bud

A. Lateral view of the nephric system development during the fifth week, from the pronephros to the mesonephros to the metanephros.

B. Coronal view of the nephric system development.

Fig. 1.3 Lateral view of the nephric system development during the fifth week, from the pronephros to the mesonephros to the metanephros.

of the nephron: the glomerulus, proximal and distal convoluted tubules, and loop of Henle. Failure of the ureteric bud to associate with the metanephrogenic blastema will result in renal agenesis and often also results in maldevelopment of the other Wolffian duct structures, including the ipsilateral epididymis, vas, seminal vesicle, and ejaculatory duct [2–4]. Thus, when unilateral vasal agenesis is encountered, the patient should be referred for renal ultrasound to evaluate the upper tracts. Urine production begins around week 10 of development. The number of glomeruli increases throughout gestation until week 36, so that nephron formation is complete at birth, with each kidney containing 200 000–2 million nephrons [5]. The paired metanephroi develop within the pelvis, and with abdominal growth, they gradually rise and rotate, until the hilum faces anteromedially and the superior portion of the kidney is in contact with the adrenal glands. The renal blood supply changes with ascent, initially coming from the common iliac arteries, then the abdominal aorta, as it reaches its final position in the retroperitoneum. For this reason, it is not uncommon for kidneys to have accessory renal arteries.

Bladder Development The cloaca is the early chamber where the hindgut and allantois empty, and it is covered by the cloacal

membrane. Two theories exist regarding separation of the cloaca into the dorsal rectum and ventral urogenital sinus. Traditionally, cloacal division was thought to occur between weeks 4 and 7 through a combination of coronal folding and caudal descent of the urorectal septum. Fusion of the urorectal septum with the cloacal membrane was thought to form the perineal body. However, microscopic studies, aided with computer-assisted three-dimensional reconstruction, have suggested that although the distance between the urorectal septum and the cloacal membrane decreases, the two structures do not fuse. Rather, there is flattening, and then apoptosis, of the cloacal membrane, ending in rupture around week 7. This leads to exposure of two surfaces: the anterior urogenital groove and the posterior anal orifice. The area in between, which is the tip of the urorectal septum, becomes the future perineum [6, 7]. Early rupture of the cloacal membrane and failure of mesenchymal cell migration between the abdominal ectoderm and the cloacal endoderm result in the bladder exstrophy–epispadias complex. The bladder forms from the urogenital sinus above the mesonephric (Wolffian) ducts. It begins as a cylindrical tube lined by a single layer of cuboidal cells and is surrounded by loose mesenchymal tissue. From weeks 7 to 12, the surrounding connective tissue condenses and smooth muscle appears. Mature urothelial epithelium is present by week 17. Bladder development is guided by several signaling factors, including Shh,

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Table 1.1. Derivatives and vestigial remnants of embryonic urogenital structuresa Embryonic structure

Female

Male

Indifferent gonad

Ovary

Testis

Cortex

Ovarian follicles

Seminiferous tubules

Medulla

Rete ovarii

Rete testis

Gubernaculum

Ovarian ligament Round ligament of uterus

Gubernaculum testis

Mesonephric tubules

Epoophoron Paroophoron

Efferent ductules of testis Paradidymis

Mesonephric duct

Appendix vesiculosa Duct of epoophoron Longitudinal duct (Gartner duct)

Appendix of epididymis Duct of epididymis Ductus deferens Ejaculatory duct and seminal gland

Stalk of ureteric bud

Ureter, pelvis, calices, and collecting tubules

Ureter, pelvis, calices, and collecting tubules

Paramesonephric duct

Hydatid (of Morgagni) Uterine tube Uterus, cervix

Appendix of testis

Urogenital sinus

Urinary bladder Urethra Vagina Urethral and paraurethral glands Greater vestibular glands

Urinary bladder Urethra (except navicular fossa) Prostatic utricle Prostate Bulbourethral glands

Sinus tubercle

Hymen

Seminal colliculus

Primordial phallus

Clitoris Glans clitoris Corpora cavernosa of clitoris Bulb of vestibule

Penis Glans penis Corpora cavernosa of penis Corpus spongiosum of penis

Urogenital folds

Labia minora

Ventral aspect of penis

Labioscrotal swellings

Labia majora

Scrotum

a

Functional derivatives are in italics. (From Moore KL, Persaud TVN, Torchia MG. The Developing Human: Clinically Oriented Embryology, 11th ed, 2020. Table 12.1: Derivatives and Vestigial Remnants of Embryonic Urogenital Structures)

Bmp4, and Fgfr2 [8]. Around week 10, when urine production begins, the ratio of elastic-to-collagen fibers increases and that of thick-to-thin collagen fibers decreases, improving bladder compliance. The layers of the adult bladder are the urothelium, lamina propria, muscularis mucosa, smooth detrusor muscle, and perivesical fat. The trigone of the bladder is formed by a combination of smooth muscle from the bladder and incorporation of the ureters from the mesonephric (Wolffian) duct. The mesonephric duct caudal to the take-off of the ureteric bud is known as the common excretory (nephric) ducts, as shown in Fig. 1.5 [8]. As the common excretory duct contacts the developing bladder, it undergoes apoptosis, separating from the developing ureter, as seen in Fig. 1.5C and D. Through

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remodeling, expansion, and differentiation, the ureteral orifices move superolaterally and ultimately enter obliquely through the base of the bladder within the trigone (Fig. 1.5D, E) [9]. Adequate detrusor backing is required to prevent the reflux of urine from the bladder to the ureter/renal pelvis as the bladder fills. Vesicoureteral reflux can result in urinary tract infections, renal parenchymal damage, and hypertension. In males, the Wolffian ducts migrate caudally, entering the prostatic urethra at the verumontanum as the ejaculatory ducts (Fig. 1.5E). External striated muscle fibers appear around week 15 at the caudal end of the urogenital sinus, while internal smooth muscle layers thicken at the bladder neck, laying the groundwork for the future continence mechanism.

Chapter 1: Anatomy and Embryology

Minor calyces

Renal pelvis Ureter

Major calyces

Remnant of pronephros Nephrogenic cord

Branching ureteric bud

Developing liver

Mesonephros

Ureteric bud Cloaca Mesonephric duct Ureteric bud

Metanephrogenic blastema

Mesonephric duct Metanephrogenic blastema

Fig. 1.4 Successive stages of ureteric bud development, from the fifth to eighth weeks: ureter, renal pelvis, and major and minor calices.

Key Points • The fetal kidney passes through three stages before arriving at its adult form: the pronephros, mesonephros, and metanephros. The pronephros and mesonephros largely degrade, but the mesonephric duct persists as Wolffian structures in the male: the ejaculatory duct, vas deferens, epididymal tail, and seminal vesicles. In both male and female fetuses, the mesonephric duct gives off the ureteric bud. • The metanephros forms via reciprocal induction between the ureteric bud and the metanephrogenic blastema, resulting in the collecting system and renal parenchyma, respectively. • The urorectal septum divides the cloaca into the anterior urogenital sinus and posterior rectum (hindgut). Urine production begins

around week 10 of gestation, coinciding with bladder development from the urogenital sinus.

Male Gonadal and Genital Duct Development To review, there are a series of ducts that give rise to the genitourinary system. The pronephric duct is primarily for the rudimentary urinary system and gives rise to the mesonephric duct, which becomes coopted by the genital system in males. The paramesonephric duct is also primarily for the genital system; however, the fates of the mesonephric and paramesonephric ducts are determined by the chromosomal makeup of the fetus. Due to the lack of anti-Müllerian hormone (AMH) in 46XX fetuses, 46XX fetuses retain their paramesonephric ducts while the paramesonephric duct will largely involute in 46XY fetuses.

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A

Wolffian B duct

Genital ridge

BI

Hindgut

Aorta Primordial germ cells

Allantois

Ureteric bud Ureter Common nephric duct D

C

BI

BI

Ejaculatory duct

Urethra

E Bladder Ureter

Trigone

Urethra

Prostate

Ejaculatory duct Fig. 1.5 Origin of the ureteric bud from the Wolffian duct (A) and remodeling of the positions of the ureters and Wolffian ducts in male embryos (B–D). (E) shows the final positions of the ureters and ejaculatory ducts, in relation to the trigone.

For both sexes, gonadal development begins in the fifth week with development of the gonadal ridge, a thickened area of mesothelium medial to the mesonephros. Gonadal cords grow into the underlying mesenchyme and are joined by primordial germ cells, which are the earliest undifferentiated sex cells. These cells begin in the yolk sac and migrate along the dorsal mesentery of the hindgut to the gonadal ridge (Fig. 1.6), forming the primitive sex cords.

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Fig. 1.6 Primordial germ cells, shown in blue, migrating from the wall of the yolk sac along the dorsal mesentery of the hindgut to the developing mesenchyme of the bilateral genital ridges.

In the seventh week of gestation, this indifferent gonad is identical between 46XX and 47XY embryos, with an epithelial cortex and a mesenchymal medulla. In the 46XX embryo, the cortex of the indifferent gonad will differentiate into an ovary, with subsequent degeneration of the medulla. Conversely, the cortex will regress and the medulla will develop into a testis in the 47XY embryo. The short arm of the Y chromosome contains the sexdetermining region (SRY gene), which produces the SRY protein, also known as testis-determining factor (TDF). Without the SRY gene, the ovary is the default future for the indifferent gonad. The SRY protein induces the cells in the medullary region of the indifferent gonad to form testicular epithelial cords, which will eventually become seminiferous tubules. The primordial germ cells develop into spermatogonia, while the surrounding mesenchyme becomes Sertoli cells. The SRY gene influences seminiferous tubule differentiation through multiple regulating genes: Sox9, Wnt4, Foxl2, Fst, and Rspo1 [1]. Fig. 1.7 demonstrates the effect of TDF on the indifferent gonad (left), in comparison to the genital course taken without the presence of TDF (right). In the sixth and seventh weeks, Sertoli cells begin secreting AMH, which suppresses paramesonephric duct development, thereby preventing formation of the uterus, Fallopian tubes, and upper vagina. The mesenchyme separating the seminiferous cords matures into interstitial Leydig cells, which begin producing

Chapter 1: Anatomy and Embryology

Aorta

(A) Undifferentiated gonad

Mesonephric duct

Primordial germ cells Gonadal ridge

Primary sex cord

TDF

No TDF

Seminiferous cords

Cortical cords from surface epithelium

Rete testis Mesonephric duct and tubule

(B) Epididymal duct

(C)

Degenerating rete ovarii Ser to

li cell

Dissolving paramesonephros

s

Paramesonephros developing into the uterine tube

Primordial ovarian follicle

Stroma Spermatogonium

Oogonium

Fig. 1.7 (A) Undifferentiated gonads of the 5-week embryo. (B) Testis development is shown on the left. The testis-determining factor on the Y chromosome forces regression of the cortex and development of the medulla into the testicle. The gonadal cords become the rete testis and seminiferous tubules. The bottom left of this image shows the two main cells of the testis: spermatogonia (from the primordial germ cells) and Sertoli cells (from the mesenchyme). Note the lack of the paramesonephros on the left, as this degenerates in the male due to anti-Müllerian hormone. (C) Ovarian development is shown on the right. The primordial germ cells enter the cortical cords, which have developed from the surface epithelium. In contrast to the testis, the medulla of the undifferentiated gonad regresses and the cortex becomes the ovary. Note that the paramesonephros persists and develops into the uterine tube.

testosterone in the eighth to ninth weeks under the influence of placental hCG. Later, as the pituitary gland develops, fetal gonadotropins take over androgen production. Testosterone and its more potent derivative, dihydrotestosterone, masculinize the mesonephric duct (Wolffian duct), genital tubercle, and urogenital sinus (as noted in Table 1.1) [5]. Type 2 5-alpha reductase, secreted by the urogenital sinus and genital tubercle, converts testosterone to dihydrotestosterone, which is essential for normal penile and prostatic development.

Testosterone from fetal Leydig cells stimulates the mesonephric duct to become the Wolffian structures. Although the lumen of the seminiferous tubules will not develop until puberty, the walls of the seminiferous cords are made of spermatogonial stem cells and Sertoli cells. The seminiferous cords condense and extend into the rete testis, which grows into 15–20 mesonephric tubules. These tubules will become the efferent ductules. Distally, the mesonephric duct develops into the duct of the epididymis and, as it becomes enveloped in smooth

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muscle, thickens into the vas deferens. The caudal end of the mesonephric duct enters medially at the prostatic urethra as the ejaculatory duct. Laterally, the mesonephric duct gives rives to the seminal vesicles. There is very limited research on seminal vesicle development. Brewster [10] documented the first evidence of distal bulbous dilations from the mesonephric duct in the 12th week of gestation. Over the following 13 weeks, these dilations grow, forming branching diverticula and folding into concentric circular layers of undifferentiated mesenchyme surrounding an epithelial-lined lumen. The mesenchyme gradually differentiates into the lamina propria, surrounded by the tunica muscularis and adventitia postnatally. Fig. 1.8 shows the progression in seminal vesicle development from 14 weeks’ gestation to a 1-day-old infant to an 18-year-old male. Similar to the prostate, branching

morphogenesis of the seminal vesicle is directed by local androgens and fibroblast growth factor, Hedgehog, and transforming growth factor-beta factors [11]. The seminal vesicle contributes fructose, ascorbic acid, prostaglandins, and coagulation factors to the male ejaculate [12]. The prostate is unique amongst the other male accessory sex glands in that it is derived from the endodermal urogenital sinus. Around weeks 10–12, endodermal outgrowths at the base of the fetal bladder grow into the surrounding mesenchyme. Hox genes control glandular morphogenesis, which results in mesenchymal differentiation into the dense stroma and smooth muscle of the prostate [13]. Fibroblast growth factor 7, activin A, insulin-like growth factor 1, and bone morphogenetic proteins 4 and 7 have been implicated in the repeated branching of the prostatic ducts from their urethral insertion to the

Fig. 1.8 (A) A 14-week embryo, showing the beginnings of a tubular seminal vesicle, growing from the ampulla. AA, ampulla; ED, ejaculatory duct; P, prostate; PD, paramesonephric duct; SV, seminal vesicle. (B) Section through the seminal vesicle of a 1-day-old infant. A diverticulum is branching from the main lumen. The lumen contains cellular debris and mucoid substance. BC, basal cells; D, diverticulum; L, lumen. M, smooth muscle; PC, principal cells. (C) Section through the seminal vesicle of an 18-year-old patient, showing two layers of tunica muscularis and the circumferential nature of the epithelial folding. BC, basal cells; L, lumen; LP, lamina propria; PC, principal cells; TM, tunica muscularis. (From Reference [10].)

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Fig. 1.9 Section of a prebud 9-week human fetal urogenital sinus (UGS) in the region of the verumontanum, a dorsal hillock projecting into the UGS (dotted line). The Wolffian ducts (WDs) flank the Müllerian-derived prostatic utricle (marked), and both open into the UGS at/near the apex of the verumontanum. The section is stained by FOXA1, an endodermal marker. (From Reference [5].)

Fig. 1.8 (cont.)

distal ductal tips [5]. As previously noted, prostate differentiation is guided by fetal androgens, namely dihydrotestosterone, which peak during the second trimester. Prostate development is highly sensitive to varying steroid hormones. In utero exposure to elevated maternal or exogenous estrogens, such as bisphenol A, during prostate development results in decreased prostatic growth and increased risk of prostate cancer in adulthood [14]. This is likely secondary to permanent alterations in DNA methylation patterns. With regard to male fertility, the prostate is the main source of acid phosphatase, citric acid, inositol, calcium, zinc, magnesium, and liquefying factors found in the ejaculate [12]. The verumontanum elongates in a craniocaudal fashion on the dorsal wall of the male urethra, and is the entrance for the terminal mesonephric (Wolffian) ducts and any remnant Müllerian duct, which takes the form of a prostatic utricle. As seen in Fig. 1.9, the verumontanum is an interface between the mesodermal Wolffian duct and the endodermal Müllerian duct [5]. Similarly, smaller glands derived from the spongy urethra and the adjacent

mesenchyme develop into the paired bulbourethral glands. The bulbourethral glands contribute about 5 percent of seminal volume, mostly as a lubricating mucus within the male urethra [12]. Testicular development begins in the upper lumbar region near the developing kidney. The germ cells, Sertoli cells, and Leydig cells become surrounded by a thick, fibrous capsule of connective tissue known as the tunica albuginea. As shown in Fig. 1.10A, the testis is supported by a cranial and caudal suspensory ligament, the latter of which becomes the gubernaculum. As the mesonephros and paramesonephros degrade and the fetal intra-abdominal pressure increases, the testes descend caudally along the posterior abdominal wall, typically between weeks 10 and 15. As the testis descends, the cranial suspensory ligament degrades and the gubernaculum thickens (Fig. 1.10B), eventually bulging obliquely beyond the future external inguinal ring and into the scrotum. This process is facilitated by hollowing of the peritoneum, allowing for a ventral evagination of the peritoneal cavity known as the processus vaginalis. As the gubernaculum shortens, pulling the testicle through the inguinal canal into the scrotum, it drags the surrounding processus vaginalis, and thus the abdominal wall layers with it, to surround the testis and spermatic

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CSL

WD MD G CGRP A

B

development of the female sex organs from the paramesonephric ducts. Midline fusion of the paramesonephric ducts, regulated by HoxA10, results in formation of the uterus, as well as of the upper twothirds of the vagina, broad ligament, rectouterine pouch, and vesicouterine pouch [1]. The remainder of the paramesonephric duct that does not fuse gives rise to the Fallopian tubes. The distal third of the vagina, as well as the hymen, appears to originate from the urogenital epithelium [5].

C

Fig. 1.10 The stages of testicular descent. (A) Before descent, the developing testis is held in the urogenital ridge by the cranial suspensory ligament (CSL) and the gubernaculum (G). The adjacent Wolffian duct (WD) forms the epididymis and vas deferens in the male, and the Müllerian duct (MD) forms the uterus and Fallopian tubes in the female. (B) Around week 15, the transabdominal journey has completed and the testis is held near the future inguinal ring by dilation of the gubernaculum. (C) The inguinoscrotal phase requires the gubernaculum to elongate into the scrotum, under control of androgens and calcitonin gene-related peptide (CGRP). After migration is complete, the peritoneum of the processus vaginalis (PV) closes and then completely involutes and disappears. (From Reference [5].)

cord: the transversalis fascia, internal oblique fascia, and external oblique fascia. The opening of the transversalis fascia becomes the internal inguinal ring, and the opening of the external oblique the external inguinal ring. The processus vaginalis typically involutes, leaving the testis almost completely surrounded by the tunica vaginalis, except at the entrance of the vas deferens and testicular blood vessels (Fig. 1.10C). A persistent patent processus vaginalis may result in an inguinal hernia and/ or a hydrocele [15]. Testicular descent is controlled by fetal androgens and is typically complete by 32 weeks. Cryptorchidism, or undescended testes, is present in up to 9 percent of all newborns, with approximately 40 percent of these testes descending spontaneously by three months of age [16].

Female Gonadal Development Bladder development above the entry of the mesonephric ducts is the same between male and female fetuses. In the 46XX fetus, the caudal portion of the urogenital sinus develops into the urethra in a similar fashion to the 46XY fetus, although without phallic lengthening. Due to the lack of AMH and testosterone in the female, there is persistence of the paramesonephric ducts and involution of the Wolffian ducts. Maternal estrogen facilitates

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Key Points • The short arm of the Y chromosome contains the SRY gene, which produces TDF. TDF promotes Sertoli cell development and differentiation of the primitive germ cells into spermatogonia. Sertoli cells produce AMH, which prevents formation of the uterus, Fallopian tubes, and upper two-thirds of the vagina from the paramesonephric ducts. Leydig cell production of testosterone influences the mesonephric duct to form the Wolffian structures. • The prostate originates from endodermal outgrowths at the base of the bladder around week 10. This is guided by dihydrotestosterone, which is produced from testosterone via 5-alpha reductase. The testicles form in the abdomen, then descend from weeks 10 to 32 to the scrotum, guided by the gubernaculum (rudder) and processus vaginalis.

Male External Genitalia The external genitalia develop from weeks 4 to 12 of gestation. They begin as a genital tubercle at the cranial end of the cloacal membrane, flanked on either side by the labioscrotal swellings and urogenital folds. The future of the labioscrotal swellings and urogenital folds in the 46XX and 46XY fetuses are shown in Table 1.1. There are many gene loci, in addition to the SRY gene, needed for successful male differentiation. The urogenital ridge is driven by Wt1, Sf1, and Dax1 [17]. Sertoli cell production of AMH depends on Sox9, FGF9, SRY, Fog2, and Igf1 [18–21]. Finally, Leydig cell production of testosterone is influenced by enzymes produced by the following genes: StAR, Cyp11a1, Cyp17, and 3beta-HSD [22]. The urogenital sinus and genital tubercle produce 5-alpha reductase, so that under the influence of testosterone

Chapter 1: Anatomy and Embryology

and dihydrotestosterone, the distance between the phallus and the anus increases and the genital tubercle elongates into the primordial phallus and then the penis. There is some debate regarding male urethral development. The classic fusion theory, proposed by Glenister in 1954, maintains that the urethral plate canalizes on the ventral aspect of the primordial phallus using the urogenital sinus endoderm. The urethral groove folds and fuses to form the spongy urethra, which then meets an ingrowth of the surface ectoderm distally at the glans penis [23]. However, this theory does not explain the wide variety of anomalies seen in men with hypospadias. Hadidi et al. [24] conducted a histological study of 15 male fetuses, ranging from 6 to 14 weeks of development, and found the glandular urethra to develop in parallel to the spongy urethra. They observed three stages of development within the spongy urethra: a solid epithelial plate, a deep urethral groove, and a fused urethra. The glandular urethra appeared to have four stages of development: a solid epithelial plate, a blind central canal, a deep glandular groove, and distal migration of the glans/ prepuce, which fuses with the distal-most edges of the glandular groove to form the floor of the glandular urethra. The authors did not find evidence of ectodermal ingrowth, as previously suggested by Glenister. Using optical projection tomography, Li et al. [25] proposed a “double zipper theory,” where the first (opening) zipper begins at the scrotal urethral meatus and is characterized by high levels of cellular proliferation within the dorsal aspect of the urethral plate, allowing for urethral plate canalization. The second (closing) zipper is represented by midline fusion of the epithelial edges lining the urethral groove, which also moves distally from the scrotum to the terminal urethral meatus at the mid-glans. The authors noted a lack of caspase 3 within their histological analysis, which would have suggested a role for apoptosis or ectodermal ingrowth, as previously suggested.

The corpus cavernosum and corpus spongiosum develop from the mesenchyme within the primordial phallus. In the male, the paired labioscrotal swellings meet in the midline and fuse to form the scrotum. The future prepuce forms simultaneous with the glandular urethra and is dependent on normal urethral development. This is evidenced by the lack of a normal ventral prepuce in hypospadias and the lack of a normal dorsal prepuce in epispadias.

Key Points • The male urethra likely develops via a “double zipper” process, where the opening zipper is represented by urethral canalization, followed by midline fusion (the closing zipper), moving the meatus from the proximal phallus to the distal glans. • Dihydrotestosterone directs the formation of the external male genitalia from the ambiguous genital tubercle. The urogenital folds and labioscrotal swellings fuse to form the ventral penis and scrotum, respectively. In the female, these structures become the labia minor and labia majora, respectively.

Conclusion Embryology of the genital system is intimately associated with the urinary tract. Greatly impacted by chromosomal makeup, cellular proliferation and apoptosis, correct timing of events, and reciprocal differentiation, there are many opportunities for missteps and congenital anomalies. Although the majority of these malformations are not fatal, they do have lasting impacts on the infant into adulthood. An understanding of how normal anatomy is formed is essential to a solid foundation in male infertility diagnosis, evaluation, and management.

Further Reading

References

Wein AJ, Kavoussi LR, Novick AC, Partin AW, Peters CA (eds). Campbell-Walsh Urology, 10th ed. Saint Louis, MO: Elsevier, 2011.

1.

Moore KL, Persaud TVN, Torchia MG. The Developing Human: Clinically Oriented Embryology, 11th ed. Philadelphia, PA: Saunders, 2020.

Moore KL, Persaud TVN, Torchia MG. The Developing Human: Clinically Oriented Embryology, 11th ed. Philadelphia, PA: Saunders, 2020.

2.

Ochsner MG, Brannan W, Goodier EH. Absent vas deferens associated with renal agenesis. JAMA 1972;222:1055–6.

3.

McCallum T, Milunsky J, Munarriz R, Carson R, Sadeghi-Nejad H, Oates R. Unilateral renal agenesis associated with congenital bilateral absence of the vas deferens: phenotypic findings and genetic considerations. Hum Reprod 2001;16:282–8.

4.

Holt SA, Peterson NE. Ectopia of seminal vesicle. Associated with

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5.

6.

7.

8.

9.

agenesis of ipsilateral kidney. Urology 1974;4:322–4. Wein AJ, Kavoussi LR, Novick AC, Partin AW, Peters CA (eds). Campbell-Walsh Urology, 10th ed. Saint Louis, MO: Elsevier, 2011. Nievelstein RA, van der Werff JF, Verbeek FJ, Valk J, Vermeij-Keers C. Normal and abnormal embryonic development of the anorectum in human embryos. Teratology 1998;57:70–8. Nebot-Cegarra J, Fàbregas PJ, Sánchez-Pérez I. Cellular proliferation in the urorectal septation complex of the human embryo at Carnegie stages 13–18: a nuclear area-based morphometric analysis. J Anat 2005;207:353–64. Liaw A, Cunha GR, Shen J, et al. Development of the human bladder and ureterovesical junction. Differentiation 2018;103:66–73. Viana R, Batourina E, Huang H, et al. The development of the bladder trigone, the center of the anti-reflux mechanism. Development 2007;134:3763–9.

human semen and the formulation of a semen simulant. J Androl 2005;26:459–69. 13. Huang L, Pu Y, Alam S, Birch L, Prins GS. Estrogenic regulation of signaling pathways and homeobox genes during rat prostate development. J Androl 2004;25:330–7. 14. Prins GS, Tang WY, Belmonte J, Ho SM. Perinatal exposure to oestradiol and bisphenol A alters the prostate epigenome and increases susceptibility to carcinogenesis. Basic Clin Pharmacol Toxicol 2008;102:134–8. 15. Rafailidis V, Varelas S, Apostolopoulou F, Rafailidis D. Nonobliteration of the processus vaginalis. J Ultrasound Med 2016;35:805–18. 16. Kolon TF, Herndon CD, Baker LA, et al. Evaluation and treatment of cryptorchidism: AUA guideline. J Urol 2014;192:337–45.

10. Brewster SF. The development and differentiation of human seminal vesicles. J Anat 1985;143:45–55.

17. Nachtigal MW, Hirokawa Y, Enyeart-VanHouten DL, Flanagan JN, Hammer GD, Ingraham HA. Wilms’ tumor 1 and Dax-1 modulate the orphan nuclear receptor SF-1 in sex-specific gene expression. Cell 1998;93:445–54.

11. Thomson AA, Marker PC. Branching morphogenesis in the prostate gland and seminal vesicles. Differentiation 2006;74:382–92. 12. Owen DH, Katz DF. A review of the physical and chemical properties of

18. Sekido R, Bar I, Narváez V, Penny G, Lovell-Badge R. SOX9 is upregulated by the transient expression of SRY specifically in Sertoli cell precursors. Dev Biol 2004;274: 271–9.

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19. Colvin JS, Green RP, Schmahl J, Capel B, Ornitz DM. Male-to-female sex reversal in mice lacking fibroblast growth factor 9. Cell 2001;104:875–89. 20. Tevosian SG, Albrecht KH, Crispino JD, Fujiwara Y, Eicher EM, Orkin SH. Gonadal differentiation, sex determination and normal Sry expression in mice require direct interaction between transcription partners GATA4 and FOG2. Development 2002;129:4627–34. 21. Nef S, Verma-Kurvari S, Merenmies J, et al. Testis determination requires insulin receptor family function in mice. Nature 2003;426:291–5. 22. Brennan J, Tilmann C, Capel B. Pdgfr-α mediates testis cord organization and fetal Leydig cell development in the XY gonad. Genes Dev 2003;17:800–10. 23. Glenister TW. The origin and fate of the urethral plate in man. J Anat 1954;88:413–25. 24. Hadidi AT, Roessler J, Coerdt W. Development of the human male urethra: a histochemical study on human embryos. J Pediatr Surg 2014;49:1146–52. 25. Li Y, Sinclair A, Cao M, et al. Canalization of the urethral plate precedes fusion of the urethral folds during male penile urethral development: the double zipper hypothesis. J Urol 2015;193: 1353–9.

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Cellular Architecture and Function of the Testis Siwen Wu, Lingling Wang, and C. Yan Cheng

Introduction Cellular Architecture of the Testis The cellular architecture of the mammalian testis that supports testis function, which, in turn, maintains spermatogenesis throughout adulthood to produce millions of sperm on a daily basis from rodents to humans, has been eminently reviewed by investigators in recent years, based on morphological, biochemical, and molecular studies [1– 10]. As such, in order to avoid a repetitive account on this topic, compared with earlier reviews, we attempt to provide insightful information on this topic based on recent studies which have not been evaluated in details, as noted in the following sections. Thus, this avoids redundancy because readers can refer to the earlier reviews pertinent to the cellular architecture of the testis that supports spermatogenesis as reviewed in [1–10]. It is conceivable that some necessary discussion still overlap with earlier reviews or earlier editions of this reference work. This is done such that readers can follow through the discussion here without the necessity of going through other contents to grasp related facts and concepts. Nonetheless, we will refer to other reviews for additional discussion on specific topics in this chapter when a detailed and redundant description is not warranted. In brief, the general cellular architecture of the mammalian testis is shown in Fig. 2.1, using the adult rat as an example. As noted in Fig. 2.1A, this is the schematic drawing of the cross-section of a stage VII seminiferous tubule from an adult rat testis, illustrating the spatial relationship between the Sertoli cells and germ cells in the seminiferous epithelium, with the support of the basement membrane of the tunica propria. These structures all work in concert to support spermatogenesis through different epithelial cycles, as noted in both rodents and humans (Figs 2.2, 2.3, and 2.4). In the seminiferous epithelium of a tubule, which is the functional unit that produces millions of sperm daily after puberty in humans (at approximately 12 years of age) or in rodents (approximately 35 days or 45 days of age in mice or

rats) [5, 8, 11–13]. The most notable feature is the testisspecific and actin-rich adherens junction (AJ) at the Sertoli cell–cell and Sertoli–spermatid (steps 8–16 or 8–19 in the mouse or rat testis) interface designated basal ectoplasmic specialization (ES) and apical ES, respectively [14–18] (Fig. 2.1A). More importantly, studies have shown that the orderly alignment of developing spermatids, most notably step 17–19 spermatids in stage IV–VIII tubules, has distinctive apico-basal polarity wherein developing spermatids are orderly aligned, with their heads pointing toward the basement membrane and their tails toward the tubule lumen, as noted in Fig. 2.1B. Studies have shown that this orderly alignment of developing spermatids is supported by three distinctive cell polarity complexes, including the Par (Partitioning defective), Crumbs-, and Scribblebased polarity modules [19]. It is noted that these three cell polarity protein modules, including their partner proteins that were initially found in fruit flies and worms [20, 21], have now been identified in the testis. These include: the Par-based polarity module of Par3/Par5/Par6 and their protein partners, including atypical protein kinase C (aPKC) and Cdc42 [22, 23]; the Crumbs-based polarity module of Crumbs homolog-3 (Crb3) and its protein partners of Protein associated with Lin-7 1 (Pals1) and Pals1associated tight junction protein (PatJ) [24]; and the Scribble-based polarity module of Scribble and its protein partners of Lethal giant larvae 2 (Lgl2) and Discs large 1 (Dlg1) [25]. Many of these proteins have been identified in the testis, and their physiological function to support cell polarity in the testis during spermatogenesis have also been partially characterized. More importantly, the orderly alignment of polarized spermatids across the plane of the seminiferous epithelium, mimicking planar cell polarity (PCP), has also been characterized, based on the use of confocal microscopy, as reported earlier [26, 27]. This is analogous to PCP found in other epithelia such as hair cells in the cochlea and also in insects hair cuticle cells [28–32]. Studies have also shown that Sertoli and/or germ cells indeed express the PCP protein complexes Van

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Fig. 2.1 Morphological features of the seminiferous epithelium in rat testes that support spermatogenesis. (A) Schematic drawing of a cross-section of a typical stage VII seminiferous tubule that illustrates the most typical morphological features across the seminiferous epithelium in adult rat testes. The blood–testis barrier (BTB), constituted by the actin-based tight junction, basal ectoplasmic specialization (ES), and gap junction, as well as

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Gogh-like 2 (Vangl2)/Prickle 1 [26, 33] and Frizzled (Fzd)/ Disheveled 3 (Dvl3) [34] that support spermatid PCP. In brief, this orderly alignment of spermatids across the seminiferous epithelium supported by cell polarity proteins and PCP proteins is essential to maintain spermatogenesis. They are also necessary to maintain daily germ cell production, since cross-talks and spatial interaction between Sertoli cells and germ cells (most notably spermatids during spermiogenesis) mimic a well-run manufacturing plant. Thus, components necessary to support morphological transformation of developing germ cells, such as proteins across the developing germ cells, can be “installed” or “removed” (either for recycling or for degradation) at different stages of the epithelial cycle (Figs 2.2, 2.3, and 2.4). This concept has now been confirmed in studies by selective knockdown (KD) of these cell polarity or PCP proteins by RNA interference (RNAi) using corresponding siRNA duplexes, which was found to perturb spermatid polarity, causing defects in spermatogenesis [22, 24–26, 34]. Collectively, these finding thus illustrate the physiological significance of these cell polarity and PCP proteins to support spermatogenesis.

Key Points • Developing spermatids and Sertoli cells display apico-basal polarity across the seminiferous epithelium conferred by cell polarity protein complexes (or modules). • Developing spermatids, in particular step 17–19 spermatids in stage V to VIII tubules, display apico-basal polarity, but also planar cell polarity (PCP). • PCP refers to the alignment of polarized step 17–19 spermatids displaying apico-basal polarity across the plane of the seminiferous epithelium conferred by PCP proteins.

Seminiferous Tubules and the Epithelial Cycle of Spermatogenesis In the mammalian testis, there are unique, but cyclic, changes in the association of Sertoli cells with specific classes of developing germ cells, most notably haploid spermatids, which can be divided into stages from I to XII, I to XIV, and I to XII in the mouse, rat, and human testis, respectively [5, 8, 35, 36]. The stages of the epithelial cycle of spermatogenesis corresponding to the rat, mouse, and human are depicted in Figs 2.2, 2.3, and 2.4, respectively. In humans, spermatogonia type B first appear at stage I and preleptotene spermatocytes at stages V and VI, and elongating/elongated spermatids begin their journey being transported to the adluminal edge of the tubule lumen from stages VIII to VI, so that the release of fully developed spermatids (i.e., spermatozoa) at spermiation takes place in late stage VI of the epithelial cycle (Fig. 2.4). Meiosis takes place at stage XII of the epithelial cycle [13] (Fig. 2.4). It is noted that the current epithelial cycle scheme in humans (Fig. 2.4) is a revised cycle staging of I–VI, as earlier reported [12, 37]. In rat testes, spermatogonia type B appear at stage V, and preleptotene spermatocytes at stage VII, which are to be transported across the immunological barrier at the blood–testis barrier (BTB) at stage VII to late stage VIII/early stage IX (Fig. 2.2). On the other hand, elongating/elongated spermatids are being transported to the adluminal edge of the tubule from stages VI to early stage VIII, so that the release of sperm at spermiation can take place at late stage VIII, with meiosis taking place at stage XIV [38–40] (Fig. 2.3). In brief, when a specific section of a tubule is examined by stereomicroscopy, the duration of a seminiferous epithelial cycle (e.g., sperm release at spermiation at stage VIII in rats or humans at stage VI) represents the time it takes to complete the series of morphological changes between two appearances of the same developmental stage. Thus, the duration of the seminiferous epithelium cycle is estimated

Fig. 2.1 (cont.) by the intermediate filament-based desmosome, segregates the seminiferous epithelium into the basal and adluminal (apical) compartments. The cytoskeletons, shown here are the actin- and microtubule (MT)-based polarized cytoskeletons with distinctive plus (+) and minus (
) ends, which provide the structural support to Sertoli cells, but also to developing germ cells. Preleptotene spermatocytes transformed from type B spermatogonia residing in the basal compartment are to be transported across the BTB, beginning in late stage VII through to late stage VIII of the epithelial cycle, so that they can be differentiated into leptotene, zygotene, pachytene, and eventually diplotene spermatocytes to undergo meiosis I/II to form haploid spermatids. Besides preleptotene spermatocytes, developing spermatids and other germ cells, as well as organelles and cellular structures (e.g., residual bodies, phagosomes, endocytic vesicles), are also to be transported across the seminiferous epithelium to support spermatogenesis. These transports are supported by MT-based motor proteins, such as dynein 1 (an MT minus (
) end-directed motor protein) and kinesins (e.g., kinesin 15, an MT plus (+) end-directed motor protein), myosin VI (an actin pointed (
) end-directed motor protein), and myosin VIIa (an actin barbed (+) end-directed motor protein), as noted herein. Apical ES is found at the Sertoli–spermatid (steps 8–19 in rat testes) interface, whereas basal ES is at the Sertoli cell–cell interface and coexists with the tight junction and gap junction to support BTB function. (B) Cross-section of a stage VII tubule in adult rat testes showing different germ cell types which are supported by Sertoli cells. The morphological features noted in rodent testes are similar to those found in human testes. Scale bar, 40 μm.

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Fig, 2.2 Spermatogenesis in rats. (A) The various steps of spermatogenesis in male rats. Asingle (As) is considered to be a spermatogonial stem cell (SSC) [126]. Apaired (Apr) and Aaligned (Aal) are considered to be undifferentiated spermatogonia. Aal transform into A1–A4 spermatogonia, which become intermediate spermatogonia (In) and differentiate to become type B (B) spermatogonia. Type B spermatogonia transform to preleptotene (Pl). These germ cells reside in the basal compartment of the seminiferous epithelium and Pl is the only germ cell type that is transported across the blood– testis barrier, while differentiating to become leptotene spermatocyte (L). Once in the adluminal (apical) compartment, L differentiates to become zygotene (Z), pachytene (P), and then diplotene (Di) spermatocytes (tetraploids), which enter meiosis I to form two secondary spermatocytes (Ss) (diploids), and Ss rapidly undergo meiosis II to form the haploid spermatids. In the rat, haploid round spermatids undergo spermiogenesis via 19 steps to become elongated spermatids with extensive morphological changes, including condensation of the genetic materials in the nucleus, formation of the acrosome, and elongation of the tail. The duration of spermatogenesis, that is, from A1 spermatogonia to fully developed step 19 elongated spermatids in stage VIII tubules is about 58 days in the rat [127–129] which was estimated by using [3H]-thymidine-labeled type A spermatogonia. (B) Stages of the seminiferous epithelial cycle are divided into 14 stages in the rat testis, from stage I through to stage XIV based on periodic acid–Schiff (PAS) staining of the spermatid head, and each stage is composed of several distinctive germ cell types. For instance, at stage VIII, only type A1 spermatogonia, preleptotene, and pachytene spermatocytes, as well as step 8 and step 19 spermatids, are found across the seminiferous epithelium. Some notable cellular events are also detected in these stages, including the release of sperm at spermiation which takes place at stage VIII and meiosis I/II which take place at stage XIV. The duration of each stage of the cycle is noted in the bottom panel in hours (hr) and the relative percentages of different staged tubules in cross-sections of rat testes are also shown. In brief, a complete epithelial cycle in the rat testis is about 12.8 days, indicating that if an investigator visualizes a tubule at stage VIII, this will take 12.8 days for the tubule to run through a complete epithelial cycle. Thus, it takes 4.5 epithelial cycles for A1 spermatogonia to become fully developed haploid elongated spermatids, which is equivalent to 58 days. This figure was prepared based on findings of earlier reports [39, 130–133].

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Fig. 2.3 Spermatogenesis in mice. (A) The steps of spermatogenesis in male mice, which are similar to the information noted in the rat testis. However, in the mouse, spermiogenesis comprises only 16 steps. The duration of spermatogenesis in the mouse is about 35 days [134, 135]. (B) In the mouse testis, there are only 12 stages in the epithelial cycle, with a duration of 8.6 days, and the time for each stage is noted in the bottom panel in hours (hr), and the percentage of each stage is also shown. In the mouse testis, it takes four epithelial cycles for A1 spermatogonia to become fully developed haploid elongated spermatids, which is equivalent to 35 days. This figure was prepared based on findings of earlier reports [8, 135, 136].

to be 12.8, 8.6, and 16 days in rat (Fig. 2.2), mouse (Fig. 2.3), and human (Fig. 2.4) testes, which comprises stages I–XIV, I–XII, and I–XII, respectively. However, the duration of spermatogenesis, representing the time it takes for a type As (Asingle) spermatogonium in rats or mice versus Adark spermatogonium in humans to develop into multiple functional haploid spermatids (such as via use of [3H]-thymidine-labeled spermatogonia to track the duration of development), takes 58, 35, and 70 days,

respectively. Thus, this represents 4.5 cycles in rodents (Figs 2.2 and 2.3) and 5 cycles in humans (Fig. 2.4) to complete a round of spermatogenesis because it takes considerable time for undifferentiated spermatogonia (e.g., Apale (Ap) spermatogonia in humans or A1 spermatogonia in rodents) to differentiate into preleptotene spermatocytes in the basal compartment. Furthermore, across the seminiferous epithelium, there is an intricate spatial relationship between developing germ cells and

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Fig. 2.4 Spermatogenesis in humans. (A) In the human testis, the steps of spermatogenesis are also similar to those in rodents, except that Ad (Adark) spermatogonia are considered to be spermatogonial stem cells [137–139], with only 12 steps of haploid spermatids during spermiogenesis. The duration of human spermatogenesis in men is about 64 days [12, 140, 141]. (B) In the human testis, there are 12 stages in the epithelial cycle, with a duration of 16 days from stage I through to stage XII. It takes five epithelial cycles for Ap spermatogonia to become fully developed haploid elongated spermatids, which is equivalent to ~70 days. This figure was prepared based on findings of earlier reports [13].

Sertoli cells at a 30–50:1 ratio [41–43] wherein germ cells rely exclusively on the support of Sertoli cells for their development and nutritional support. This is analogous to a well-designed assembly line of a manufacturing plant, such that components, including proteins and genes, are

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tightly regulated and coordinated – for instance, through the expression of genes and also post-translational processing of proteins, including the addition of newly formed proteins to, and timely removal of, unwanted (e.g., aged) proteins from the developing haploid spermatids, to

Chapter 2: Cellular Architecture

support the daily production of 30–50 million versus 300 million of sperm corresponding to rodents and humans per testes pair on a daily basis.

Key Points • During spermatogenesis in rodents and humans, a distinctive association of developing haploid spermatids with Sertoli cells is noted, so that the stages of the epithelial cycle can be divided into stages I–XII in human and mouse testes, but into stages I–XIV in rat testes. • Developing spermatids across the seminiferous epithelium in the testis during spermiogenesis can be separated into 12, 16, and 19 steps in humans, mice, and rats, respectively.

Cell Polarity, Planar Cell Polarity, and Cell Polarity and Planar Cell Polarity Proteins During spermatogenesis, the most notable cell polarity is the alignment of developing spermatid heads across the seminiferous epithelium, wherein the heads of spermatids at spermiogenesis are pointing toward the basement membrane, with their elongating tails toward the tubule lumen (Fig. 2.1B). Studies have shown that this apico-basal alignment of developing elongating/elongated spermatids across the seminiferous epithelium is mediated through the concerted efforts of the spatiotemporal expression of three cell polarity protein complexes, namely the Par[22, 23], the Crumbs- [24], and the Scribble-based [25] polarity complexes. This also includes the expression of their corresponding partner proteins initially reported in Drosophila or Caenorhabditis elegans of the Par3/Par6/ aPKC/Cdc42 [22], the Crb3/Pals1/PatJ [24], and the Scribble/Lgl2/Dlg1 [25] polarity complexes [20, 44–46]. Studies have shown that these polarity protein complexes display mutually exclusive distribution patterns and distinctive functionality wherein the Par- and Crb3-based complexes are usually localized adjacent to another near the tight junction (TJ) at the apical region, and the Scribble-based complex is found distinctively expressed at the basal region, of an epithelium [20, 21, 47]. Such differences in the distribution of these cell polarity protein complexes thus recruit different partner proteins and cell organelles at the corresponding sites, which, in turn, lead to apico-basal cell polarity. This is essential to support tissue and organ development during embryogenesis. In

the testis, utilizing these apico-basal polarity proteins, developing spermatids can be orderly aligned across the epithelium, enabling easier access of supplies (e.g., nutrients, cytokines, and pertinent biomolecules) from the Sertoli cells, including structural, functional, and nourishment supports, to facilitate spermatid development. Besides the apico-basal alignment of developing spermatids, recent studies have shown that the alignment of directional spermatids across the plane of the seminiferous epithelium also mimics the PCP noted in hair cell distribution in the epidermis and inner ear [48–51]. This additional spermatid polarity has recently been shown to be modulated through the PCP complex Vangl2/Prickle 1 [26, 33] and also Fzd/Dvl3 [34].

Cell Polarity and Regulatory Proteins Studies of cell polarity in the testis are lagging far behind those in other epithelia, since the presence of cell polarity proteins to support spermatid polarity during spermiogenesis was not reported until 2008 when the Par-based protein complex (e.g., Par3, Par6) and its partner proteins (e.g., aPKC, Cdc42) were first reported in the testis [22]. Since then, Par1 [23], Scribble (and its partner proteins Lgl2 and Dlg1) [25], and Crb3 (and its partner proteins Pals1 and PatJ) [24] have been identified and studied in the testis. These studies have shown that all three cell polarity protein complexes that confer apico-basal polarity to spermatids, to support the orientation of the spermatid heads by aligning perpendicularly to the basement membrane, exert their effects through changes in the cytoskeletal organization of actin, consistent with reports in the literature in other model systems [52]. Furthermore, the physiological function of the Par- and Crb3-based polarity complexes is somewhat different from the Scribble-based polarity complex. For instance, a knockdown of either Par3 (or Par6) and Crb3 by RNAi perturbs Sertoli cell TJ barrier function, making the Sertoli cell BTB barrier “leaky” [22, 24]. On the other hand, a triple knockdown of Scribble/ Lgl2/Dlg1 promotes TJ barrier function, making the immunological barrier “tighter” [25]. Collectively, these findings illustrate that their functions in the testis are mutually exclusive, consistently with their functional role in other epithelia/endothelia [20, 21, 47]. More importantly, studies have shown that these cell polarity proteins also exert their regulatory effects through changes in the cytoskeletal organization of F-actin across the seminiferous epithelium (Fig. 2.5). For instance, a triple knockdown of Scribble/Lgl2/Dlg1 in the testis in vivo by RNAi, which were shown to promote the TJ barrier function in the

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Fig. 2.5 Localization of actin- and vimentin-based cytoskeletons across the seminiferous epithelium in the adult rat testis. F-actin (red fluorescence) and the two actin regulatory proteins Eps8 (green fluorescence) and Arp3 (green fluorescence), as well as vimentin (green fluorescence), were visualized

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Sertoli cell epithelium cultured in vitro, was also found to promote the integrity of F-actin network at the BTB in stage VIII tubules in vivo [25]. Interestingly, this is also the stage wherein the BTB undergoes remodeling to facilitate the transport of preleptotene spermatocytes across the barrier site in control testes (treated with non-targeting small interfering ribonucleic acid (siRNA) duplexes) [25]. Thus, this shows that a knockdown of the Scribble complex promotes BTB integrity by maintaining the F-actin network at the site even in late stage VIII tubules when F-actin should have undergone remodeling to facilitate the transport of preleptotene spermatocytes across the barrier. As such, occludin was shown to be retained at the site when it should have been downregulated in stage VIII tubules to facilitate BTB remodeling [25]. On the other hand, a knockdown of Crb3 in the testis in vivo was found to promote BTB remodeling, perturbing F-actin organization at the BTB, but also across the seminiferous epithelium, leading to premature release of spermatids in stage V–VII tubules [24]. This finding is consistent with the observation in vitro that a knockdown of Crb3 in the Sertoli cell epithelium indeed induced extensive, but reversible, truncation of F-actin network across the Sertoli cell cytosol [24]. Taking these findings collectively, it is obvious that cell polarity proteins are working in concert with cell cytoskeletons to confer polarity function in the testis through their effects on cytoskeletal organization.

Key Points • The three cell polarity proteins that support apicobasal proteins are the Par-, Crb3-, and Scribblebased protein complexes (modules), which, in turn, confer Sertoli cell and spermatid polarity. • The Par- and Crb3-based complexes work in concert with each other, but are mutually exclusive with respect to the Scribble-based complex regarding their localization and also functionality.

Planar Cell Polarity Studies in the testis have shown that multiple PCP proteins, including Vangl2 (a homolog of Van Gogh) [26, 33] and Dvl3 [34], expressed both by Sertoli and germ cells in adult rat testes, are essential to support spermatid PCP. In fact, virtually all PCP proteins necessary to maintain PCP earlier found in insects (e.g., Drosophila) are expressed in the testis. These include: PCP core proteins (e.g., Vangl2, Dvl2, Dvl3, Fzd), PCP ligands (e.g., Wnt5A), PCP effectors (e.g., Fuzzy), and PCP signaling proteins (e.g., Dchs1), via the use of specific primers for reverse transcriptase polymerase chain reaction (RT-PCR) and also immunologicallybased staining [33, 34]. Interestingly, the two PCP proteins that have been carefully examined in the testis, namely Vangl2 (also known as Strabismus, or FlyBase, first found in Drosophila [53]) and Dvl3, are also localized across the seminiferous epithelium through different stages of the epithelial cycle, co-localizing with microtubules (MTs) and appearing as track-like structures that stretch across the entire seminiferous epithelium [33, 34]. More importantly, a knockdown of Vangl2 by RNAi in the testis in vivo was found to perturb spermatid PCP considerably, based on the use of confocal microscopy, by re-constructing the seminiferous epithelial architecture, revealing the notable loss of spermatid PCP across the epithelium [26]. These changes, in turn, impede the status of spermatogenesis, in particular spermiogenesis [33]. This observation is important because it illustrates that Vangl2 supports spermatid PCP in the testis. Furthermore, these studies also illustrate that Vangl2 exerts its regulatory effects in the testis through actinand MT-based cytoskeletons [26]. Consistent with these findings, Dvl3, another PCP protein that works in concert with Fzd, also exerts its regulatory effects in Sertoli cells to support PCP in the testis through actin- and MTbased cytoskeletons [34]. As such, a knockdown of Dvl3 or Dvl1 or Dvl2 in the Sertoli cell epithelium, using corresponding specific siRNA duplexes with an

Fig. 2.5 (cont.) in cross-sections of adult rat testes by immunofluorescence microscopy, as earlier described [112, 142]. F-actin was most prominently localized at the apical and basal ectoplasmic specialization (ES)/blood–testis barrier (BTB) site. F-actin dynamics are supported functionally by the actin barbed end capping and bundling protein Eps8, but also by the branched actin polymerization protein Arp3. In brief, Eps8 (and other actin bundling proteins such as palladin and fascin 1) confer actin filaments in the seminiferous epithelium as bundles to support ES function. On the other hand, Arp3 that binds to Arp2 to form the Arp2/3 complex induces branched actin polymerization and can modify linear actin filaments into a branched network. In brief, the concerted effects of Eps8 and Arp2/3 complex confer actin network the plasticity to assume a bundled or unbundled/branched configuration to support cellular changes across the epithelium during the epithelial cycle. F-actin, Eps8, and Arp3 all appeared as bulb-like structures, localized at the concave (ventral) side of spermatid heads at the apical ES. Vimentin is the structural component of the intermediate filaments across the epithelium, most notably near the base of the seminiferous epithelium as short, track-like structures to support cellular functions. Scale bar, 250 µm; enlarged image enclosed in yellow box, 80 µm; magnified image in inset, 30 µm. Cell nuclei visualized by 40 ,6-diamidino-2-phenylindole (DAPI) staining.

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established functional barrier by RNAi, was found to perturb the TJ permeability barrier function [34]. However, a knockdown of Dvl3 alone was more effective in perturbing the Sertoli cell TJ barrier function than either Dvl1 or Dvl2 alone [34]. More importantly, Dvl3 in the testis exerts its effects by maintaining the integrity of actin filaments and also MTs across the Sertoli cell cytosol. This notion was supported by findings that Dvl3 knockdown in Sertoli cells cultured in vitro was able to induce extensive truncation of actin and MT filaments, but also perturbed actin and MT polymerization activity in Sertoli cells, based on biochemical assays [34]. These data are consistent with studies investigating the role of PCP proteins on cytoskeletal organization in other study models [54, 55]. However, much work is needed to explore the role of other PCP proteins in the testis, in particular Prickles (which work closely with Vangl2) and Fzd and inversin (which works closely with Dvl3), as noted in studies of other epithelia [56, 57]. Furthermore, it is also important to examine how PCP proteins are working in concert with cell polarity protein complexes to confer spermatid polarity, to support spermatogenesis through actin and MT cytoskeletons. In this context, it is of interest to note that based on studies of other epithelia, the Vangl2/Prickle and Fzd/ Dvl–diversin–inversin complexes are two important PCP regulatory protein complexes. In these protein complexes, Vangl2 and Fzd are integral membrane proteins that partner with the corresponding cytoplasmic multifunctional adaptor proteins Prickle 1/2/3 and Dvl/diversin/inversin (known as Diego, dgo, in Drosophila) to create unique functional PCP complex modules [54, 55, 58, 59]. On the other hand, Celsr (cadherin EGF LAG seven-pass G-type receptor; its Drosophila counterpart is Flamingo (Fmi), also known as Starry night (Stan), which is a family of atypical cadherins) 1, 2, or 3 are members of a family of integral membrane PCP proteins and the third PCP protein complex [60, 61]. However, the role of Celsr in conferring PCP in the testis has not been examined. Emerging evidence has shown that the Fzd/ Dvl/inversin complex exerts its effects to modulate actin cytoskeleton, wherein Dvl promotes actin polymerization by activating Rho GTPase and Rho-associated kinase (ROK), which, however, can be inhibited by PCP protein Multiple wing hairs (Mwh) [62]. Interestingly, Mwh is also the downstream PCP protein of the Vangl2/Prickle complex [62, 63], but its ortholog in rodents and humans remains to be identified. Taken collectively, these findings illustrate that the effects of Vangl2/Prickle and Fzd/

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Dvl/diversin/inversin on actin cytoskeletal function are mutually exclusive. On the other hand, numerous diseases are found in humans following mutations and/or genetic variations of these PCP proteins [64]. For instance, genetic variations of Dvl1 and/or Dvl3, via clustered frameshift of the last axon and also other genetic variations of partner proteins of Wnt signaling (e.g., Ror2, Wnt5a) were found to mediate autosomal dominant Robinow syndrome, displaying cardiac abnormalities in approximately 75% of patients with Dvl3 genetic mutations [65, 66]. Robinow syndrome is a rare pathological condition that affects the development of many parts of the body in humans, including the skeleton and also underdeveloped genitalia in males and females, plus dental problems with crowded teeth and gum overgrowth [67]. On the other hand, Vangl2 mutations in humans lead to cranial neural tube defects [68].

Key Points • Vangl2/Prickle and Fzd/Disheveled/diversin/ inversin are the two PCP protein complexes that have been studied in the testis. • These PCP proteins confer alignment of directionally oriented developing spermatids across the plane of the seminiferous epithelium during the epithelial cycle, so that developing haploid spermatids can be aligned properly within the limited space of the seminiferous epithelium in seminiferous tubules to support spermatogenesis.

A Local Regulatory Axis to Support Changes of Cellular Architecture during the Epithelial Cycle There are dynamic interactions that take place between Sertoli cells, but also between Sertoli and germ cells, at the cellular and molecular levels in the seminiferous epithelium to support different cascades of cellular events during the epithelial cycle such as spermatogonial differentiaton, meiosis, spermiogenesis, and spermiation [2, 69]. However, the cellular architecture across the epithelium undergoes continuous remodeling due to structural re-organization of actin- (Fig. 2.5), vimentin- (Fig. 2.5), and MT-based (Fig. 2.6) cytoskeletons through different stages of the epithelial cycle, analogous to a modern-day manufacturing plant. In order for this to occur, different sets of cellular functions will need to be tightly

Chapter 2: Cellular Architecture

Fig. 2.6 Localization of the microtubule (MT) cytoskeleton across the seminiferous epithelium in the adult rat testis. MTs were visualized by staining of α-tubulin (which, together with β-tubulin, creates α-/β-tubulin oligomers, which are the building blocks of MTs) (green fluorescence). MTs are supported by MARK4 (a Ser/Thr non-receptor protein kinase, by phosphorylating microtubule affinity proteins [MAPs] [such as MAP1a] which bind onto MTs to stabilize the MT filaments, the phosphorylated MAPs are then detached from MTs, destabilizing the cytoskeleton to undergo catastrophe). MTs are also supported by EB1, a microtubule plus (+) end tracking protein (+TIP) known to stabilize MTs. It is noted that both MT regulatory proteins co-localize with MTs. Scale bar, 250 µm and 80 µm. Cell nuclei visualized by 40 ,6-diamidino-2-phenylindole (DAPI) staining.

coordinated, so that the manufactured parts (i.e., organelles such as phagosomes, residual bodies, and Golgi apparatus; cellular components such as constituents of germ cell plasma membrane and the acrosome, genetic material of the germ cell nucleus) can be transported (via endocytic vesicles) across different assembly lines using

proper conveyer belts (such as MT and/or F-actin conferred polarized tracks) for the assembly of germ cells at different stages of their development [1, 3]. Studies have shown that the seminiferous epithelium produces several regulatory peptides locally that are used to modulate and coordinate cellular functions across the epithelium

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through changes in cytoskeletal organization. These peptides, in turn, support cellular functions pertinent to the epithelial cycle and to maintaining the cellular architecture of the epithelium.

The F5-Peptide Studies in different epithelia have shown that some peptides generated from laminin chains via proteolytic cleavage endogenously, almost exclusively in the extracellular matrix (ECM) such as basal lamina (known as basement membrane in the testis, located at the base of the seminiferous epithelium), are biologically active molecules that modulate a wide range of cellular events. These include angiogenesis, cell adhesion, junction assembly and/or disassembly, differentiation, cell movement and cell apoptosis. In the mammalian testis, most of the laminin chains, such as laminin-α2 chains, similar to other epithelia, are found at the basement membrane, which is a modified form of extracellular matrix (ECM) in the testis [70, 71]. Unlike other epithelia, laminin-γ3 chain is exclusively expressed at the Sertoli–elongating/elongated spermatid interface, known as the apical ES, a testis-specific and actin-rich cell–cell AJ [72, 73], located at the opposite end of the basement membrane. It is noted that a functional laminin-based ligand is a trimeric structure composed of three laminin chains, one each of α, β, and γ chains. Studies have shown that the functional trimeric ligand at the apical ES is composed of laminin-γ3, -α3, and -β3, and designated laminin-333 [73, 74]. In most epithelia, laminins serve as the ligand that specifically interacts with an integrin-based receptor to induce signaling function. Interestingly, laminin-333, exclusively expressed by elongating/elongated spermatids in the adult rat testis [73, 74], forms a bona fide adhesion complex with α6β1 integrin, which is expressed by Sertoli cells [75– 77]. In late stage VIII of the epithelial cycle, matrix metalloprotease 2 (MMP2) is robustly expressed at the apical ES to induce degradation of the apical ES [74], which is capable of generating a short stretch of a peptide designated the F5-peptide via proteolytic cleavage of the laminin-γ3 chain at its domain IV [73, 78] (Fig. 2.7). Studies have shown that the F5-peptide is composed of 50 amino acid residues, capable of inducing remodeling of the Sertoli cell BTB in vitro and in vivo, by making the immunological barrier “leaky” transiently, through the use of its purified recombinant protein or via its overexpression in the Sertoli cell epithelium in vitro or in the testis in vivo [78, 79]. This transient BTB “opening” thus facilitates the transport of preleptotene

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spermatocytes across the immunological barrier at stage VII-VIII of the epithelial cycle. Besides these effects on the BTB, the F5-peptide is also capable of inducing apical ES degeneration, illustrating an initial breakdown of the apical ES that facilitates the release of mature spermatids (i.e., spermatozoa) from the apical ES site at spermiation. Also, this generation of the F5-peptide also potentiates the cellular events of apical ES break-down to facilitate spermiation [79] (Fig. 2.7). These findings thus illustrate that the testis generates an endogenous bioactive peptide during spermiation to coordinate the cellular events of spermiation and BTB remodeling that take place at the opposite ends of the seminiferous epithelium in stage VIII tubules. Furthermore, studies have shown that the F5-peptide exerts its effects through the signaling protein FAK downstream, by downregulating, and also causing disruptive spatial expression of, p-FAK-Y407 across the seminiferous epithelium [78, 79] (Fig. 2.8). Earlier studies using different mutants of p-FAK-Y407, including the phosphomimetic mutants of p-FAK-Y407E (the constitutively active mutant) and p-FAK-Y407F (the constitutively inactive mutant), cloned into the mammalian expression vector pCI-neo for their overexpression in Sertoli cells cultured in vitro with an established functional TJ barrier, have shown that p-FAK-Y407 is a crucial regulator of the Sertoli cell BTB by promoting BTB integrity [80]. The BTB-promoting effects of p-FAK-Y407 have also confirmed, using the human Sertoli cell BTB model in vitro, that overexpression of the human p-FAK-Y407E phosphomimetic (i.e., constitutively active) mutant in primary human Sertoli cells was capable of rescuing PFOS-induced Sertoli cell injury by blocking PFOS-mediated disruptive effects on the Sertoli cell TJ permeability barrier function [81].

The NC1-Peptide Background One of the major constituents of the basement membrane, a modified form of the ECM [70, 71, 82], at the base of the seminiferous epithelium is collagen α3(IV) chain. In this context, it is of interest to note that collagens and laminins are the predominant structural proteins and building blocks of the ECM [70, 83]. The building block of the collagen IV network (e.g., collagen α3(IV)) in the basement membrane of the seminiferous tubule (Fig. 2.3) is a triple helical structure (i.e., a monomer) composed of three α chains. There are six genetically distinct α chains, designated α1 to α6, known to date [84, 85]. Each monomer is characterized by a non-collagenous 7S domain of

Chapter 2: Cellular Architecture

Fig. 2.7 A hypothetical model in which bioactive peptides work in concert with the corresponding motor proteins to support transport of spermatids and preleptotene spermatocytes across the seminiferous epithelium and blood–testis barrier (BTB), respectively, during spermatogenesis.

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1

2

3

Laminin-J3 chain

Collagen-D3 (IV) chain

Laminin-D2 chain

MMP-2-mediated degradation

MMP-9-mediated degradation

F5-peptide

MMP-9-mediated degradation LG3/4/5-peptide

NC1-peptide

 p-FAK-Y407

mTOR Raptor

mTORC1

mTORC1





p-rpS6

N-WASP

p-rpS6





p-Akt1/2

Arp2/3 complex

mTOR Raptor

p-Akt1/2

 Cdc42 GTPase

Promote BTB/basal ES and apical ES re-structuring and degeneration



EB1

Promote BTB/basal ES and apical ES integrity and re-assembly



Endocytic vesicle-mediated protein trafficking

Eps8

Promote BTB/basal ES and apical ES restructuring and degeneration Fig. 2.8 The signaling molecules and pathways used by the three bioactive peptides to regulate spermatogenesis, using the rat testis as a study model.

15 amino acid residues from the N-terminus, a long middle collagenous domain of approximately 1400 residues composed of the Gly–Xaa–Yaa repeats, and a Cterminal non-collagenous (NC1) domain of approximately 230 residues. At least 19 different collagen subtypes are known to date [86, 87], and they can be classified into the following subgroups. These include: (1) fibrillar collagens (e.g., types I, II, III, V, and XI, all of which have long

triple helical structures), (2) network-forming collagens (e.g., types IV, VIII, and X, which have interrupted domains in the triple helix), (3) beaded filament-forming collagens (e.g., type VI), (4) anchoring fibril-forming collagens (e.g., type VII), (5) fibril-associated collagens (e.g., types IX, XII, XIV, XVI, and XIX), and (6) transmembrane collagens (e.g., types XIII and XVII) [86]. While the collagen V and XVIII subtypes are not part of these

Fig. 2.7 (cont.) F5- and NC1-peptide generated at the apical ectoplasmic specialization (ES) and basement membrane, respectively, are used to support remodeling of the apical ES and also basal ES/BTB at stages VII–VIII of the epithelial cycle to prepare for the release of sperm at spermiation and preleptotene spermatocyte transport at the BTB. On the other hand, LG3/4/5-peptide generated at the basement membrane is used to support apical ES and basal ES/BTB integrity in early stage VII tubules, but also to facilitate the re-assembly of a new BTB underneath the preleptotene spermatocytes when they are transported across the BTB to maintain the immunological barrier integrity (see text for details).

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Chapter 2: Cellular Architecture

subgroups, they also serve as structural components in some basement membranes [88]. Interestingly, the trimeric collagen monomers interact and associate with one another to form dimers and tetramers. For instance, the C-terminal NC1 domains of monomers can selfassociate with others to create dimers, whereas the Nterminal 7S domains also associate with others to form spider-like tetramers [85]. These dimers and tetramers, in turn, create a suprastructure of collagen network. Type IV Collagens in the Testis Collagen α1(IV) and α2(IV) chains are ubiquitously found in mammalian tissues, whereas collagen α3(IV), α4(IV), and α5(IV) chains have a more restrictive tissue distribution. Interestingly, α1(IV) to α5(IV) chains are found in the rat and mouse testis [89–92], whereas all six chains α1 to α6 are found in the bovine basement membrane in the testis and also in the kidney glomerular basement membrane [93, 94]. Besides testes, α3(IV) chain is also expressed in the ovary, kidney, lung, inner ears, and neuromuscular junction in rodents [89–92, 95, 96]. In this context, it is of interest to note that the expression of collagen α3(IV) chain peaked at 10–20 days postpartum (dpp) in the mouse testis, coinciding with the assembly of the BTB at 13–15 dpp [90]. These findings thus implicitly support the possibility that collagen α3(IV) chains in the basement membrane of the tubules may play a role in regulating BTB function, in particular its postnatal assembly to facilitate the onset of meiosis in rodents [97]. Collagen α3(IV) is likely the product of Sertoli and peritubular myoid cells in the testis since collagen α1(IV) and α2(IV) chains are synthesized by Sertoli cells and peritubular myoid cells in vitro [89, 98, 99]. NC1-Peptide from Collagen α3(IV) Chain Studies have shown that inclusion of an anti-collagen IV antibody in the apical chamber of Matrigel-coated bicameral units wherein primary Sertoli cells were cultured was capable of perturbing the assembly of a functional Sertoli cell TJ permeability barrier [100]. This finding is important because it illustrates that perturbation of collagen chain function in the basement membrane could disrupt Sertoli cell TJ barrier integrity. This observation is also consistent with an earlier report whereby modifications of the basement membrane by passive transfer of antibodies against seminiferous tubule basement membranes caused local epithelial sloughing that mimicked orchitis in rats [101]. It also showed that tumor necrosis factor alpha (TNFα), known to induce Sertoli TJ disruption

[102], was involved in this event by activating matrix metalloprotease 9 (MMP9), which, in turn, induced proteolytic cleavage of collagen α3(IV) chains to release the NC1-peptide to exert its biological effect on Sertoli cell TJ barrier function [100]. Subsequent studies by using purified recombinant NC1-peptide [103] or through overexpression of the NC1-peptide cDNA (cloned into the mammalian expression vector pCI-neo) [27] in primary Sertoli cells cultured in vitro have shown that either treatment is capable of perturbing Sertoli cell TJ barrier function. These findings thus confirm the notion that the NC1-peptide, similar to the F5-peptide, is an endogenously generated bioactive peptide from the structural component at the basement membrane (versus apical ES for the F5-peptide) (Fig. 2.7). However, unlike the F5-peptide that exerts its regulating effects on actin- and MT-based cytoskeletal function downstream through the signaling protein FAK [78, 79], the NC1peptide has recently been shown to exert its effects through Cdc42-mediated effects on actin- and MTbased cytoskeletons [104] [Fig. 2.8]. Furthermore, following cleavage of the NC1-peptide from collagen α3 (IV) chains in the basement membrane, likely involving MMP9 [27], the NC1-peptide is capable of being transported across the seminiferous epithelium to the adluminal compartment, via an MT-dependent transport mechanism, to exert its regulating effects at the apical ES by modulating the organization of actin- and MTbased cytoskeletons [27]. More importantly, the NC1peptide, similar to the F5-peptide [78], exerts its regulating effects via use of an integrin-based “inside–out” or “outside–in” signaling cascade. This notion is supported by the observation that the presence or absence of a signal peptide inserted into the cDNA clone encoding the NC1peptide has no notable differences regarding the potency of their effects on Sertoli cell TJ barrier function following the expression of these two cDNA clones [27]. Studies by confocal microscopy have confirmed disruptive changes in the distribution of both TJ (e.g., CAR, ZO-1) and basal (e.g., N-cadherin, β-catenin) proteins at the Sertoli cell–cell interface, due to defective organization of F-actin and MTs across the Sertoli cell epithelium, thereby leading to disruption of the Sertoli cell TJ-barrier [27]. The defects noted in the Sertoli cell epithelium following overexpression of the NC1-peptide were reproduced faithfully when the NC1-peptide was overexpressed in the testis in vivo. In brief, the NC1-peptide was overexpressed in the testis, which led to considerable defects in the status of spermatogenesis, including

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extensive exfoliation of germ cells from the testis due to epithelial damage, largely the results of defects in BTB integrity in the testis, as noted in a functional BTB integrity assay in vivo [27] (Fig. 2.7).

The LG3/4/5-Peptide In the basement membrane of the mammalian testis, besides collagen α3(IV) chains, laminin-α2 chains are another notable structural component [70, 71]. In contrast to the F5-peptide, which is released from domain IV of laminin-γ3 chain that perturbs Sertoli cell TJbarrier function, the 80-kDa fragment (or LG3/4/5peptide) from the C-terminal region of laminin-α2 chains is capable of promoting Sertoli cell TJ-barrier function, making the barrier “tighter” [105, 106] [Fig. 2.7]. This conclusion was reached based on the study wherein use of short hairpin RNA (shRNA) by targeting the laminin-α2 chain for its knockdown by RNAi was found to perturb the Sertoli cell TJ-barrier function [106], and this effect was subsequently shown to be mediated via the mTORC1/p-rpS6 signaling cascade downstream [105] (Fig. 2.8). In brief, disruption of the LG3/4/5-peptide function by blocking the laminin-α2 chain through its knockdown would lead to an upregulation of p-rpS6, the phosphorylatable protein translation regulator [105], which is the downstream signaling protein of mTORC1 [107] (Fig. 2.8). These findings are also consistent with earlier reports which showed that activation of the mTORC1/rpS6 signaling pathway would lead to disruptive changes in Sertoli cell BTB function both in vitro [108–110] and in vivo [111, 112]. While the effects of the LG3/4/5-peptide per se may not be this important, the combined physiological effects of the F5- and NC1-peptides, compared with the LG3/4/5peptide, in the testis as a whole are crucial to support spermatogenesis. The antagonistic effects of the LG3/4/ 5-peptide versus the F5- and NC1-peptides make these bioactive peptides serve as molecular switches to turn the BTB “on” or “off” to support the timely transport of preleptotene spermatocytes, and perhaps other substances, at the immunological barrier to support spermatogenesis (Fig. 2.7). This is important, as a recent report has shown that the immunological barrier at the BTB is highly selective, capable of blocking some, but allowing other, cellular products (e.g., sperm autoantigens) to move “in” or “out” of the BTB in the seminiferous epithelium [113]. At the time of writing, the biological function of the LG3/4/5-peptide and its downstream signaling cascade are being actively investigated,

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but two recent reports [105, 106] have demonstrated unequivocally the physiological significance of this novel bioactive peptide. For instance, using several antibodies against the laminin-α2 chain, including one that recognizes the 80-kDa fragment (the precursor of the LG3/4/5peptide), it was shown that the LG3/4/5-peptide generated at the basement membrane was transported to the adluminal compartment, such as the apical ES, to exert its effects via an MT-dependent mechanism [106]. This conclusion was reached by a study showing that in the adult rat testis, Taxol (via intratesticular injection, as described [114]) which specifically blocks MT depolymerization, was able to block transport of the LG3/4/5-peptide across the epithelium, when compared with control rats [106]. Also a knockdown of the laminin-α2 chains by shRNA was found to block the Sertoli cell TJ-barrier function via disruptive changes in the distribution of both TJ (e.g., CAR, ZO-1) and basal ES (e.g., N-cadherin, β-catenin) proteins through cytoskeletal disorganization of F-actin and MT cytoskeletons across the Sertoli cell cytoplasm [106]. Additionally, the effects of laminin-α2 knockdown on cytoskeletons were mediated through disruptive changes in the distribution of actin-regulating proteins (e.g., Arp3, Eps8, palladin), but also of MTregulating proteins (e.g., EB1) [106], which are necessary to maintain the homeostasis of actin (Fig. 2.5) and MT (Fig. 2.6) dynamics to support cellular events across the epithelium during the seminiferous epithelial cycle [7, 115–117].

Key Points • Fragments from laminin chains at the apical ES and the basement membrane generate the corresponding F5- and LG3/4/5-peptides to support spermatogenesis. • Fragments from the collagen α3(IV) chain in the basement membrane also generates the NC1peptide to support spermatogenesis. • These three bioactive peptides exert their effects through distinctive downstream signaling proteins.

Future Perspectives and Conclusion Here, we have summarized recent findings regarding the regulation of cellular events across the seminiferous

Chapter 2: Cellular Architecture

epithelium during the epithelial cycle pertinent to spermatogenesis through the actions of three locally produced bioactive peptides, as noted in Fig. 2.7. The involved signaling proteins and the plausible signaling cascade utilized by these bioactive peptides to modulate cellular events during spermatogenesis are summarized in Fig. 2.8. It is expected that much of this information will be updated in the next decade, as more information is available in the literature. Nonetheless, the model depicted in Fig. 2.6 will serve as a helpful guide to investigators. At present, many questions remain unanswered. For instance, what is the upstream signaling protein(s) that govern the production of these bioactive peptides during spermatogenesis? Are these cytokines and chemokines? Since these peptides serve as ligands, as noted in other studies, wherein fragments of collagens and/or laminins are ligands that exert their effects by binding onto integrin-based receptors through subsequent

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Acknowledgments Studies performed in C.Y.C.’s laboratory were supported, in part, by grants from the National Institutes of Health (NICHD, HD029990, Project 5 and R01 HD056034).

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129. Franca LR, Ogawa T, Avarbock MR, Brinster RL, Russell LD. Germ cell genotype controls cell cycle during spermatogenesis in the rat. Biol Reprod 1998;59:1371–7. 130. Clermont Y, Harvey SC. Duration of the cycle of the seminiferous epithelium of normal, hypophysectomized and hypophysectomized-hormone treated albino rats. Endocrinology 1965;76:80–9. 131. Russell LD, Brinster RL. Ultrastructural observations of spermatogenesis following transplantation of rat testis cells into mouse seminiferous tubules. J Androl 1996;17:615–27. 132. Aslam H, Rosiipen G, Krishnamurthy H, et al. The cycle duration of the seminiferous epithelium remains unaltered during GnRH antagonist-induced testicular involution in rats and monkeys. J Endocrinol 1999;161:281–8. 133. Leblond C, Clermont Y. Definition of the stages of the cycle of the

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seminiferous epithelium in the rat. Ann N Y Acad Sci 1952;55:548–73. 134. Oakberg EF. Duration of spermatogenesis in the mouse and timing of stages of the cycle of the seminiferous epithelium. Am J Anat 1956;99:507–16. 135. Clermont Y, Trott M. Duration of the cycle of the seminiferous epithelium in the mouse and hamster determined by means of 3Hthymidine and radioautography. Fertil Sertil 1969;20:805–17. 136. Russell LD, Franca LR, Brinster RL. Ultrastructural observations of

spermatogenesis in mice resulting from transplantation of mouse spermatogonia. J Androl 1996;17:603–14. 137. de Rooij DG, Russell LD. All you wanted to know about spermatogonia but were afraid to ask. J Androl 2000;21:776–98. 138. Ehmcke J, Schlatt S. A revised model for spermatogonial expansion in man: lessons from non-human primates. Reproduction 2006;132:673–80. 139. Ehmcke J, Wistuba J, Schlatt S. Spermatogonial stem cells: questions, models and perspectives.

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Hum Reprod Update 2006;12:275–82. 140. Heller CG, Clermont Y. Kinetics of the germinal epithelium in man. Recent Prog Horm Res 1964;20: 545–75. 141. Heller CG, Clermont Y. Spermatogenesis in man: an estimate of its duration. Science 1963;140:184–6. 142. Wu S, Yan M, Li L, et al. mTORC1/ rpS6 and spermatogenic function in the testis – insights from the adjudin model. Reprod Toxicol 2019;89:54–66.

Section 1 Chapter

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Scientific Foundations of Male Infertility

Maturation and Function of Sperm Caroline Kang, Nahid Punjani, and Dolores J. Lamb

Introduction Human sperm are created in the testis via a highly regulated and complex hormonal pathway arising from the hypothalamic–pituitary–testicular (HPT) axis under complex paracrine control by growth factors and cytokines. Following their creation, sperm must travel through the seminiferous tubules, epididymis, vas deferens, ejaculatory ducts, and urethra until they are expelled through ejaculation into the female genital tract. Fertilization capabilities of the sperm require morphologic and molecular changes that are acquired during transit of sperm through the male and female reproductive tracts. In this chapter, maturation and transport of sperm through the male reproductive tract, with a focus on the epididymis, are discussed. The process of normal spermatogenesis is presented, followed by sperm transport from the testis into the epididymis. Normal epididymal anatomy and the role of each segment in sperm maturation and transport, followed by cellular function, including the blood–epididymis barrier (BEB) and gene expression, are reviewed. Disorders of the epididymis related to male infertility and general management options for epididymal-related infertility are then examined.

functions of the epididymis is maturation of sperm, resulting in their acquisition of motility [1]. The peritubular connective tissue (lamina propria) and basement membrane of the seminiferous tubules contain myofibroblasts [2]. These myofibroblasts are large, flat cells connected through a network of collagen and microfibrils, and arranged in discontinuous layers within the extracellular matrix [3]. Myofibroblasts are critical for seminiferous tubule peristalsis which, together with hydrostatic pressure, propels newly produced, immotile sperm toward the rete testis [2]. Following their arrival in the rete testis, sperm then traverse through the efferent ducts into the epididymis.

Key Points • Spermatogenesis occurs in the seminiferous tubules of the testis. • Testicular sperm are immotile, and acquisition of motility occurs in the epididymis and female reproductive tract.

Spermatogenesis

Epididymal Development, Structure, and Function Epididymal Development

Spermatogenesis, or formation of mature sperm, occurs in the seminiferous tubules of the testis. This intricate biological process requires the coordination of various testis factors, including Sertoli and Leydig cells, and hormones stimulated by the HPT axis to occur correctly. Spermatogenesis consists of three phases, including the mitotic and meiotic phases and spermiogenesis. Mature sperm consist of a head, a midpiece, and a tail (Fig. 3.1) and are spermiated from the apical surface of Sertoli cells into the lumen of the seminiferous tubule. Sperm produced within the testis are immotile, and one of the many

The epididymis was first described in the nonscientific literature by Ben Johnson, a playwright, in the late 1500s and early 1600s [4]. In the eighteenth century, William Hunter, an anatomist, placed quicksilver in the epididymal lumen, followed by placement in turpentine, which helped to delineate the tubular structure, which varies across the three morphologically different segments – the caput (head), the corpus (body), and the cauda (tail) [4]. A primitive ductal system discovered in the vertebrate Chrondrichthyes may serve as a homeostatic maneuver for fertilization, playing a role in post-testicular

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Acrosome Head

Nucleus

Fig. 3.1 Normal sperm structure. Normal sperm structure includes a head, a midpiece, and a tail. Located in the head are the nucleus and the acrosome.

Midpiece

Tail

ducts from the residual mesonephric tubules [10]. These efferent ducts attach to the rete testis, and those failing to attach may atrophy or persist as an appendix of the epididymis (mesonephric duct remnant) or paraepididymides (also known as organ of Giraldes) [11]. Furthermore, prior to becoming the final epididymal structure, the anterior Wolffian duct undergoes a series of complex and coordinated steps. The first step includes elongation and expansion where the duct elongates along a caudal path, which occurs through tightly regulated cellular proliferation regulated by genes such as Gata3, as well as fluid secretion into the tubular lumen [9]. At this point, the elongated tubule also must undergo coiling, which is believed to occur from the creation of a morphogenic gradient along the duct. This, in turn, induces folding, along with cellular proliferation in critical areas, which results in space constraints that induce folding [9]. This folding is induced by predetermined signals along the duct, which may explain why the vas deferens does not coil [9]. Following elongation, the duct reorganizes into morphologically and functionally distinct segments. A series of genes, including the Homeobox (Hox) gene family, control the expression of morphogens in the duct, conferring different phenotypes because of their distance from the morphogenic source. Other possible drivers include the secreted fluid, which induces cellular proliferation. Overall this variable differentiation along the same tubule occurs secondary to septa along the tubule, which ultimately create the various distinct segments [9].

General Epididymal Structure sperm maturation and storage [5]. This revealed that all vertebrate species conducting internal fertilization by the male gamete had an epididymal structure which permitted maturation upon leaving the testis [6]. During normal embryologic development, sexual differentiation begins at the seventh week, at a time where both the Müllerian and Wolffian ductal systems are present [7]. Masculinization into a male occurs from release of anti-Müllerian hormone (AMH) and testosterone, both produced by the testis [8]. AMH causes regression of the Müllerian structures, following which the Wolffian ducts develop into male reproductive structures [9]. By day 84 of gestation, the upper (anterior) portion forms the epididymis, vas deferens, seminal vesicles, and ejaculatory ducts. More specifically, the epididymis arises from the mesonephric (Wolffian) duct, and efferent

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The human epididymis is adherent to the superior pole of the human testis (Fig. 3.2). It is continuous with the seminiferous tubules through efferent ducts in the rete testis, and attached along the lateral aspect of the testis via epididymo-testicular connective tissue and more distally by an epididymal fat pad [12]. Given its shape, it is often described as a “comma-shaped” organ. On average, the human epididymis is 10–12 cm in length in the adult human male, as measured from the epididymal head to the convoluted portion of the vas deferens [12]. The epididymis consists of a concentric layer of fibrous tissue, with an interconnected vascular network rich in blood vessels and nerves. In general, there is peristaltic movement of smooth muscle within the ducts to facilitate sperm flow through the epididymis and eventually the vas deferens.

Chapter 3: Maturation and Function of Sperm

Straight portion of vas Caput

Epididymis

Efferent ducts

Rete testis Corpus

Seminiferous tubules Testis

Cauda Convoluted vas Fig. 3.2 Structure of the epididymis and testis. Illustration showing the cross-sectional anatomy of the testis and epididymis. Seminiferous tubules are depicted in the cross-sectional image of the testis. The seminiferous tubules are linked to the efferent ducts by the rete testis. The epididymis is composed of the caput (magenta), corpus (gray), and cauda (green).

Arterial supply of the epididymis comes from multiple sources. The epididymal artery, which arises from the internal spermatic artery, supplies the efferent ducts, caput, and corpus and is surrounded by a network of epididymal veins. The deferential artery, which arises from the iliac or hypogastric artery, supplies the cauda and epididymo-testicular connective tissue [13]. The venous return of the epididymis occurs via the pampiniform plexus, deferential vein, and cremasteric vein [14]. The epididymis is controlled by autonomic innervation, and is thought to be almost completely devoid of somatic innervation [15]. The distal epididymis receives sympathetic innervation from the inferior mesenteric ganglion, which lies caudal to the mesenteric artery, and additional spermatic nerves provide sympathetic (middle and inferior spermatic nerves) and parasympathetic (inferior spermatic nerves) innervation to the epididymis, both of which arise from the hypogastric nerves [15]. Denervation studies in rats have shown a significant role of the autonomic, primarily sympathetic, nervous system in the regulation of sperm maturation, transport, and storage within the epididymis.

Initial Segment In animal models, an additional structure exists, known as the initial segment, which is situated between the caput

and the efferent ducts [16]. The number of segments varies between species (e.g., two segments exist in mice, and four segments in rats) [17]. The role of the initial segment for infertility is demonstrated by mutations in three genes localized to this region, including Ros1, Gpx5b, Pten, in mouse models. In all cases, sperm have hairpin bends at the sperm midpiece [17]. Furthermore, this segment has significant vascularity and is uniquely regulated in animal models, but no distinct anatomical counterpart exists in humans [17]. Instead, humans have efferent ducts, discussed below, which act as the pathway from the rete testis to the caput epididymis [18].

Efferent Ducts As previously described, the efferent ducts arise from the rete testis at the superior pole of the testis, and are straight or mildly convoluted and entrenched within fat and the superior epididymal ligament (Fig. 3.2) [19]. These ducts form a large section of the caput (head) of the epididymis and act as a conduit for sperm transport from the testis [19]. The main function of the efferent ducts is luminal fluid reabsorption (approximately 90% of the total fluid) from columnar principal cells, which results in concentration of sperm [20]. The fluid is isosmotic, and reabsorption involves numerous ion transporters and

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aquaporin channels which are modulated by estrogens [20]. The abundance of estrogen receptors in this location suggests that this region may provide a unique target for male contraception, but spermatogenic failure due to deficient fluid resorption seen in the estrogen receptor knockout mouse models showed that this male contraceptive target is not ideal [21, 22]. Sperm can traverse the efferent duct, in part, by constant fluid secretion from the seminiferous tubule epithelium, contraction of peritubular myofibroblasts, and a vacuum generated by ejaculation and pressure from the branching and convergent anatomic ductule system [23]. The efferent ducts also are unique in containing both absorptive cells along with cells containing motile cilia, which also function to propel sperm into the epididymis [24]. Other functions of the efferent ducts include secretion of glycoconjugates and enzymes and phagocytosis of abnormal spermatozoa [25, 26].

Caput A large majority of the epididymal caput is composed of efferent ducts, as previously mentioned (Fig. 3.2). The caput plays significant roles in sperm maturation [27]. Upon arrival to the caput, most of the fluid has been resorbed (approximately 99%), resulting in high sperm and protein concentrations [28]. The caput contains principal cells, which are involved in secretion of proteins that adhere to the sperm membrane and subsequently alter the plasma membrane composition [29]. In fact, the caput is the most metabolically active region of the epididymis, with cells in this region secreting up to 80% of total protein found in the epididymal lumen [28]. In the caput epididymis, sperm acquire clusterin, a cell-aggregating protein involved in lipid transport, inflammation regulation, and cellular differentiation. This molecule plays a role in caudal cytoplasmic droplet migration of sperm [29]. There are conflicting reports as to whether cytoplasmic droplets promote sperm maturation and motility or inhibits these processes. In addition, the caput contains high amounts of water (aquaporin) channels, calcium channels, chloride channels, sodium/potassium transporters, potassium channels, and ion exchangers, in comparison to the corpus and cauda, suggesting a role of these ion channels in sperm maturation [27]. Clear cells are present in the caput, and these cells are responsible for phagocytosis of cytoplasmic droplets and luminal proteins and debris [30]. Clear cells are also thought to secrete and maintain appropriate levels of immobilin, a protein which serves to

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physically restrict flagellar propagation, rendering sperm immotile [30, 31].

Corpus The corpus of the epididymis also plays an important role in late sperm maturation (Fig. 3.2) [27]. This region is composed of principal cells, which have large amounts of lipids, contributing to lipid content modification in the sperm plasma membrane [29]. The corpus and cauda also have higher expressions of innate immunity molecules, such as defensins, which are present throughout various regions of the epididymal epithelium [27]. The human corpus is thicker than that in other mammals [6].

Cauda The cauda of the epididymis acts as a storage reservoir for mature male sperm since maturation occurred prior to sperm arrival (Fig. 3.2) [27]. While variable in different species, about 50–80% of sperm in the epididymis are stored here [1]. The epithelium in this region is shorter than in the other regions, and is composed of clear cells which phagocytose cytoplasmic droplets and luminal debris that are shed from maturing spermatozoa [29]. Here, excess luminal proteins are resorbed, with subsequent secretion of immobilin, maintaining mature sperm in a quiescent, dormant state prior to ejaculation [29]. Structurally, the cauda is surrounded by two smooth muscle layers, as opposed to the caput, the earlier portion of the epididymis, only containing a single layer [1]. Additionally, the cauda in humans is thought to have a reduced sperm reservoir capacity than is present in other species [6].

Effects of Aging on the Epididymis As males age, there are genetic and cellular changes in the epididymis that can have an effect on spermatogenesis. Most of this work has been elucidated in the Brown Norway (BN) rat, a model used extensively in the study of the aging male reproductive system [32]. In male BN rats, similar to human males, aging results in declining testosterone levels and decreased spermatogenesis. These rats are ideal models for studying the reproductive system, as they gain little weight and very rarely develop pituitary, testicular, or other tumors that may confound male reproductive findings [33]. With age, the epididymal epithelium accumulates lipofuscin, lysosomes, and vacuoles and the basement membrane thickens, all of which are characteristic signs of aging [34]. Different

Chapter 3: Maturation and Function of Sperm

segments of the epididymis are affected more than others and the effects of aging occur, regardless of whether spermatozoa are present [35]. Jervis and Robaire used complementary DNA (cDNA) microarrays to analyze gene expression during aging in the BN rat epididymis. Overall levels of gene expression decreased with age and the discrepancy between old and young epididymides was more pronounced in the corpus and cauda, compared to the initial segment and caput [32]. Genes encoding ribosomal components and adenosine triphosphate (ATP) synthesis machinery were globally decreased in all epididymal segments in older male BN rats, compared with younger males [32]. Additionally, heat shock protein expression was decreased in all epididymal segments. It is also well known that expression of heat shock proteins, which are activated by numerous stressors, including exposure to heat or reactive oxygen species (ROS), is important for repairing damaged proteins. Only several genes had increased expression in all four segments, including tissue inhibitor of metalloproteinase-2 (TIMP-2), a protease involved in extracellular matrix remodeling [32]. This may explain the thicker basement membranes noted in the epididymides of older BN rats. Lysosomal cathepsins and components of the proteasome had decreased expression in the epididymides of older BN rats, with a more dramatic decrease in the corpus and cauda, compared to the initial segment and caput [32]. Given the multiple functions of the different segments in the epididymis, it is not surprising that gene expression levels are different and dependent on age. Furthermore, loss of BEB function has been noted in older BN rats, allowing an increased number of halo cells (lymphocytes and monocytes) to be present in the epididymis [36].

General Function of the Epididymis The epididymis is a critical male reproductive organ and plays a role in sperm transport, concentration, protection, storage, and maturation [1]. Transit time through the epididymis from the rete testis to the vas deferens is approximately 10–15 days, and occurs through rhythmic contractions of smooth muscle layers surrounding the epididymis [1, 37]. Contractions occur along the entirety of the epididymis but are most amplified in the cauda. In addition to smooth muscle contraction, cilia on luminal epididymal cells also propel sperm through the epididymis [37]. While sperm move through the epididymis, fluid resorption occurs through epithelial

cells. Resorption of fluid occurs along all segments of the epididymis, but most resorption occurs in the efferent ducts and initial segment [1]. These epithelial cells also have a high metabolic rate, resulting in the generation of ROS which may be harmful to sperm. To protect sperm, epithelial cells secrete various antioxidant enzymes, such as superoxide dismutase, into the epididymal lumen to neutralize ROS [1]. Additional protection from toxins or immune cells in the blood is afforded by the BEB, which will be discussed in the following section. As sperm are transported through the epididymis, maturation (acquiring motility and other factors necessary to fertilize an ovum) occurs. For maturation to occur, sperm must directly contact specific factors within the epididymal lumen, and the composition of luminal contents and epithelial layers varies along the length of the epididymis. Changes in nuclear compaction, plasma membrane composition, cytoskeletal structure, and protein and noncoding RNA content of sperm occur during epididymal transit [1]. Furthermore, both testosterone and dihydrotestosterone are involved in sperm transmit and maturation through the epididymis. Deprivation of testosterone reduces sperm quantity and modifies the epididymal epithelium, hindering proper sperm maturation [29]. Restoration of testosterone reverses cellular changes in the entire epididymis, except in the initial segment [28]. Progressive motility, which allows sperm to propel themselves toward the ovum, during passage through the epididymis. Immotile sperm have the machinery required for full motility, and different types of motility are achieved in different epididymal segments. In the caput, sperm become motile but exhibit different flagellar bending in the cauda. One proposed mechanism for acquisition of motility is through cyclic adenosine monophosphate (cAMP)-dependent protein kinase A (PKA) phosphorylation of proteins [38]. Mice lacking ADCY10 (the soluble adenylyl cyclase gene present in sperm), which converts intracellular ATP to cAMP, had morphologically normal, but immotile sperm [39]. Another proposed mechanism is a cAMP-dependent pathway involved in the exchange of proteins activated by cAMP (EPACs, formerly known as RAPGEF3 and RAPGEF4) [38]. Serine phosphorylation of GSK3 increases, as sperm pass through the epididymis, and is associated with acquisition of sperm motility [38]. Further work is needed to determine the precise signaling pathways involved in the acquisition of sperm motility. Caudal migration and loss of the cytoplasmic droplet are associated with sperm maturation and are the most obvious change in sperm during epididymal

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transit [40]. The cytoplasmic droplet is an organelle that contains the germ cell cytoplasm not phagocytosed by Sertoli cells and left over from the spermiogenesis process. It has been proposed that the cytoplasmic droplet plays a role in regulating sperm ion homeostasis via potassium, chloride, and water channels [41]. However, the precise role of the cytoplasmic droplet is unknown. Retention of the cytoplasmic droplet may result in reduced fertility in animal models [40]. Morphologically, the sperm plasma membrane composition is altered by varying concentration gradients of specific luminal enzymes and molecules, and results in narrowing of the acrosome [1]. High concentrations of glycohydrolases and glycosyltransferases are present in the epididymal lumen and cause alterations of the membrane, including removal or modification of glycoproteins and polysaccharides [40]. These changes may promote further maturation of sperm or facilitate membrane changes that are essential for egg fertilization [40]. Finally, the epididymis functions to store sperm until ejaculation occurs. The majority of sperm in the epididymis are located in the cauda epididymis [40]. Epididymal epithelial cells within the cauda epididymis secrete factors that promote an environment that maintains sperm in a quiescent state. Further maturation occurs in the female genital tract and will be discussed in the next sections.

Key Points • The epididymis is formed from a portion of the Wolffian duct during male fetal development. • The human epididymis consists of various segments, including the efferent ducts, caput (head), corpus (body), and cauda (tail). • Acquisition of motility and caudal migration of the cytosolic droplet are two major changes in sperm that occur during transport through the epididymis.

Molecular Composition of the Epididymis Cellular Composition of the Epididymis and Blood–Epididymal Barrier The BEB is a structural barrier that limits the exchange of molecules between the epididymal lumen and blood in order to maintain immunoprivilege, or

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exemption from immune surveillance, for maturing spermatozoa [36]. Epididymal tubules are lined with pseudostratified epithelium made up of principal cells. Principal cells are connected to one another by tight junctions, which comprise proteins that link adjacent cells to each other, allowing for segregation of proteins from the basolateral surface from the apical (or luminal) surface and limiting movement of blood components to the epididymal lumen, and vice versa [36]. The BEB begins to form during embryonic development, and decrease in permeability of the BEB likely continues to develop after birth [36]. Immune cells are present in all segments of the human epididymis [42]. The peritubular zone and epithelium of the caput epididymis contain the highest number of immune cells, with decreasing numbers of immune cells toward the cauda epididymis and vas deferens. The immune cell types present in the epididymis include macrophages, dendritic cells (DCs), basal cells, and halo cells, or lymphocytes [42, 43]. The number of immune cells is highest in the cauda epididymis and progressively declines, with the lowest number in the cauda epididymis and vas deferens [42].

Macrophages Macrophages are the most abundant immune cells present in the epididymis and are located predominantly in the interstitium and peritubular layers [44]. Macrophages in the stroma typically express major histocompatibility complex (MHC) II antigens which are required for CD4+ T cell activation. However, macrophages within the epididymal epithelium do not express MHC II and the majority of lymphocytes present in the epithelium are CD8+ T cells [45]. The predominance of CD8+ T cells may prevent the development of an autoimmune response to maturing sperm [45].

Dendritic Cells DCs also are present in the basal region of the epithelium and extend their processes between the epithelial cell tight junctions toward the epididymal tubule lumen [36, 46]. These DCs are thought to sample antigens within the epididymal lumen, and present antigen to CD4+ T cells to regulate the immune response to spermatozoa and any pathogens present in the epididymis.

Basal Cells Basal cells, also present in the basal region of the epididymal epithelium, are dendritic-like cells that can extend

Chapter 3: Maturation and Function of Sperm

their processes into the epididymal tubule lumen [36, 42, 43]. These cells may sample the epididymal lumen and express the angiotensin II receptor, allowing them to detect angiotensin II in the lumen [43, 46, 47]. Cross-talk between basal cells and adjacent clear cells results in proton secretion by clear cells after binding of angiotensin II (to the angiotensin II receptor on basal cells) [47]. Additionally, basal cells also may control electrolyte and water transport by principal cells. Leung et al. hypothesized that tRNA-pro (anticodon TGG) 3–1 (Trp3) and cyclooxygenase 1 (COX-1) in basal cells result in activation of the cystic fibrosis transmembrane conductance regulator (CFTR) and secretion of anions, and subsequently water, into the epididymal lumen [48]. Furthermore, expression of aquaporins in the epididymal epithelium is modulated by hormones, such as androgens and estrogens, and also may allow movement of water [43].

Blood–Epididymis Barrier Loss of BEB function, determined by decreased expression of tight junction proteins between epididymal principal cells, in BN rats was associated with increased halo cell number in the epididymis. In humans, loss of BEB function can result in decreased fertility. Epididymal cell lines derived from infertile patients had lower expression and mislocalization of tight junction proteins, resulting in the inability to form functional tight junctions [49]. The BEB is more permeable than the blood–testis barrier, and immune cells can directly interact with epididymal luminal antigens, as previously mentioned [42]. Sperm granulomas, or masses containing leaked sperm in the interstitium surrounded by macrophages, form readily in the epididymis, but not in the testis [50]. Additionally, the epididymis is more susceptible to infection than the testis, and there is a gradual decrease in this susceptibility, the closer the epididymal segment is to the testis [50]. Expression of immunoregulatory proteins in the proximal epididymis is similar to that observed in the testis, suggesting that the proximal epididymis has protein expression profiles that promote a more immunoprivileged status [50]. Indoleamine 2,3-dioxygenase (IDO) is an immunoregulatory enzyme that controls the cytokine milieu in the epididymis. Proinflammatory cytokine expression is increased in the caput epididymis of IDO-deficient mice. Activin A regulates IDO expression via the SMAD 2/3/4 signaling pathway. Both IDO and activin A expression are highest in the caput epididymis [51]. Additionally, Toll-like receptors (TLRs), which are important for sensing pathogens, are expressed

differentially in the epididymis. TLR-4 levels in the caput epididymis are more similar to those seen in the testis, and expression of TLR-4 decreases distally toward the cauda epididymis and vas deferens [52].

Gene Expression in the Epididymis Since the structure of the epididymis is so unique, gene expression is highly regulated and varies throughout its entire structure along the various segments. However, the mechanisms regulating gene expression remain largely unknown [6]. Research on the epididymis in various animal models suggests variable gene expression amongst species [6]. In humans, while the segments (caput, corpus, and cauda) are distinct, gene expression along each segment appears not to be as discrete as present in other species, as confirmed by studies revealing reasonable similarity in both the epididymal secretome and proteome [6]. Furthermore, studies in men with epididymal obstruction suggest that maturation occurs in more proximal regions (closer to the testis) than described in other species; however, this is refuted by the observed rapid transport of sperm through the more proximal epididymis [6]. Transcriptome analysis demonstrated 78% of human proteins are expressed in the epididymis and 412 genes have elevated expression in the epididymis [53]. Highlighted below are some of the important genes with a more in-depth understanding.

Androgen Receptor (AR) Androgens are heavily involved in gene expression in the epididymis, based on in vitro studies on various androgen-dependent signaling pathways and genes regulated through transition of androgen response elements [16]. Furthermore, since AR is expressed throughout the entirety of the epididymis, gene expression changes due to the effects of androgen may result from differential or variable expression of AR coregulators and/or a concentration gradient of androgen along the course of the epididymis [54, 55]. In addition to androgens, estrogens also play a role in gene expression, and it is possible that stimulation by both androgen and estrogen is required for appropriate epididymal gene and protein function. One example of coregulation by androgen and estrogen is regulation of aquaporin (AQP9) [21].

Estrogen Receptor (ESR) The ESR family of genes encode estrogen receptors. ESR1 and ESR2 are expressed in the human genital tract. In the

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efferent ducts, ESR1 is localized to epithelial cells and ESR2 to ciliated cells, but only ESR2 is found in the caput [56]. As previously discussed, estrogen receptors are involved in fluid regulation and therefore play a critical role in sperm transport and maturation through the epididymis. Mice lacking ESR1 have impaired spermatogenesis and sperm morphology, motility, and viability [57]. Mice lacking estrogen receptor alpha had severely dilated efferent ducts and rete testis due to diminished capability of the efferent duct cells to resorb fluid [58, 59]. This defect resulted in decreased spermatogenesis in these mice [58, 59].

Phosphatase and Tensin Homolog (PTEN) PTEN is a tumor suppressor involved in various cellular pathways, including ERK and AKT pathways. Selective removal of Pten expression from the initial segment in mice resulted in AKT activation and ERK pathway suppression [60]. Furthermore, loss of Pten in the initial segment resulted in altered cell shape, size, and organization, signifying epididymal epithelial cell dedifferentiation [60]. In these mice, sperm leaving the testicle were normal; however, epididymal maturation did not occur. Thus, these sperm had bent flagella and diminished motility, resulting in male mouse infertility [60].

Dicarbonyl and L-Xylulose Reductase (DCXR, P34H) DCXR is an ortholog of P26h, a glycosyl phosphatidylinositol (GPI)-anchored hamster sperm protein, and is expressed in the corpus. In animal models, P26h has roles in binding of sperm to the zona pellucida, and in the human corpus epididymis, DCXR accumulates on the surface of sperm to cover the acrosome, which suggests a role as a sperm maturation marker [6]. This protein is undetectable in the sperm of 15% of men presenting with infertility, and defects in DCXR have been demonstrated in 40% of men with idiopathic infertility [6, 61]. Sperm from these men display defective ability to bind to the zona pellucida and also have elevated failure rates of in vitro fertilization [6, 61]. Levels of DCXR have also been shown to decrease following vasectomy. Therefore, DCXR may be used as a surrogate for men with vasectomy reversal, such that lower rates suggest a longer period of obstruction [6].

Adhesion G Protein-Coupled Receptor (ADGRG2) The ADGRG2 gene encodes a family of G proteincoupled receptors which are expressed in the efferent ducts of the epididymis [62]. It was discovered in

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ADGRG2 mutations in mice that fluid reabsorption was disrupted which led to an obstructive infertility phenotype, similar to that seen in men with congenital bilateral absence of the vas deferens [63].

Human Sperm Antigen 11 (SPAG11, HE2) SPAG11 is related closely to beta-defensins, which are broadly acting antimicrobials and androgen-dependent, and secrete specific epididymal proteins [64]. These proteins are observed in seminal plasma, luminal fluid, and the epididymal epithelium [65]. Within SPAG11, there are two genes – SPAG11A and SPAG11B [66]. SPAG11A is androgen-dependent and expressed in the principal cells of the mouse epididymis, suggesting its role in maturation [67]. SPAG11B is localized to the sperm neck and head regions and expressed in the epididymal epithelium, and is thought to be involved in signaling pathways implicated in sperm motility and maturation [66].

Binder of Sperm Protein Homolog 1 (BSPH1) BSPH1 is a member of the binder of sperm family of proteins expressed in the epididymis and is also localized to the neck of human sperm, the equatorial segment, and the postacrosomal segment [68]. Bovine and murine studies have demonstrated an increase in capacitation due to BSP (bovine and murine protein) interactions with the sperm plasma membrane. It is postulated that BSPH1 may interact with, and stabilize, the sperm plasma membrane, and once in the female reproductive tract, BSPH1 becomes dissociated from the sperm plasma membrane, allowing for sperm plasma membrane changes that promote capacitation [68].

Niemann Pick Disease Type C2 (NPC2, HE1) NPC2 encodes the human epididymal secretory protein. This gene is highly conserved across mammals and is involved in cholesterol binding, which alters the efflux of lipids, stabilizing the sperm plasma membrane during epididymal transit [69]. Gene expression protein products have been detected across the entirety of the epididymis, but not in vasectomized men who exhibit higher cholesterol content in sperm [70].

ADAM Metallopeptidase Domain 7 (ADAM7) ADAM7 encodes disintegrin- and metalloproteasedomain-containing protein 7. ADAM7 is expressed in the caput epididymis in mice [71]. It is transferred to the sperm surface during its transit through the

Chapter 3: Maturation and Function of Sperm

epididymis and therefore may play a role in the function of the sperm plasma membrane [72]. Its exact function remains unknown.

Cysteine-Rich Secretory Protein 1 (CRISP1) CRISP1 encodes a cysteine-rich secretory protein acquired by sperm during transit through the epididymis, and may play roles in fertilization, as well as in decapacitation [73, 74]. It is expressed in the corpus and caudal segments of the human epididymis, and upregulation of this gene has been demonstrated in vasectomized and azoospermic men [75, 76]. Furthermore, CRISP1 is involved in binding of sperm to the zona pellucida and plasma membrane of the oocyte. However, its role remains to be clearly defined [77].

Cluster of Differentiation 52 (CD52) CD52 encodes GPI, which is expressed on lymphocytes [78]. Large amounts are present in the seminal fluid, and its occurrence on the rat sperm surface is thought to be related to some aspects of sperm maturation. However, knowledge of its function is limited in humans [78]. While in animal models, it has been referred to as “maturation-association sperm antigen,” its precise function, however, still remains unknown [79].

Adenylate Kinases 1 and 7 (AK1 and AK7) AK1 and AK7 encode two adenylate kinases which are localized to the sperm flagella. AK1 is thought to play a role in sperm motility, and AK7 has been shown to be important for proper ciliary function [80].

Defensin Beta (DEFB) Beta-defensins are small antimicrobial peptides expressed in the testis and epididymis, and conserved amongst species [81]. In addition to their role in immunity, betadefensins also contribute to male fertility by promoting sperm motility and maturation and capacitation [16]. In humans, beta-defensins are mostly expressed in the corpus and mutations are associated with impaired sperm function [82]. DEFB106b is thought to be involved in motility and antimicrobial activity, DEFB1 in the caput, and DEFB125 and DEFB126 in the cauda. The latter is involved in zona pellucida binding and sperm penetration through cervical mucus [82, 83].

Lipocalin (LCN) Lipocalins are a family of proteins involved in hydrophobic ligand transport. Ten different lipocalins are

expressed in the epididymis, and prostaglandin D2 synthase is the most highly expressed [84]. In humans, these genes are expressed in distinct segments of the epididymis and thought to be involved in maturation; as opposed to animal models such as rodents, these are only expressed in the initial segment and caput [16].

Aquaporin (AQP) AQP encodes proteins in the family of aquaporins. Aquaporins have a distinct role in water transport and play a critical role in fluid reabsorption necessary for concentration of sperm along the epididymis. Various aquaporins have been expressed in the epididymis (AQP1, 3, 5, 7, 9, and 11), of which AQP9 is highly expressed in the caput and corpus, and AQP1 in the distal portions of the epididymis [85].

Other Studies in animal models suggest additional factors that may regulate gene expression include lumicrine factors (those from Sertoli and germ cells), various growth factors (i.e., fibroblast growth factor 2), sperm factors produced directly from spermatozoa, and thermodynamic factors including temperature and pressure [16, 86]. Additionally, it has been suggested that small noncoding RNAs, such as microRNAs (19- to 22nucleotide regions), may also be critical in regulating epididymal function [16]. Many of these areas require further and additional studies.

Key Points • The BEB limits the exchange of material between the epididymal lumen and the bloodstream. • Various genes are expressed throughout the epididymis. However, more research is needed to determine their precise role in sperm maturation and transport.

Sperm Transport and Maturation Beyond the Epididymis Sperm Transport from the Epididymis Sperm are stored in the epididymis prior to ejaculation. During ejaculation, two processes occur – emission and expulsion. During emission, the bladder neck closes and prostatic secretions are released, consisting of zinc, citric

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acid, and acid phosphatase, and become mixed with sperm stored in the distal epididymis and vas deferens [87]. This fluid combines with alkaline fluid from the seminal vesicles (which composes the majority of the ejaculate), as well as with a small volume of fluid from periurethral and Cowper’s glands [87]. During expulsion, the bladder neck first closes to prevent retrograde ejaculation into the bladder and seminal fluid is expelled via rhythmic muscular contractions of the ischiocavernosus and bulbocavernosus muscles [88].

Sperm Transport Within the Female Genital Tract Following ejaculation and expulsion during coitus, sperm are deposited into the vaginal canal. The vaginal environment is acidic and harmful to sperm. However, rapid progression toward the cervix occurs via contact with cervical mucus [89]. Strategies to tackle the acidic environment include a buffering ability of seminal plasma (pH range 6.7–7.4), and immune inhibitors [89]. Cervical mucus comprised mostly of water but poses some barriers to sperm, ultimately filtering out abnormal or poorly progressive sperm [90]. There is some evidence to suggest that the cervix can also mount an immune response to sperm via leukocyte invasion, but overall this appears to have minimal impact on sperm [91]. Upon progressing through the cervical os where mucus is most concentrated, sperm enter the uterus assisted by myometrial contractions as well as cervical mucus. Damaged sperm undergo immunologic attack at this point [89]. Sperm then traverse the uterotubal junction into the fallopian tube, which does not have the same degree of immunologic response, as in previously encountered organs [89]. At this point, in animals, it is suspected that sperm may be stored in the fallopian tube, but this has not been seen in humans [92]. Finally, sperm must undergo capacitation and hyperactivation in the female genital tract prior to fertilization of the oocyte.

Sperm Maturation Beyond the Epididymis and Within the Female Genital Tract To fertilize an ovum, sperm must undergo maturation processes in both the epididymis and the female genital tract. Capacitation, or changes in sperm mobility to produce hyperactivated movement, was first described in 1950s [93, 94]. The process of capacitation is associated

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with sperm plasma membrane changes (loss of cholesterol, increased fluidity, hyperpolarization of the membrane) and intracellular changes (changes in intracellular ion concentrations, intracellular alkalinization, increased PKA activity, and protein phosphorylation) [95]. In the epididymis at baseline, luminal fluid contains low concentrations of sodium (approximately 40 mM), chloride (approximately 40 mM), and bicarbonate (approximately 4 mM), and high concentrations of potassium [96]. These low bicarbonate levels and low pH allow sperm to maintain a quiescent state prior to ejaculation [95]. Several active transporters located in epididymal epithelial cells, including Na/H antiporters, Na/HCO3 cotransporter, and vacuolar H–ATPase, along with carbonic anhydrase, are critical to the maintenance of low tubular fluid pH levels [96]. In seminal fluid, pH is increased due to basic secretions of the seminal vesicles. Additionally, seminal fluid concentrations of sodium, chloride, and bicarbonate are increased (hence increased pH), while potassium levels are decreased [97]. Upon exposure to higher levels of the resulting bicarbonate, increased intracellular transport of bicarbonate is observed. Inside sperm, bicarbonate targets ADCY10, which catalyzes the conversion of ATP to cAMP [95, 98]. Increased cAMP concentrations result in activation of PKA, which is localized in the sperm flagellum, and PKA activation is thought to be the initial step in various signaling pathways associated with capacitation [95].

Capacitation of Sperm The initial event of capacitation is likely intracellular alkalinization, followed by membrane hyperpolarization and calcium influx and signaling. Increase in pH occurs upon entry into the alkalinized portion of the female genital tract and secondary to the action of proton transporters, Hv1, and sodium–hydrogen exchangers (NHEs), as well as CFTR, or the cystic fibrosis transmembrane conductance regulator, which is a cAMP-activated chloride and bicarbonate transporter. Membrane hyperpolarization occurs through the actions of Na+/K+ ATPase, which, at rest, maintains a membrane potential of about 40 mV in human sperm [99]. Activated SLO3 (or sperm-specific potassium (KSper)) channels allow efflux of potassium out of the cell, which changes the membrane potential of the plasma membrane. The change in membrane potential activates various voltagegated channels, including Hv1, ion exchangers, and

Chapter 3: Maturation and Function of Sperm

calcium channels [99]. This resulting increase in intracellular calcium concentration is required for hypermotility and the acrosome reaction, and occurs via CatSper (cation channels of sperm) [99]. Ion channels that are important in capacitation are discussed in the next section. As previously described, intracellular alkalinization must occur for capacitation to take place. The vagina and uterus are relatively harsh environments for sperm. However, once sperm reach the uterotubal junction, there is an increase in pH of the surrounding fluid to approximately 8 and capacitation is initiated [89, 95, 100]. Increase in intracellular pH is thought to occur via outward movement of protons or inward movement of bicarbonate anions, which results in sperm hypermobility through activation of channels which allow calcium influx [101, 102]. Movement of protons extracellularly has been proposed to occur via Hv1, a voltage-gated H+ transporter located in the flagella, or a sperm-specific Na/ H exchanger (sNHE) [103, 104]. Movement of bicarbonate intracellularly has been proposed to occur via CFTR and associated Cl /HCO3 transporter or via a member of the Na/HCO3 cotransporter family [105]. Overall intracellular alkalinization results in activation of CatSper, a calcium-permeable channel, along with KSper. CatSper is thought to be responsible for hyperactivity of the sperm flagellum, chemotaxis toward the ovum, and the acrosome reaction [102]. The KSper channel also is activated by intracellular alkalinization, which results in hyperpolarization of the sperm plasma membrane due to outward flow of potassium ions [95, 106]. Furthermore, sperm with mutated KSper were found to undergo membrane depolarization, rather than hyperpolarization, and these sperm were unable to undergo the acrosome reaction [99]. Signaling pathways involved in sperm capacitation include bicarbonate-mediated activation of ADCY10, which converts ATP to cAMP. cAMP then activates PKA to phosphorylate downstream effectors, which results in rapid changes in sperm motility, and may activate CatSper to allow calcium influx [99]. Other possible cross-talks between the different channels and processes during capacitation include direct PKA-mediated phosphorylation of KSper channels due to activation of Src tyrosine kinase, or cAMP-binding to sNHE inducing intracellular alkalinization which activates KSper channels [95]. Further work is needed to determine the precise pathways activated by specific channel activities and how these pathways intersect.

Sperm Channels Involved in Capacitation Various ion channels are implicated in sperm capacitation. Specifically, ion channels, as mentioned in the previous section, are involved in intracellular alkalinization, calcium signaling, and membrane hyperpolarization, all of which are required for capacitation. Intracellular Alkalinization Proton efflux across the sperm plasma membrane, which results in intracellular alkalinization, occurs via two wellstudied channels – NHEs and Hv1. Both channel types are found within the principal piece of the flagella [99]. NHEs are encoded by the Slc9 gene and transport sodium ions into, and hydrogen ions out, of cells [99]. Two particular NHEs, sNHE and NHA1, are expressed in the sperm flagella and are important regulators of sperm pH regulation [99]. sNHE is thought to be voltage-sensitive [107]. Loss of sNHE or NHA1 results in male infertility in mouse studies, with defects observed in sperm motility [107, 108]. In these studies, addition of ammonium chloride partially rescued the motility and fertility defects in mice lacking sNHE [107]. A separate proton channel, Hv1, is also important for regulation of intracellular pH of human sperm [99]. Hv1 activation is voltagedependent and its activity is inhibited by zinc [99]. Super high-resolution microscopy demonstrates Hv1 channels to be situated along the side of the flagellar membrane, arranged in bilateral longitudinal lines [109]. The specific distribution of these channels affects the rotation of the sperm flagella, depending on whether the channels are activated or inhibited [109]. Bicarbonate movement into the cell can occur via CFTR, which is permeable to chloride and bicarbonate ions and activated by cAMP [110]. Mutations of CFTR not associated with congenital absence of the vas deferens have been observed in healthy men with decreased sperm count or quality [99]. Lower CFTR levels decrease the ability of sperm to undergo membrane hyperpolarization and cAMP production in response to bicarbonate, resulting in lower motility and subfertility [99]. Calcium Signaling An intracellular increase in calcium concentration is required for sperm hypermotility and the acrosome reaction, and the primary regulator of calcium influx in sperm is CatSper. CatSper is a sperm-specific, low voltage-dependent calcium channel that is essential for male fertility [102]. The channel is composed of multiple

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subunits, including four pore-forming channel proteins (CatSper 1–4), each of which contains six transmembrane domains and three auxiliary subunits (beta, gamma, and delta) [95, 111–114]. The pH-sensitive domain of CatSper1 is in the intracellular N-terminal His-rich domain [115, 116]. Disruption of CatSper2 may result in asthenozoospermia or immotile sperm and failed hyperactivated motility, whereas disruption of CatSper3 and CatSper4 may have implications in allowing sperm to undergo the acrosome reaction [102]. Because of the significant impact of CatSper mutations on sperm motility, CatSper has been the target of potential male contraceptives. Antibodies specific for the transmembrane domain and pore regions of CatSper1 resulted in reduced fertility in mice [102, 117]. Additionally, drugs such as calcium channel blockers have been studied to inhibit the actions of CatSper, preventing the calcium influx required for sperm hypermotility [102]. Membrane Polarization The sperm membrane potential is typically maintained by Na+/K+ ATPase, and the α4 subunit is specific to male germ cells [99, 118]. Mice lacking Atp1α4, which encodes the α4 subunit, are infertile, and sperm from these mice are morphologically abnormal (bent at the midpiece) and have depolarized membrane potentials [99]. The α4 subunit is thought to maintain intracellular sodium concentrations, which, in turn, affect both intracellular pH and calcium concentrations. Membrane hyperpolarization requires efflux of potassium ions and this occurs primarily via KSper activation. KSper is a voltage-gated potassium channel and activated by intracellular alkalinization and calcium [99, 119]. KSper is thought to function downstream of CatSper due to the requirement of intracellular calcium for KSper activation [99]. Mice lacking SLO3, the murine form of KSper, are infertile due to impaired hyperpolarization during capacitation. Mutation of these ion channels may not impair sperm production (therefore, sperm count may be normal). However, the inability of sperm to undergo capacitation and the acrosome reaction renders sperm incapable of fertilization. Thus, understanding the physiology and function of these ion channels in sperm capacitation remains a focus for the development of treatments to improve a subset of men with infertility, as well as for the design of nonhormonal male contraceptive agents.

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Key Points • Capacitation is the change in sperm motility to produce hyperactivated movement, and occurs in the female genital tract (uterotubal junction). • Capacitation is associated with sperm plasma membrane alteration and intracellular pH and ion concentration changes. • Molecular processes underlying capacitation include intracellular alkalinization, membrane hyperpolarization, and calcium ion influx and signaling. • Various channels, including CatSper and KSper, are involved in sperm capacitation.

Epididymal Dysfunction and Male Infertility Epididymal Disorders and Male Infertility Various disorders of the epididymis can result in male infertility. These include congenital defects of the epididymis, infection or inflammation of the epididymis, neoplasms, and obstruction. Infertility in men with a history of epididymal disorders can be due to obstruction. In men with normal spermatogenesis, assisted reproductive technologies can be employed, with sperm harvested from either the testis or a normal area of the epididymis. This section highlights disorders of the epididymis that may relate to male infertility, and management of epididymal-related infertility.

Congenital Defects of the Epididymis Congenital defects of the epididymis have been noted in patients with congenital absence of the vas deferens and cryptorchidism. Defects in the development of the epididymis may contribute to obstructive and nonobstructive causes of male infertility. Congenital Absence of the Vas Deferens Men with congenital unilateral or bilateral absence of the vas deferens (CUAVD/CBAVD) can have varying degrees of epididymal abnormalities, ranging from mild epididymal abnormalities with a mostly intact epididymis to moderate abnormalities with absence of the cauda or severe abnormalities with absence of the distal caput, corpus, and cauda (Fig. 3.3) [120]. The caput is usually

Chapter 3: Maturation and Function of Sperm

identified are similar to the previously described classification [125, 128]. Marshall and Shermeta found that 33 percent of their patients had atresia of a portion of the epididymis, along with loss of continuity of the epididymis with the testis [128]. The severity of impact of these defects on fertility potential is likely. However, additional studies are needed to determine the overall contribution of epididymal abnormality in infertility in patients with cryptorchidism. In addition to issues with fertility, abnormal attachment of the epididymis to the testis can result in torsion and infarction of the epididymis [126].

Epididymitis and Other Inflammatory Conditions of the Epididymis

Fig. 3.3 Congenital unilateral absence of the vas deferens. Vasogram depicting the absence of the right vas deferens. (Imaged provided by Dr. Marc Goldstein.)

present in patients with CBAVD [121]. The etiology for malformation of the epididymis in CBAVD is thought to be due to in utero epididymal obstruction from dehydrated secretions or mesonephric developmental abnormalities [122]. Spermatogenesis is largely normal in cases of isolated CBAVD and sperm can be recovered surgically and used for assisted reproductive technology [123, 124]. Cryptorchidism Structural abnormalities of the epididymis are observed in 32–61 percent of patients with cryptorchidism at the time of orchiopexy for cryptorchid testis or testicular torsion [125–128]. Heath et al. classified these abnormalities as type A (complete absence of the vas and epididymis in the presence of a normal testis) and type B (abnormal attachment of the epididymis and testis) [129]. Type B abnormalities can be subdivided into four anatomic findings: group 1, normal attachment of the head and tail, with loose attachment of the body (noted as a looped epididymis in some studies) (48%); group 2, normal attachment of the head and epididymis, with a detached body and tail (40%); group 3, normal attachment of the tail, with a detached head (4%); group 4, near-complete separation of the testis and epididymis (7%) [129]. Other studies have divided these epididymal anomalies into different categories, but the abnormalities

Epididymitis is a common urologic problem that can be associated with high morbidity [42, 130, 131]. Epididymitis, or inflammation of the epididymis, can be due to infectious or noninfectious causes, and can affect males of all ages. Although the etiologies differ, patient symptoms are typically unilateral scrotal pain and swelling [42]. However, patients with epididymitis can be asymptomatic or have other symptoms, including pelvic pain, scrotal masses or nodules, irritative urinary symptoms, fever, purulent ejaculate, urethral discharge, and infertility [42, 132]. Rare sequelae of epididymitis are fistula, abscess, hematospermia, systemic infection, and sepsis [42, 132, 133].

Infectious Epididymitis Retrograde ascent from the urinary tract into the male genital tract via the ejaculatory ducts and vas deferens is thought to be the route of entry for urinary pathogens to gain access to the epididymis [42, 131, 134]. In men younger than 35 years, epididymitis is contracted most commonly by sexual intercourse. The most common pathogens in this population are Chlamydia trachomatis and Neisseria gonorrhoeae [42, 135]. Men with sexually transmitted epididymitis typically present with urethritis, dysuria, urethral irritation, and mucopurulent urethral discharge, although a large percentage of men may be asymptomatic. In men older than 35 years, Gramnegative organisms are more commonly the cause of epididymitis [42, 135]. In these patients, Escherichia coli, a common urinary pathogen, can cause epididymitis when patients have urinary obstruction and incomplete bladder emptying. Higher voiding pressures in the setting of urinary obstruction can result in reflux of urine, which

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may be infected, into the vas deferens and epididymis, which may result in vasitis and/or epididymitis [42]. Other bacterial pathogens frequently found to be the cause of epididymitis include Klebsiella species (spp.), Proteus spp., and Pseudomonas aeruginosa [42]. Treatment of infectious epididymitis includes appropriate use of antimicrobial agents against the specific pathogen(s) [42, 135, 136]. Rare cases of severe epididymitis can lead to sepsis, formation of abscesses or fistulae, or ischemia/infarction of the testis [132, 133, 137].

Epididymitis in the Pediatric Population Pediatric epididymitis demonstrates a bimodal age distribution, with the disease occurring in young boys (under 5 years of age) and prepubertal boys [138]. In boys less than five years of age, anatomical defects are strongly associated with epididymitis [136, 139]. Genitourinary defects, including posterior urethral valves, urethral strictures or hypoplasia, may result in bladder outlet obstruction and high bladder pressure voiding with subsequent reflux of urine into the ejaculatory duct, vas deferens, and epididymis. In prepubertal boys, dysfunctional voiding, including bladder hyperactivity and detrusor-sphincter dyssynergia, may result in high bladder voiding pressures and reflux epididymitis [136, 140]. Venereal pathogens are less commonly observed, although can be seen in sexually active children or importantly in cases of sexual abuse. More common bacteria that can cause pediatric epididymitis include E. coli and Haemophilus influenzae, and less commonly Salmonella and Pseudomonas [136]. Additionally, with the decline of vaccination, mumps epididymo-orchitis, caused by a member of the Paramyxovirus family, is now more commonly seen [140]. Infection with the mumps virus is characterized by orchitis and parotiditis. However, it can also affect the prostate, central nervous system, pancreas, heart, joints, and liver. The disease is typically self-limited and involves supportive care [141]. Pediatric epididymitis typically presents as an acute scrotum. Severe scrotal pain due to epididymitis must be distinguished from testicular torsion, which is a surgical emergency. Epididymitis has been reported to occur almost as frequently as testicular torsion [142, 143]. Physical examination and scrotal ultrasonography with color Doppler can help distinguish epididymitis from testicular torsion. Doppler allows for evaluation of the vasculature, and the appearance of epididymitis on scrotal ultrasonography typically demonstrates a hypervascular epididymis [144]. Infectious epididymitis in the

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pediatric population should be treated with appropriate antimicrobial therapy, whereas noninfectious or viral epididymitis is treated with supportive care.

Rare Infectious Causes of Epididymitis Other rare infectious causes of epididymitis include opportunistic infections of the epididymis in immunocompromised individuals, tuberculous epididymitis, and parasitic epididymitis. Opportunistic infections of the epididymis can occur in patients who are immunocompromised. Fungal, bacterial, and viral pathogens, including Candida, Coccidioides, Blastomyces, Actinomyces, Histoplasma, Aspergilla, Nocardia, and cytomegalovirus, have been reported in the literature [145, 146]. Epididymitis in this setting can occur, as described previously, through a retrograde route from the urinary tract, although these pathogens also may disseminate to the epididymis hematogenously. Aggressive treatment with antimicrobial agents is necessary in this patient population, as opportunistic epididymitis may be an early sign of systemic infection which may have increased morbidity and mortality in this patient population [147]. Tuberculous epididymitis is a rare phenomenon and often is usually associated with concomitant infection of the genitourinary tract. Spread of mycobacteria to the epididymis can occur via lymphatics, as well as via the bloodstream. Iatrogenic tuberculous epididymitis can occur when patients with bladder cancer receive intravesical instillations of bacille Calmette–Guérin (BCG) [148, 149]. Parasitic epididymitis is rarely seen in the United States but can be due to schistosomiasis, enterobiasis, trypanosomiasis, leishmaniasis, and filariasis [150–154]. Treatment of tuberculous and parasitic epididymitis involves administration of appropriate antimicrobial or antifungal agents.

Rare Noninfectious Causes of Epididymitis Other noninfectious forms of epididymitis include druginduced, iatrogenic, traumatic, and stress-induced epididymitis [147]. Drug-induced epididymitis has been reported in patients using amiodarone, an antiarrhythmic drug [155, 156]. Histologic examination of the epididymis demonstrates infiltration of inflammatory cells as well as crystals within histiocytes which may be deposits of amiodarone metabolites [156]. Drug concentrations in the epididymis are 25- to 400-fold higher than that detected in serum, and the effects of this drug on the epididymis may be dose- and

Chapter 3: Maturation and Function of Sperm

duration-dependent [157, 158]. Treatment involves dose adjustment or discontinuation of amiodarone [157]. Iatrogenic epididymitis can occur in patients undergoing genitourinary procedures, including intermittent or indwelling urethral catheterization, cystoscopy, or prostate biopsy or surgery [159–162]. Traumatic epididymitis may occur when iatrogenic injury to the epididymis occurs during scrotal surgery or scrotal trauma. Physical examination typically reveals a swollen and tender epididymis, and ultrasonography with color Doppler reveals epididymal hyperemia [163]. Patients with traumatic epididymitis can develop chronic epididymitis or infertility due to obstruction of epididymal tubules. Conservative management with scrotal support, ice, elevation, and rest often improves symptoms. If superinfection is suspected, antibiotics can be administered. Stress-induced epididymitis can occur when a man undergoes physical stress such as performing heavy lifting or other strenuous activities [164]. The mechanism of disease is still thought to involve reflux of urine through the vas deferens to the epididymis, resulting in inflammation. Treatment involves use of conservative management with anti-inflammatory medications, ice, scrotal support, and rest.

Epididymal Neoplasms Primary malignancy of the epididymis is a rare phenomenon, and tumors account for 0.03 percent of male cancers [165]. The majority of epididymal tumors are benign. Given the cellular composition of the epididymis, the most likely malignancy is adenocarcinoma; other types include sarcomas, germ cell tumors, lymphoma, and plasmacytoma, with the latter two often associated with metastatic disease [166]. Adenocarcinoma has metastatic potential, with spread to regional lymph nodes, as well as to visceral organs [167]. The most common benign subtype of epididymal neoplasm are adenomatoid tumors, which are of mesothelial origin and are differentiated from adenocarcinomas by histopathology [168]. Other benign tumors include papillary cystadenomas as part of von Hippel– Lindau (VHL) disease, and these tumors have clear cells similar to those seen in clear cell renal cell carcinoma [166]. Lastly, metastasis to the epididymis is extremely rare but has been detected in some genitourinary (prostate and kidney) and gastrointestinal malignancies [169].

Fig. 3.4 Epididymal obstruction. Photograph depicting dilated epididymal tubules. (Image provided by Dr. Marc Goldstein.)

conditions such as tuberculosis) or trauma, as noted previously, and are generally only recognized during workup and treatment of an infertile male [170, 171]. Amongst obstructive men with azoospermia, epididymal obstruction affects up to 67 percent of those with folliclestimulating hormone (FSH) concentrations less than twice the upper limit of normal [172]. Iatrogenic injuries may occur during procedures in childhood, including herniorrhaphy or hydrocelectomy, or during any scrotal procedure such as exploration for testicular torsion [173]. Many cases of iatrogenic obstruction may be missed if only occurring unilaterally, as a functional contralateral side may compensate for the obstruction. Congenital obstruction, on the other hand, occurs most commonly in the head and body of the epididymis and/or may consist of atresia or agenesis of a segment of the epididymis [172, 174]. This may be diagnosed preoperatively during physical examination of the epididymis with fullness or with radiographic findings (ultrasound or magnetic resonance imaging (MRI)) of abnormal epididymal structures such as agenesis or dilation [175]. Otherwise diagnosis may occur at the time of surgical intervention (Fig. 3.4), and options for sperm retrieval in these men would include surgical sperm aspiration or retrieval from the testis or epididymis, or microsurgical reconstruction of the vas deferens to a portion of unobstructed epididymis [176].

Epididymal Obstruction

Management of Epididymal-Related Male Infertility

Obstruction of the epididymis may occur idiopathically, iatrogenically, or from infection (epididymitis or rare

Sperm may be retrieved surgically from the epididymis either percutaneously or by using a surgical operating

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Fig. 3.5 Epididymal sperm aspiration techniques. Illustration depicting microsurgical (MESA) and percutaneous (PESA) epididymal sperm aspiration. PESA is performed through the skin without any skin incision. Depicted is the needle piercing the skin and entering the caput epididymis where epididymal fluid is retrieved. MESA is performed through a skin incision and using a standard operating microscope. A single tubule is identified and incised. Fluid is retrieved directly from the visualized tubule using a pipette.

microscope (Fig. 3.5). Percutaneous epididymal sperm aspiration (PESA) often is completed in the clinic with local anesthesia and offers a minimally invasive procedure with a short recovery period [177]. Retrieval rates with PESA in obstructive azoospermia are 61–96 percent [178, 179]. However, this procedure has high complication rates and may result in long-term epididymal obstruction [177]. Microsurgical epididymal sperm aspiration (MESA) is performed in the operating room under general anesthesia and requires a standard operating microscope and a surgeon with experience and training in microsurgery. Retrieval rates with MESA are higher than those seen with PESA, with success of sperm retrieval in 96–100 percent of patients [180, 181]. Given direct visualization of the tubules during the procedure, MESA is associated with fewer complications and reduced risk of future epididymal obstruction (Fig. 3.5) [181]. As sperm mature and develop along the course of the epididymis, the quality of surgically retrieved sperm varies. Aspiration from the cauda yields poorer-quality sperm with debris and macrophages, but with better motility [182]. Retrieval generally begins in the cauda and toward the caput until appropriate sperm are

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retrieved [182]. There are some data to suggest that epididymal sperm may have higher rates of abnormal DNA fragmentation, compared to sperm in the testis, as measured by terminal deoxynucleotidyl transferase deoxyuridine triphosphate nick end labeling (TUNEL) assay [183]. With the exception of increased DNA fragmentation, limited data exist with regard to assisted reproductive techniques and epididymal sperm retrieval. As previously mentioned, if there is suspected epididymal obstruction, an attempt can be made to microsurgically reconstruct the connection between the vas deferens and an unobstructed tubule within the epididymis (vaso-epididymostomy). Success results in normal sperm transport and men can conceive naturally after successful vaso-epididymostomy. Typically, the time to patency after vaso-epididymostomy is 2.8–6.6 months [184]. This procedure is technically challenging, and even in the hands of skilled microsurgeons, the mean patency rate is approximately 64 percent. These, however, may range from 31 to 92 percent, and pregnancy rates in the range of 10–50 percent [185, 186]. Given the structural differences along the epididymis, it has been suggested that reconstruction closer to the caput generally results in poorer fertility outcomes [6].

The Epididymis as a Contraceptive Target Given its unique role in sperm maturation and storage, the epididymis is an optimal target for male contraceptives. However, despite numerous attempts, no male contraceptive targeting the epididymis has been successfully developed. Possible targets have included: (1) alteration of peritubular epididymal contraction to reduce sperm transit time, preventing optimization throughout the organ, (2) direct attack of sperm and its function with sperm inhibitors, and (3) modification of the luminal fluid within the epididymal tubule [187]. As more research is conducted on the epididymis and its role in sperm transport and maturation, this information may be used toward the development of an effective male contraceptive in the future.

Key Points • Congenital epididymal defects are observed in men with congenital absence of the vas deferens and cryptorchidism. • Epididymitis can be infectious or noninfectious, and can result in epididymal obstruction requiring

Chapter 3: Maturation and Function of Sperm

sperm retrieval or reconstruction of the male reproductive tract for fertility. • Epididymal-related infertility can be managed with minimally invasive procedures, including PESA or MESA, and sperm retrieval rates are generally good. • The epididymis may be a future target of male contraceptive development.

Sperm maturation and transport are a complex and highly regulated process. While sperm is created in the testis, its maturation occurs predominantly in the epididymis and female genital tract. Much work has been done to elucidate the functions of various sperm and nonsperm factors required for normal sperm maturation and capacitation. However, more work is needed to elucidate this complex biologic process.

Take-Home Messages • •

Sperm formation occurs in the testis, and maturation occurs in the epididymis and female genital tract. Each section of the epididymis displays different and critical roles for sperm maturation. Capacitation, or acquisition of sperm hypermotility, occurs in the female genital tract due to various ion channels within the sperm plasma membrane.

Further Reading Cornwall GA. New insights into epididymal biology and function. Hum Reprod Update 2009;15: 213–27. James ER, Carrell DT, Aston KI, Jenkins TG, Yeste M, Salas-Huetos A. The role of the epididymis and the contribution of epididymosomes to mammalian reproduction. Int J Mol Sci 2020;21:5377. Stival C, Puga Molina Ldel C, Paudel B, Buffone MG, Visconti PE, Krapf D. Sperm capacitation and acrosome reaction in mammalian sperm. Adv Anat Embryol Cell Biol 2016;220:93–106.

•

•

Capacitation requires intracellular alkalinization, sperm plasma membrane hyperpolarization, and calcium influx and signaling. Various epididymal-related disorders may result in male infertility requiring surgical reconstruction of the male genital tract or sperm retrieval for future fertility. The epididymis remains an area of continued study and a potential target for therapeutics and nonhormonal male contraceptives.

Acknowledgments

Conclusion

•

•

The authors would like to acknowledge Vanessa Dudley for her outstanding illustrations. C.K. and N.P. are supported by the Frederick J. and Theresa Dow Wallace Fund of the New York Community Trust. D.J.L. is supported, in part, by the National Institutes of Health (NIH), National Institute of Kidney and Digestive Diseases (1R01DK078121), the Eunice Kennedy Shriver National Institute of Child Health and Human Development (1U54HD100549–01, 1R01HD095341 and 5P01HD087157), and the Frederick J. and Theresa Dow Wallace Fund of the New York Community Trust. D.J.L. serves on the Scientific Advisory Board of Roman Health (Scientific Advisory Board and Consultant; equity and compensation) Fellow (equity) and her travel was supported, in part, by the World Health Organization to attend an editorial board meeting of the organization. She serves as Secretary-Treasurer of the American Board of Bioanalysis.

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micropuncture with perivascular nerve stimulation for intracytoplasmic sperm injection to treat unreconstructable obstructive azoospermia. Arch Androl 1996;36:217–24. 125. Johansen TE. Anatomy of the testis and epididymis in cryptorchidism. Andrologia 1987;19:565–9. 126. Brisson P, Feins N, Patel H. Torsion of the epididymis. J Pediatr Surg 2005;40:1795–7. 127. Favorito LA, Riberio Julio-Junior H, Sampaio FJ. Relationship between undescended testis position and prevalence of testicular appendices, epididymal anomalies, and patency of processus vaginalis. Biomed Res Int 2017;2017:5926370. 128. Marshall FF, Shermeta DW. Epididymal abnormalities associated with undescended testis. J Urol 1979;121:341–3. 129. Heath AL, Man DW, Eckstein HB. Epididymal abnormalities associated with maldescent of the testis. J Pediatr Surg 1984;19:47–9. 130. Nicholson A, Rait G, MurrayThomas T, Hughes G, Mercer CH, Cassell J. Management of epididymo-orchitis in primary care: results from a large UK primary care database. Br J Gen Pract 2010;60: e407–22. 131. Campbell MF. Gonococcus epididymitis: observations in three thousand cases from the urological services of Bellevue Hospital. Ann Surg 1927;86:577–90. 132. Kaver I, Matzkin H, Braf ZF. Epididymo-orchitis: a retrospective study of 121 patients. J Fam Pract 1990;30:548–52. 133. Slavis SA, Kollin J, Miller JB. Pyocele of scrotum: consequence of spontaneous rupture of testicular abscess. Urology 1989;33:313–16. 134. Berger RE. Acute epididymitis: etiology and therapy. Semin Urol 1991;9:28–31. 135. Berger RE, Alexander ER, Harnisch JP, et al. Etiology, manifestations and

therapy of acute epididymitis: prospective study of 50 cases. J Urol 1979;121:750–4. 136. Hagley M. Epididymo-orchitis and epididymitis: a review of causes and management of unusual forms. Int J STD AIDS 2003;14:372–7; quiz 8. 137. Gould SW. Epididymo-orchitis: a rare, fatal, intra-abdominal cause. Ann R Coll Surg Engl 1996;78:230. 138. Likitnukul S, McCracken GH Jr, Nelson JD, Votteler TP. Epididymitis in children and adolescents. A 20year retrospective study. Am J Dis Child 1987;141:41–4. 139. Merlini E, Rotundi F, Seymandi PL, Canning DA. Acute epididymitis and urinary tract anomalies in children. Scand J Urol Nephrol 1998;32:273–5.

Aspergillus fumigatus in a patient with AIDS. Clin Infect Dis 1998;26:229–31. 147. Seçil M, Göktay AY, Dicle O, Yörükoğlu K. Bilateral epididymal Candida abscesses: sonographic findings and sonographically guided fine-needle aspiration. J Clin Ultrasound 1998;26:413–15. 148. Singh D, Fontanella M, Voci S. Tuberculous epididymitis. Ultrasound Q 2012;28:145–7. 149. O’Connell HE, Russell JM, Schultz TC. Delayed epididymitis following intravesical bacillus Calmette– Guerin administration. Aust N Z J Surg 1993;63:70–2.

140. Bukowski TP, Lewis AG, Reeves D, Wacksman J, Sheldon CA. Epididymitis in older boys: dysfunctional voiding as an etiology. J Urol 1995;154(2 Pt 2):762–5.

150. Gelfand M, Ross CM, Blair DM, Castle WM, Weber MC. Schistosomiasis of the male pelvic organs. Severity of infection as determined by digestion of tissue and histologic methods in 300 cadavers. Am J Trop Med Hyg 1970;19:779–84.

141. Singh R, Mostafid H, Hindley RG. Measles, mumps and rubella – the urologist’s perspective. Int J Clin Pract 2006;60:335–9.

151. Kollias G, Kyriakopoulos M, Tiniakos G. Epididymitis from Enterobius vermicularis: case report. J Urol 1992;147:1114–16.

142. Caldamone AA, Valvo JR, Altebarmakian VK, Rabinowitz R. Acute scrotal swelling in children. J Pediatr Surg 1984;19:581–4.

152. Shiadeh MN, Niyyati M, Fallahi S, Rostami A. Human parasitic protozoan infection to infertility: a systematic review. Parasitol Res 2016;115:469–77.

143. Tanaka K, Ogasawara Y, Nikai K, Yamada S, Fujiwara K, Okazaki T. Acute scrotum and testicular torsion in children: a retrospective study in a single institution. J Pediatr Urol 2020;16:55–60. 144. Boettcher M, Bergholz R, Krebs TF, et al. Differentiation of epididymitis and appendix testis torsion by clinical and ultrasound signs in children. Urology 2013;82:899–904. 145. Docimo SG, Rukstalis DB, Rukstalis MR, Kang J, Cotton D, DeWolf WC. Candida epididymitis: newly recognized opportunistic epididymal infection. Urology 1993;41:280–2. 146. Hood SV, Bell D, McVey R, Wilson G, Wilkins EG. Prostatitis and epididymo-orchitis due to

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153. Carvalho T, Trindade S, Pimenta S, Santos AB, Rijo-Ferreira F, Figueiredo LM. Trypanosoma brucei triggers a marked immune response in male reproductive organs. PLoS Negl Trop Dis 2018;12:e0006690. 154. Williams PB, Henderson RJ, Sanusi ID, Venable DD. Ultrasound diagnosis of filarial funiculoepididymitis. Urology 1996;48:644–6. 155. Hutcheson J, Peters CA, Diamond DA. Amiodarone induced epididymitis in children. J Urol 1998;160:515–17. 156. Shen Y, Liu H, Cheng J, Bu P. Amiodarone-induced epididymitis: a pathologically confirmed case report

Chapter 3: Maturation and Function of Sperm

and review of the literature. Cardiology 2014;128:349–51.

update. Biomed Res Int 2017;2017:4126740.

157. Nikolaou M, Ikonomidis I, Lekakis I, Tsiodras S, Kremastinos D. Amiodarone-induced epididymitis: a case report and review of the literature. Int J Cardiol 2007;121: e15–6.

168. Cazorla A, Algros MP, Bedgedjian I, Chabannes E, Camparo P, ValmaryDegano S. Epididymal leiomyoadenomatoid tumor: a case report and review of literature. Curr Urol 2014;7:195–8.

158. Greene HL, Graham EL, Werner JA, et al. Toxic and therapeutic effects of amiodarone in the treatment of cardiac arrhythmias. J Am Coll Cardiol 1983;2:1114–28.

169. Algaba F, Santaularia JM, Villavicencio H. Metastatic tumor of the epididymis and spermatic cord. Eur Urol 1983;9:56–9.

159. Donzella JG, Merrick GS, Lindert DJ, et al. Epididymitis after transrectal ultrasound-guided needle biopsy of prostate gland. Urology 2004;63:306–8. 160. Hoffelt SC, Wallner K, Merrick G. Epididymitis after prostate brachytherapy. Urology 2004;63:293–6.

170. Peng J, Yuan Y, Cui W, et al. Causes of suspected epididymal obstruction in Chinese men. Urology 2012;80:1258–61. 171. Gupta R, Singh P, Kumar R. Should men with idiopathic obstructive azoospermia be screened for genitourinary tuberculosis? J Hum Reprod Sci 2015;8: 43–7.

161. Perrouin-Verbe B, Labat JJ, Richard I, Mauduyt de la Greve I, Buzelin JM, Mathe JF. Clean intermittent catheterisation from the acute period in spinal cord injury patients. Long term evaluation of urethral and genital tolerance. Paraplegia 1995;33:619–24.

172. Dohle GR, Colpi GM, Hargreave TB, et al. EAU guidelines on male infertility. Eur Urol 2005;48:703–11.

162. Moss WM. A comparison of openend versus closed-end vasectomies: a report on 6220 cases. Contraception 1992;46:521–5.

174. Breeland E, Cohen MS, Warner RS, Leiter E. Epididymal obstruction in azoospermic males. Infertility 1981;4:49–66.

163. Gordon LM, Stein SM, Ralls PW. Traumatic epididymitis: evaluation with color Doppler sonography. AJR Am J Roentgenol 1996;166:1323–5.

175. Ammar T, Sidhu PS, Wilkins CJ. Male infertility: the role of imaging in diagnosis and management. Br J Radiol 2012;85:S59–68.

164. Sawyer EK, Anderson JR. Acute epididymitis: a work-related injury? J Natl Med Assoc 1996;88:385–7.

176. Berardinucci D, Zini A, Jarvi K. Outcome of microsurgical reconstruction in men with suspected epididymal obstruction. J Urol 1998;159:831–4.

165. Yeung CH, Wang K, Cooper TG. Why are epididymal tumours so rare? Asian J Androl 2012;14:465–75. 166. Elsasser E. Tumors of the epididymis. Recent Results Cancer Res 1977;60:163–75. 167. Zou ZJ, Xiao YM, Liu ZH, et al. Clinicopathological characteristics, treatment, and prognosis of rarely primary epididymal adenocarcinoma: a review and

173. Hopps CV, Goldstein M. Microsurgical reconstruction of iatrogenic injuries to the epididymis from hydrocelectomy. J Urol 2006;176:2077–9; discussion 80.

intracytoplasmic sperm injection. Hum Reprod Update 1998;4:57–71. 179. Lin YM, Hsu CC, Kuo TC, Lin JS, Wang ST, Huang KE. Percutaneous epididymal sperm aspiration versus microsurgical epididymal sperm aspiration for irreparable obstructive azoospermia – experience with 100 cases. J Formos Med Assoc 2000;99:459–65. 180. Schlegel PN, Palermo GD, Alikani M, et al. Micropuncture retrieval of epididymal sperm with in vitro fertilization: importance of in vitro micromanipulation techniques. Urology 1995;46:238–41. 181. Bernie AM, Ramasamy R, Stember DS, Stahl PJ. Microsurgical epididymal sperm aspiration: indications, techniques and outcomes. Asian J Androl 2013;15:40–3. 182. Esteves SC, Miyaoka R, Orosz JE, Agarwal A. An update on sperm retrieval techniques for azoospermic males. Clinics (Sao Paulo) 2013;68:99–110. 183. Hammoud I, Bailly M, Bergere M, et al. Testicular spermatozoa are of better quality than epididymal spermatozoa in patients with obstructive azoospermia. Urology 2017;103:106–11. 184. Farber NJ, Flannigan R, Srivastava A, Wang H, Goldstein M. Vasovasostomy: kinetics and predictors of patency. Fertil Steril 2020;113:774–80 e3. 185. Yoon YE, Lee HH, Park SY, et al. The role of vasoepididymostomy for treatment of obstructive azoospermia in the era of in vitro fertilization: a systematic review and meta-analysis. Asian J Androl 2018;21:67–73.

177. Coward RM, Mills JN. A step-bystep guide to office-based sperm retrieval for obstructive azoospermia. Transl Androl Urol 2017;6:730–44.

186. Chan PT. The evolution and refinement of vasoepididymostomy techniques. Asian J Androl 2013;15:49–55.

178. Meniru GI, Gorgy A, Batha S, Clarke RJ, Podsiadly BT, Craft IL. Studies of percutaneous epididymal sperm aspiration (PESA) and

187. Hinton BT, Cooper TG. The epididymis as a target for male contraceptive development. Handb Exp Pharmacol 2010;198:117–37.

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Section 1 Chapter

4

Scientific Foundations of Male Infertility

The Male Reproductive Endocrine System Ettore Caroppo

Introduction The male reproductive endocrine system function is strictly dependent on the dynamic interplay between neural and hormonal signals originating from the hypothalamus where specific neurons secrete gonadotropinreleasing hormone (GnRH) in an episodic pattern of pulses under the control of excitatory and inhibitory signals from neuromodulators, the anterior pituitary where GnRH binds to its own receptors on a specific pituitary cell type to stimulate pituitary gonadotropin secretion, and the testes where the trophic actions of gonadotropins result in the promotion of spermatogenesis and secretion of testicular steroids and peptides, which, in turn, modulate hypothalamic and pituitary function in both positive and negative feedback loops.

Experiments in rats and mice have shown that individual adult GnRH neurons exhibit a range of different spontaneous firing patterns: they include silent, continuously active, or irregularly active neurons, including what are commonly called bursting cells (Fig. 4.1). Such variability in firing behavior has been demonstrated also in vivo, as well as in cultured embryonic GnRH neurons [1]. Studies on adult and embryonic GnRH neurons indicate that most burst firing in GnRH neurons consists of 2–5 action potentials occurring over about 2 seconds, with interburst interval of 20 seconds or longer. The firing of individual, or bursts of, action potentials in

Gonadotropin-Releasing Hormone The mechanism(s) involved in the regulation of GnRH secretion are far from being completely understood, due to the anatomical complexity of the system. Measuring GnRH secretion in vivo is not possible in humans and is challenging in mice. Several studies have therefore been performed on immortalized hypothalamic neurons or on acute brain slices for electrophysiological recordings. However, to what extent such models could reproduce the in vivo physiology remains to be determined. The data presented herein are therefore not definitive and open to further refinement. GnRH is a decapeptide secreted in a pulsatile fashion by GnRH neurons (approximately 1200 in number in humans), the cell bodies of which are relatively scattered across the mediobasal hypothalamus and preoptic area. Once secreted, GnRH is released into the hypophyseal portal system in intermittent pulses, the frequency and amplitude of which are crucial to maintaining gonadotrope responsiveness to GnRH itself (so-called selfpriming effect).

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Fig. 4.1 Typical firing patterns of adult rodent GnRH neurons in vitro (top) and in vivo (bottom). The top three traces show the typical bursting, continuous, and silent types of firing exhibited by adult GnRH neurons in an acute brain slice preparation using the cell-attached recording mode. The bottom two recordings show cell-attached firing patterns from GnRH neurons in vivo in an anesthetized adult mouse. Each deflection, up or down, represents a single action potential. (Reproduced with permission from Herbison A. Physiology of the adult gonadotropin-releasing hormone neuronal network. In: TM Plant and AJ Zeleznik, eds. Knobil and Neill’s Physiology of Reproduction, 4th ed. Academic Press, 2015; pp. 399–467.)

Chapter 4: The Male Reproductive Endocrine System

spontaneously active GnRH neurons is followed by hyperpolarization that lasts from several milliseconds to several seconds and then by subthreshold afterdepolarization potential and significant reduction of the frequency of action potential firing. Modulation of calcium influx by GnRH mediates firing frequency. GnRH receptors expressed in hypothalamic GnRH neurons are important modulators of their neuronal excitability [2]. GnRH pulsatile secretion is tightly regulated by autocrine, paracrine, and endocrine mechanisms.

Autocrine Regulation of GnRH Secretion A number of studies suggest that GnRH is able to regulate its own release by means of ultrashort loop negative feedback. This hypothesis was supported by the demonstration of expression of functional GnRH receptors (GnRHRs) in immortalized hypothalamic neurons [3]. A study on brain slices from castrated adult male mice further demonstrated that low-dose GnRH significantly decreased, whereas higher doses increased, the firing rate in those GnRH neurons expressing GnRHRs [4]. In particular, the reduction in firing rate by a low-dose GnRH signal supported its ultrashort inhibitory action upon its own release. In addition, using the GnRH agonist buserelin, basal GnRH secretion, its promoter activity, and mRNA levels were decreased in immortalized GT1–1 cells [5]. The pulsatile secretion of GnRH is highly calcium-dependent and stimulated by cyclic adenosine monophosphate (cAMP). GnRH itself has been found to modulate calcium influx and consequently mediate the firing frequency and spike profile [6]. Notably, GnRHR expression, GnRH-dependent activation of calcium signaling, and autocrine regulation of GnRH release are prerogatives of early fetal GnRH neurons. Taken together, these evidence suggest that GnRH neurons could activate and suppress their own activity using GnRH itself as an intra-GnRH neural network signal. Such strategy may have important implications for the generation of GnRH surge and regulation of its pulsatile release.

Paracrine Regulation Although pulsatile secretion of GnRH release is an intrinsic property of GnRH neurons due to their spontaneous electrical activity, there are other regulatory mechanisms able to sustain this activity that is crucial to reproduction. Amongst others, a potential role for kisspeptin signaling in the generation of pulsatile hormone secretion has been

postulated, since humans with global deletions or mutations in kisspeptin or its receptor fail to exhibit normal pulsatile luteinizing hormone (LH) secretion [7]. Discovery of kisspeptin reproductive function was made from a study on consanguineous families that had members with idiopathic hypogonadotrophic hypogonadism who were homozygous for a “L148S” mutation in the GPR54 gene – otherwise known as Kiss1r. A mouse model deficient in Kiss1r was further created, with mice expressing a hypogonadotrophic hypogonadism phenotype; however, since exogenous administration of GnRH restored gonadal function, it was hypothesized that kisspeptin could play a role on GnRH release [8]. In humans, kisspeptin neurons are found in the infundibular nucleus and preoptic area. Kisspeptin seems to be essential for normal timing of puberty. Indeed, loss-of-function mutations lead to the absence of pubertal development [9]. Moreover, the electrophysiological properties of GnRH neurons in response to kisspeptin differ with age; significant depolarization of more than 90 percent of GnRH neurons is observed in adult mice, whereas only 27 and 40 percent of GnRH neurons in juvenile and prepubertal mice respond to kisspeptin administration, which suggests that the number of kisspeptin-responsive GnRH neurons increases during pubertal development [10]. Almost all kisspeptin neurons in the arcuate nucleus (ARC) coexpress the tachykinin neurokinin B (NKB), as well as the endogenous opioid peptide (EOP) dynorphin. NKB is a key regulator of fertility, since mutations in the genes encoding NKB (TAC3) or its receptor (TAC3R) have been found to lead to defects in GnRH release and subsequent hypogonadism [11]. Kisspeptin/ neurokinin B/dynorphin (KNDy) neurons in the infundibular/arcuate nucleus influence the activity of GnRH by acting on both GnRH cell bodies and neurosecretory terminals, making direct contact with GnRH cell bodies, to produce coordinated and pulsatile GnRH secretion. The stimulatory role of NKB and the inhibitory action of dynorphin coordinate the pulsatile release of kisspeptin, which, in turn, drives the pulsatile secretion of GnRH and LH [12]. A schematic diagram showing the neuroanatomy of the kisspeptin–GnRH pathway and the relationship between KNDy neurons and GnRH neurons is illustrated in Fig. 4.2. KNDy neurons are thought to be a critical component of the so-called GnRH pulse generator. The KNDy hypothesis proposes that NKB is the signal responsible for pulse onset by triggering activation amongst KNDy

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Section 1: Scientific Foundations of Male Infertility

RODENT

HUMAN Arcuate nucleus

AVPV

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PR Kisspeptin neuron

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LH/FSH Kiss1r/KISS1R (Kisspeptin receptor)

Gonads

Gonads Sex steroids

KOR (Kappa opioid receptor)

Sex steroids

Fig. 4.2 Schematic diagram showing the neuroanatomy of the kisspeptin–GnRH pathway and the relationship between KNDy neurons and GnRH neurons in humans and rodents. Kisspeptin signals directly to GnRH neurons which express the kisspeptin receptor. The location of kisspeptin neuron populations within the hypothalamus is species-specific, residing within the anteroventral periventricular (AVPV) nucleus andarcuate nucleus in rodents, and within the preoptic area (POA) and the infundibular nucleus in humans. Kisspeptin neurons in the infundibular (humans)/arcuate (rodents) nucleus coexpress neurokinin B and dynorphin (KNDy) neurons, which, via the neurokinin B receptor and kappa opioid peptide receptor, autosynaptically regulate pulsatile kisspeptin secretion, with neurokinin B being stimulatory and dynorphin being inhibitory. Negative (red) and positive (green) sex steroid feedback is mediated via distinct kisspeptin populations in rodents, via the AVPV nucleus and arcuate nucleus, respectively. In humans, KNDy neurons in the infundibular nucleus relay both negative (red) and positive (green) feedback. The role of the POA kisspeptin population in mediating sex steroid feedback in humans is incompletely explored. +, stimulatory; inhibitory; Dyn, dynorphin; ERα, estrogen receptor alpha; FSH, follicle-stimulating hormone; GnRH, gonadotropin-releasing hormone; Kiss1/KiSS1, kisspeptin; LH, luteinizing hormone; ME, median eminence; NKB, neurokinin B; PR, progesterone receptor. (Reproduced from Skorupskaite K, George JT, Anderson RA. The kisspeptin-GnRH pathway in human reproductive health and disease. Hum Reprod Update 2014;20:485–500. Open access article distributed under the terms of the Creative Commons CC BY license. The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice.)

neurons, whereas kisspeptin serves as the output signal from those neurons driving GnRH secretion and dynorphin acts as the signal terminating each pulse. Indeed, interconnected KNDy cell bodies were found to produce bursts of synchronized firing to coordinate pulsatile GnRH release, and pharmacological manipulation of postsynaptic receptors for kisspeptin, NKB, and dynorphin resulted in altered pulsatile GnRH release [13]. Finally, a number of other neuropeptides (gamma aminobutyric acid (GABA), glutamate, neuropeptide Y (NPY), and many others) act on GnRH neurons by

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binding to their cognate receptors, and contribute to the redundant mechanism that ultimately modulates GnRH secretion.

Endocrine Regulation The role of sex steroids in the endocrine regulation of GnRH secretion has been investigated by several authors. However, questions remain about the relative contributions and the respective sites of feedback of testosterone and estradiol, and the degree to which aromatization to

Chapter 4: The Male Reproductive Endocrine System

E2 is important in mediating the effects of testosterone at the hypothalamic level. A study performed in normal men and men with idiopathic hypogonadotropic hypogonadism (IHH) due to lack of endogenous hypothalamic GnRH secretion produced evidence in favor of a role of both testosterone and estradiol in the endocrine control of GnRH secretion [14]. Medical castration and inhibition of aromatase activity were induced using high-dose ketoconazole (KC), then a subgroup received testosterone replacement and another subgroup received selective add-back of estradiol, while the remaining subjects received no steroid add-back. In normal men, only testosterone replacement restored the GnRH pulse frequency to baseline levels, whereas the effect of estradiol add-back was less and the GnRH pulse frequency, although slowed, remained significantly higher, compared to baseline. Interestingly, the demonstration that E2 lowered LH levels in normal males by slowing pulse frequency, without any effect on pulse amplitude, suggests that the dominant site of E2 feedback may be at the hypothalamus, as suggested by previous studies [15]. Moreover, the demonstration that circulating E2 can modify GnRH secretion in the presence of castrate testosterone levels (the major substrate for central aromatase activity) provides indirect evidence that estrogen effects at the hypothalamus are not dependent on central aromatization. The negative feedback effects of testosterone on GnRH secretion seem to be mediated by kisspeptin neurons located in the ARC. Testosterone inhibits the expression of kisspeptin neurons in the ARC, while it stimulates the expression of kisspeptin mRNA in the anteroventral periventricular nucleus [16]. On the other hand, the site of estrogen action at the hypothalamic level has been a matter of debate, since earlier immunocytochemical studies showed a lack of estrogen receptor (ER) immunoreactivity in GnRH neurons. These results were challenged due to the lack of sensitivity of such technique when dealing with few neurons scattered throughout several nuclei; indeed, when the clonal GT1 neuronal cell line was used, functional ERα and β were detected [17] and later, both fetal and adult GnRH neurons were found to express in vivo both ERα and β mRNA [18]. Similarly, studies evaluating the estrogen effects on GnRH secretion have provided conflicting results. Estrogens seem to exert a negative feedback (but a positive feedback related to the preovulatory GnRH surge in female mice has been also demonstrated) on GnRH neuronal membrane

excitability, cAMP production, and GnRH secretion, mediated by dose-dependent activation of ERs expressed in hypothalamic GnRH neurons [18]. There are also nonclassical rapid effects of estradiol on GnRH neurons, including rapid phosphorylation of cAMP response element binding protein (CREB), potassium–adenosine triphosphate (ATP) channel modulation, increase in intracellular calcium concentrations, and firing rate alteration. However, which of these actions are physiologically relevant remains to be determined. Another site of estrogen action seems to be at the kisspeptin neuron level where estrogens bind to ERα and subsequently inhibit kisspeptin and GnRH release. Indeed, in gonadectomized animals and humans, the decline in sex steroid levels leads to an increase in kisspeptin, GnRH, and gonadotropin levels [19]. The arcuate kisspeptin neurons have been suggested to have roles in the estrogen negative feedback mechanism and GnRH pulse generation. However, the pathways through which these neurons modulate GnRH neurons have not been clarified to date.

Key Points GnRH secretion is modulated at the: • Autocrine level by GnRH itself (ultrashort loop negative feedback) • Paracrine level by kisspeptin/KNDy neurons (the GnRH pulse generator hypothesis) • Endocrine level by testosterone (probably through inhibition of kisspeptin neuron activity) and estrogens

Gonadotropins The gonadotropins follicle-stimulating hormone (FSH) and LH are glycoproteins (oligosaccharide-modified dimeric proteins) with a molecular mass of 30–40 kDa, composed of a common α subunit and a β subunit which confer biologic specificity, associated with noncovalent interactions, with cysteine residues involved in disulfide bonds within subunits. The intrinsic bioactivity of gonadotropins is, to a great extent, determined by their molecular heterogeneity, as determined by variations in carbohydrate content (glycosylation, terminal sialylation, and sulfonation within α and β subunits), which gives rise to gonadotropin isoforms of variable in vivo bioactivity, receptor binding ability, and metabolic clearance.

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A

Cis-Golgi Removal of mannoses

RER Early processing steps

Medial-Golgi Synthesis of hybrid or complex N-glycan precursors

Trans-Golgi Late processing steps: branch elongation SO4

Asn-X-Ser/Thr

Glc3Man3GlcNAc2 P P

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Dolichol GlcNAc

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Glucose

Fucose

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Biantennary

Triantennary SO4

Asn

Asn

Asn

Asn

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Asn

SO4

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Fig. 4.3 Summary of the N-glycan biosynthetic pathway. (A) N-linked glycosylation begins in the rough endoplasmic reticulum (RER) with the cotranslational transfer of a dolichol-linked Glc3Man9GlcNAc2 to a Asn-X-Ser/Thr motif. In the cis-Golgi cisterna, additional mannose residues are removed. In medial-Golgi, hybrid- and complex-type precursors are formed by addition of N-acetylglucosamine (GlcNAc) residues. In trans-Golgi, sequential addition of galactose and sialic acid occurs. Alternatively, sequential addition of N-acetylgalactosamine (GalNAc) and sulfate (SO4) produces sulfated oligosaccharides. (B) Some of the N-linked oligosaccharide structures present on human follicle-stimulating hormone (FSH). High-mannose and hybrid-type N-glycans are incomplete oligosaccharides. Bi- and triantennary oligosaccharides are complex-type N-glycans. A “bisecting” GlcNAc residue attached to β-mannose in the core may be present in complex- and hybrid-type oligosaccharides. Glycoforms lacking terminal residues, such as fucose, galactose, GalNAc, sulfate, and/or sialic acid, may also be present. (Reproduced from Campo S, Andreone L, Ambao V, Urrutia M, Calandra RS, Rulli SB. Hormonal regulation of follicle-stimulating hormone glycosylation in males. Front Endocrinol (Lausanne) 2019;10:17. Open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice.)

Gonadotropin glycosylation is an intricate process involving glycosidase and glycosyltransferases that starts in the rough endoplasmic reticulum where three glucose residues and one mannose residue are removed; then glycoproteins are transferred to the Golgi apparatus where additional mannose residues are removed; hybrid N-glycan precursors are formed by addition of Nacetylglucosamine (GlcNAc) residues, and branch elongation occurs. Sialylated oligosaccharides are formed by addition of galactose and sialic acid. Alternatively, sequential addition of N-acetylgalactosamine (GalNAc) and sulfate produces sulfated oligosaccharides, which account for almost 10 percent of human FSH [20] (Fig. 4.3) An increased number of sialic acid residues increases the half-life of human FSH in the circulation and can reduce receptor affinity at the target organ [21]. On the other hand, studies suggest that hypoglycosylated FSH (hFSH21/18) isoforms are more active than fully glycosylated isoforms in terms of cAMP production, CREB phosphorylation, and protein kinase A (PKA)

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activity, probably due to availability of more binding sites at the receptor level [reviewed in 22]. One polymorphism of the LH β subunit, due to two amino acid transversions (Ile15Thr and Trp8Arg) and a supernumerary consensus glycosylation site (Asn13-Ala-Thr), found in about 30 percent of Northern European and Australian Aboriginal populations, renders gonadotropin more biopotent, but with a shorter in vivo half-life than wildtype LH, due impaired sialylation and sulfation of terminal oligosaccharides [23]. Totally deglycosylated gonadotropins are still able to interact with their cognate receptors but are unable to evoke the generation of second messenger signals [24]. Gonadotropins play their role by interacting with their cognate receptors. The FSH receptor (FSHR) is a G-protein-coupled receptor expressed exclusively on Sertoli cells, although reports suggest that it could be expressed also on spermatogonia [25]. Inactivating mutations of the FSHR are able to affect FSH signaling only when they occur in homozygosis. Five men homozygous for the Finnish p.Ala189Val FSHR gene mutation,

Chapter 4: The Male Reproductive Endocrine System

leading to a marked reduction in ligand binding and signal transduction by the mutated receptor, had small to normal testes volume (4–15 mL), elevated FSH level, and low inhibin B level, and sperm concentrations ranged from