Color Atlas of Human Fetal and Neonatal Histology [2nd ed. 2019] 978-3-030-11424-4, 978-3-030-11425-1

Table of contents :
Front Matter ....Pages i-xvi
Front Matter ....Pages 1-1
Heart (Linda M. Ernst, Michael K. Fritsch)....Pages 3-19
Blood Vessels and Lymphatic Vessels (Linda M. Ernst, Michael K. Fritsch)....Pages 21-32
Front Matter ....Pages 33-33
Lung (Portia A. Kreiger, Jennifer E. Pogoriler)....Pages 35-48
Front Matter ....Pages 49-49
Gastrointestinal Tract (Dale S. Huff, Chrystalle Katte Carreon)....Pages 51-76
Liver (Pierre Russo)....Pages 77-90
Pancreas (Eduardo D. Ruchelli, Tricia R. Bhatti)....Pages 91-97
Salivary Glands (Eduardo D. Ruchelli)....Pages 99-103
Front Matter ....Pages 105-112
Kidney (Eduardo D. Ruchelli, Dale S. Huff)....Pages 113-124
Urinary Bladder (Eduardo D. Ruchelli, Dale S. Huff)....Pages 125-127
Testis (Dale S. Huff, Chrystalle Katte Carreon)....Pages 129-146
Epididymis (Dale S. Huff, Eduardo D. Ruchelli)....Pages 147-155
Vas Deferens (Linda M. Ernst, Eduardo D. Ruchelli, Dale S. Huff)....Pages 157-159
Seminal Vesicle (Linda M. Ernst, Eduardo D. Ruchelli, Dale S. Huff)....Pages 161-165
Prostate Gland (Linda M. Ernst, Eduardo D. Ruchelli, Dale S. Huff)....Pages 167-173
Ovary (Eduardo D. Ruchelli, Dale S. Huff)....Pages 175-181
Fallopian Tubes (Eduardo D. Ruchelli, Dale S. Huff)....Pages 183-186
Uterus (Eduardo D. Ruchelli, Dale S. Huff)....Pages 187-192
Vagina (Eduardo D. Ruchelli, Dale S. Huff)....Pages 193-196
External Genitalia (Michael K. Fritsch)....Pages 197-209
Front Matter ....Pages 211-211
Adrenal Gland (Linda M. Ernst, Lily Marsden)....Pages 213-222
Thyroid Gland (Linda M. Ernst)....Pages 223-231
Parathyroid Gland (Linda M. Ernst, Lily Marsden)....Pages 233-239
Pituitary Gland (Linda M. Ernst)....Pages 241-247
Front Matter ....Pages 249-249
Thymus Gland (Linda M. Ernst, Chrystalle Katte Carreon)....Pages 251-259
Spleen (Linda M. Ernst)....Pages 261-268
Lymph Nodes (Michele E. Paessler)....Pages 269-273
Palatine Tonsil (Linda M. Ernst, Chrystalle Katte Carreon)....Pages 275-279
Bone Marrow (Michele E. Paessler)....Pages 281-288
Front Matter ....Pages 289-289
Brain and Spinal Cord (Patrick Shannon)....Pages 291-310
Eye (Paul J. Bryar, David Gu, Samantha Agron, Sarah E. Eichinger)....Pages 311-323
Ear (Dale S. Huff)....Pages 325-354
Front Matter ....Pages 355-355
Bone (Linda M. Ernst)....Pages 357-366
Skeletal Muscle (Linda M. Ernst, Patrick Shannon)....Pages 367-374
Front Matter ....Pages 375-375
Skin (Maria Cristina Isales, Timothy Tan, Lily Marsden)....Pages 377-384
Mammary Gland (Dale S. Huff)....Pages 385-395
Front Matter ....Pages 397-397
Placenta (Linda M. Ernst, Chrystalle Katte Carreon)....Pages 399-424
Front Matter ....Pages 425-425
Histological Changes Associated with Fetal Maceration (Linda M. Ernst, Michael K. Fritsch)....Pages 427-434
Back Matter ....Pages 435-445

Citation preview

Color Atlas of Human Fetal and Neonatal Histology Second Edition Linda M. Ernst Eduardo D. Ruchelli Chrystalle Katte Carreon Dale S. Huff Editors

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Color Atlas of Human Fetal and Neonatal Histology

Linda M. Ernst  •  Eduardo D. Ruchelli Chrystalle Katte Carreon  •  Dale S. Huff Editors

Color Atlas of Human Fetal and Neonatal Histology Second Edition

Editors Linda M. Ernst Department of Pathology and Laboratory Medicine NorthShore University HealthSystem Evanston, IL USA Chrystalle Katte Carreon Department of Pathology and Laboratory Medicine The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania Philadelphia, PA USA

Eduardo D. Ruchelli Department of Pathology and Laboratory Medicine The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania Philadelphia, PA USA Dale S. Huff Department of Pathology and Laboratory Medicine The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania Philadelphia, PA USA

ISBN 978-3-030-11424-4    ISBN 978-3-030-11425-1 (eBook) https://doi.org/10.1007/978-3-030-11425-1 © Springer Nature Switzerland AG 2019 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

This book is dedicated to all the families who have struggled with fetal loss or the death of a neonate. We hope that by encouraging the careful study of fetal and neonatal tissues, we have contributed to their healing process.

Preface

The writing of this second edition was extremely important for us as authors and teachers because it was our chance to improve upon our first work with insights we did not have the first time around. Firstly, we added another editor to the team with a new, bright, and fresh perspective and the qualities needed to make dry material seem more exciting. Dr. Carreon’s contributions have certainly touched all of the chapters to some extent. In addition, with the assistance of Dr. Lily Marsden, we tried to improve every picture in the text, in terms of brightness, contrast, and sharpness. We hope in the new edition, the figures will be truly reflective of the beauty of developmental histology. This second edition contains six new chapters including blood vessels and lymphatics, external genitalia, eye, ear, skin, and maceration changes. Many existing chapters have also been expanded to address a greater breadth of fetal and neonatal histology such as postnatal testis development and the cardiac conduction system. The “Special Considerations” sections were also expanded in many chapters to address particularly problematic issues within individual organ systems. We hope these new additions will be helpful to readers when faced with more challenging dissections and histologic interpretations. It has been our pleasure to show the breadth and beauty of fetal and neonatal histology while providing a useful reference for pathologists and scientists. Evanston, IL, USA Linda M. Ernst Philadelphia, PA, USA Eduardo D. Ruchelli  Chrystalle Katte Carreon Dale S. Huff

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Preface to the First Edition

The impetus for the writing of this book came largely from the many students and residents in pathology who spent time on the perinatal autopsy service and frequently asked us what reference they could turn to for a review of normal fetal histology. The inspiration for this book is one of the few references that does exist, although out of print, the volume entitled Histology of the Fetus and Newborn by Dr. Valdes-Dapena, published in 1979. This reference served as the model for ours, and we constantly turned to it for verification and help in deciding what to include and what not to include. We hope the quality of our volume is comparable to the standards set by Dr. Valdes-Dapena. The main goal for us was to provide an atlas of fetal and neonatal histology for use by pathology trainees and attending pathologists who have to deal with fetal tissues at autopsy. However, any scientist who might be examining fetal tissues outside of routine pathology might also find this a useful volume. Since there are dramatic changes in many of the fetal tissues over gestation, it is important for the examiner to understand normal development and histology, especially when attempting to decide if a particular finding is pathologic. We have attempted to describe the histology of each organ in detail with particular emphasis on the changes throughout the fetal period. An embryology section is included with each chapter to briefly review the important developmental changes as they apply to the appearance of the tissues. The embryology sections are not intended to be a comprehensive review of the subject since the main goal of our book is to highlight the histology. All the images in this book are obtained from fetal tissues that were formalin fixed and paraffin embedded, unless otherwise stated. Fetal dates were generally obtained by clinical gestational age estimates (menstrual age) and corroborated by fetal morphologic measurements. Since embryologists and embryology texts frequently refer to postfertilization dates, dates prior to 12 weeks are generally postfertilization. The magnifications listed after each image represent the objective power used when taking the photomicrograph. No attempt to calculate exact magnification is made, since most pathologists can relate to objective power more easily. We hope this book will teach others as much as we have learned in the process of writing it, and inspire the next generation of scientists interested in the microscopic anatomy of fetal tissue. Evanston, IL, USA  Philadelphia, PA, USA  Philadelphia, PA, USA 

Linda M. Ernst Eduardo D. Ruchelli Dale S. Huff

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Contents

Part I Cardiovascular System 1 Heart���������������������������������������������������������������������������������������������������������������������������    3 Linda M. Ernst and Michael K. Fritsch 2 Blood Vessels and Lymphatic Vessels ���������������������������������������������������������������������   21 Linda M. Ernst and Michael K. Fritsch Part II Respiratory Tract 3 Lung���������������������������������������������������������������������������������������������������������������������������   35 Portia A. Kreiger and Jennifer E. Pogoriler Part III Digestive System 4 Gastrointestinal Tract�����������������������������������������������������������������������������������������������   51 Dale S. Huff and Chrystalle Katte Carreon 5 Liver���������������������������������������������������������������������������������������������������������������������������   77 Pierre Russo 6 Pancreas���������������������������������������������������������������������������������������������������������������������   91 Eduardo D. Ruchelli and Tricia R. Bhatti 7 Salivary Glands���������������������������������������������������������������������������������������������������������   99 Eduardo D. Ruchelli Part IV Genitourinary Tract 8 Kidney �����������������������������������������������������������������������������������������������������������������������  113 Eduardo D. Ruchelli and Dale S. Huff 9 Urinary Bladder �������������������������������������������������������������������������������������������������������  125 Eduardo D. Ruchelli and Dale S. Huff 10 Testis���������������������������������������������������������������������������������������������������������������������������  129 Dale S. Huff and Chrystalle Katte Carreon 11 Epididymis�����������������������������������������������������������������������������������������������������������������  147 Dale S. Huff and Eduardo D. Ruchelli 12 Vas Deferens �������������������������������������������������������������������������������������������������������������  157 Linda M. Ernst, Eduardo D. Ruchelli, and Dale S. Huff 13 Seminal Vesicle ���������������������������������������������������������������������������������������������������������  161 Linda M. Ernst, Eduardo D. Ruchelli, and Dale S. Huff

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14 Prostate Gland�����������������������������������������������������������������������������������������������������������  167 Linda M. Ernst, Eduardo D. Ruchelli, and Dale S. Huff 15 Ovary�������������������������������������������������������������������������������������������������������������������������  175 Eduardo D. Ruchelli and Dale S. Huff 16 Fallopian Tubes���������������������������������������������������������������������������������������������������������  183 Eduardo D. Ruchelli and Dale S. Huff 17 Uterus�������������������������������������������������������������������������������������������������������������������������  187 Eduardo D. Ruchelli and Dale S. Huff 18 Vagina�������������������������������������������������������������������������������������������������������������������������  193 Eduardo D. Ruchelli and Dale S. Huff 19 External Genitalia�����������������������������������������������������������������������������������������������������  197 Michael K. Fritsch Part V Endocrine System 20 Adrenal Gland�����������������������������������������������������������������������������������������������������������  213 Linda M. Ernst and Lily Marsden 21 Thyroid Gland�����������������������������������������������������������������������������������������������������������  223 Linda M. Ernst 22 Parathyroid Gland ���������������������������������������������������������������������������������������������������  233 Linda M. Ernst and Lily Marsden 23 Pituitary Gland���������������������������������������������������������������������������������������������������������  241 Linda M. Ernst Part VI Hematolymphoid System 24 Thymus Gland�����������������������������������������������������������������������������������������������������������  251 Linda M. Ernst and Chrystalle Katte Carreon 25 Spleen�������������������������������������������������������������������������������������������������������������������������  261 Linda M. Ernst 26 Lymph Nodes�������������������������������������������������������������������������������������������������������������  269 Michele E. Paessler 27 Palatine Tonsil�����������������������������������������������������������������������������������������������������������  275 Linda M. Ernst and Chrystalle Katte Carreon 28 Bone Marrow�������������������������������������������������������������������������������������������������������������  281 Michele E. Paessler Part VII Central Nervous System and Special Sense Organs 29 Brain and Spinal Cord���������������������������������������������������������������������������������������������  291 Patrick Shannon 30 Eye �����������������������������������������������������������������������������������������������������������������������������  311 Paul J. Bryar, David Gu, Samantha Agron, and Sarah E. Eichinger 31 Ear �����������������������������������������������������������������������������������������������������������������������������  325 Dale S. Huff

Contents

Contents

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Part VIII Musculoskeletal and Connective Tissues 32 Bone���������������������������������������������������������������������������������������������������������������������������  357 Linda M. Ernst 33 Skeletal Muscle���������������������������������������������������������������������������������������������������������  367 Linda M. Ernst and Patrick Shannon Part IX Integument 34 Skin�����������������������������������������������������������������������������������������������������������������������������  377 Maria Cristina Isales, Timothy Tan, and Lily Marsden 35 Mammary Gland�������������������������������������������������������������������������������������������������������  385 Dale S. Huff Part X Placenta 36 Placenta���������������������������������������������������������������������������������������������������������������������  399 Linda M. Ernst and Chrystalle Katte Carreon Part XI Special Considerations for Fetal Tissues 37 Histological Changes Associated with Fetal Maceration���������������������������������������  427 Linda M. Ernst and Michael K. Fritsch Index������������������������������������������������������������������������������������������������������������������������������������� 435

Contributors

Samantha  Agron, BA Department of Ophthalmology, Northwestern University Feinberg School of Medicine, Chicago, IL, USA Tricia  R.  Bhatti, MD  Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA Paul J. Bryar, MD  Departments of Ophthalmology and Pathology, Northwestern University Feinberg School of Medicine, Chicago, IL, USA Chrystalle  Katte  Carreon, MD  Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA Sarah  Eichinger, BA Department of Ophthalmology, Northwestern University Feinberg School of Medicine, Chicago, IL, USA Linda M. Ernst, MD, MHS  Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA Michael K. Fritsch, MD, PhD  Department of Pathology and Laboratory Medicine, University of Wisconsin, Madison, Madison, WI, USA David Gu, BA  Department of Ophthalmology, Northwestern University Feinberg School of Medicine, Chicago, IL, USA Dale  S.  Huff, MD Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA Maria  Cristina  Isales, MD, MPH Department of Pathology, Northwestern Memorial Hospital, Northwestern University Feinberg School of Medicine, Chicago, IL, USA Portia A. Kreiger, MD  Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA Lily Marsden, MD  Utah State Office of the Medical Examiner, Salt Lake City, UT, USA Michele E. Paessler, DO  Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA Jennifer E. Pogoriler, MD, PhD  Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA

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Eduardo  D.  Ruchelli, MD Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA Pierre  Russo, MD Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA Patrick  Shannon, Msc, MD, FRCC  Department of Pathology and Laboratory Medicine, University of Toronto, Mount Sinai Hospital, Toronto, ON, Canada Timothy Tan, DO  Department of Pathology, Northwestern Memorial Hospital, Northwestern University Feinberg School of Medicine, Chicago, IL, USA

Contributors

Part I Cardiovascular System

1

Heart Linda M. Ernst and Michael K. Fritsch

Introduction The fetal myocardium has a distinctive histological appearance that differs from the histological appearance of adult myocardium. Familiarity with these differences is important when examining the fetal heart. In addition, there are specialized structures, such as the fibrous skeleton, cardiac valves, and cardiac conduction system, which show important changes during fetal and neonatal life.

Embryology Formation of the human heart is a complex developmental process that is reviewed more extensively in embryology texts [1, 2] and recent reviews [3–5]. It is reviewed briefly in this chapter for background purposes and as needed to assist in the understanding of cardiac anatomy and specialized structures of the heart.

General Cardiac Formation The heart begins to form from bilateral angiogenic clusters located in the splanchnic mesoderm of the embryo. These clusters form into the bilateral endocardial tubes that fuse with the folding of the embryo around 21–22 days postconception. The developing heart tube is suspended in the pericardial cavity by a dorsal mesocardium and surrounded by two mesodermal layers, the cardiac jelly and the epimyocardial mantle. L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected] M. K. Fritsch Department of Pathology and Laboratory Medicine, University of Wisconsin, Madison, Madison, WI, USA e-mail: [email protected]

Specialized cells from various sources, in addition to first heart field cells, including second heart field cells, neural crest cells, and proepicardial cells, some of which invade the cardiac jelly, contribute in the eventual formation of the three principal layers of the myocardial wall: endocardium, myocardium, and epicardium. The cardiac tube is initially a straight tube with its cranial end attached to the branchial arches. The intrapericardial portion consists of the bulbus cordis and ventricular tissues, and the caudal end, or atrial portion, is embedded in the septum transversum. As the heart tube elongates, it also bends, a process known as formation of the cardiac loop, which begins on day 23. The cephalic portion of the heart tube moves ventrally, caudally, and to the right, while the caudal portion moves dorsally, cranially, and to the left. During this process of looping, which is completed around day 28, the cranial portion of the heart tube gives rise to the truncus arteriosus, the conus cordis, and the trabeculated part of the right ventricle (derived from the bulbus cordis). The ventricular portion of the heart tube gives rise to the primitive left ventricle, and the atrial portion of the heart tube becomes incorporated into the pericardium as it takes its dorsal position. Partitioning of the chambers begins around day 27 postconception. The atrial septum forms from two distinct membranes: the septum primum and septum secundum. Septum primum appears first and grows downward toward the endocardial cushions but does not reach the endocardial cushions, leaving an opening known as the ostium primum, which is later closed by growth of the endocardial cushions. As the ostium primum is obliterated by the continued expansion of the septum primum and endocardial cushions, fenestrations begin to appear in the upper portion of the septum primum. These openings coalesce and form an opening known as the ostium secundum. Subsequently, the septum secundum, the second interatrial membrane, begins to form to the right of the septum primum. It is an incomplete septum, with the upper solid portion overlapping the ostium secundum and a more inferior ovoid defect known as the

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_1

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foramen ovale. Therefore, shunting blood from the right atrium occurs across the interatrial septum through the foramen ovale, over the free edge of the septum primum, and into the left atrium. The common atrioventricular (AV) canal, which encompasses the lower atrial and upper ventricular portions in the center of the heart, is closed by formation of the lateral, superior, and inferior endocardial cushions, which appear as bulges of myocardium. The endocardial cushions are formed by a subset of endocardial cells and neural crest cells which invade the cardiac jelly at critical inflow and outflow points. They fuse to form two separate left and right AV canals. Most of the ventricular septum is formed, while the two primitive ventricles are enlarging. The medial walls of the two enlarging ventricles fuse and form a nearly complete interventricular septum with a small opening just inferior to the endocardial cushions. This opening persists until conal septal formation is complete and the upper portion of the ventricular septum, also known as the membranous septum, is closed by the union of portions of the right and left conal swellings and the inferior AV cushion. The outflow of the ventricles is created by the formation of the conal swellings within the conus cordis and the truncal swellings within the truncus arteriosus. In about the fifth week postconception, the truncal swellings appear as ridges that form from the right superior and left inferior truncal walls. These ridges grow toward each other and twist around one another, creating the spiral aorticopulmonary septum. Within the more proximal cordis conus, similar bilateral swellings appear and grow upward, meeting the aorticopulmonary septum. The conal swellings also fuse with the inferior AV cushion to close the upper portion of the interventricular septum.

Cardiac Valve Formation All four cardiac valves will develop from the mesenchymal cells of the endocardial cushions. Valve development is closely related to cardiac septation and the development of the AV canal and outflow tracts. As mentioned above, in early heart tube development, the myocardial layer is separated from the endocardium by cardiac jelly, a mixture of acellular extracellular matrix material consisting mainly of collagens (I, III, IV), glycoproteins, mucopolysaccharides, and glycosaminoglycans. In the early heart tube, prior to looping, unidirectional blood flow is achieved because the entire lumen collapses during contraction due to the presence of cardiac jelly in the wall, thereby preventing reversal of blood flow during relaxation of the heart muscle. As looping commences, the cardiac jelly disappears in most of the heart except at the junctions of the AV canal and the

L. M. Ernst and M. K. Fritsch

outflow tract, where the endocardial cushions develop. The endocardial cushions contribute to both cardiac septation and valve formation. The AV valve leaflets are predominantly derived from endocardial cells, whereas the semilunar valves of the outflow tract are derived from endocardial and neural crest cells, both of which locally invade the cardiac jelly and undergo an epithelial-to-mesenchymal transition. In order to maintain unidirectional blood flow as the heart enlarges, cardiac valves must develop. The development of the AV valves (tricuspid and mitral valves) is complex; they develop independently of each other. In brief, beginning in the fifth week of development, the mesenchymal tissue at the corresponding endocardial cushions of the AV canal will begin remodeling, and eventually, by about the eighth week, it will detach from the underlying myocardium (delaminate). This complex process of forming each leaflet involves the differentiation of mesenchymal cells into fibroblasts, laying down of collagen, cell proliferation, apoptosis, and local blood flow dynamics. The chordae tendineae are predominantly derived from the mesenchyme of the AV cushions; over time, they become fibrotic. The papillary muscles are formed from the myocardial cells via compaction of the trabecular layer. There are subtle differences in the development of each leaflet and each AV valve [3, 4]. The cusps of the semilunar valves at the outflow tract are derived from the four corresponding outflow endocardial cushions (ridges), which include the truncal and conal cushions involved in septation as well as two smaller intercalated cushions. Therefore, the formation of the semilunar valves is intimately tied to the formation of the conal septum. Rather than delamination, the sinuses of Valsalva form via apoptosis, with resulting excavation of these spaces. Again each cusp forms uniquely in a complex interplay between apoptosis, cell proliferation, and local flow dynamics to generate the final valve morphology.

Cardiac Conduction System Formation The different components of the cardiac conduction system develop independently, and each possesses unique gene and protein expression programs during development [3–8]. The initial cardiac pacemaker is found in the caudal part of the left heart tube. As heart development proceeds, cells in the sinus venosus take over as the pacemaker. The sinus venosus will form part of the right atrium and will give rise to the eventual cardiac pacemaker, the sinoatrial (SA) node, located along the ridge of the right atrium near the superior vena cava. The SA node electrical depolarization signals are conducted rapidly to the AV node via anterior, middle, and posterior internodal tracts in the right atrium [3–8].

1 Heart

The AV node forms at the base of the interatrial septum in the right atrium, just anterior to the opening of the coronary sinus. Its primary function is to slow the conduction signal from the SA node so that the atria can complete contraction before ventricular contraction. The AV node develops in situ from myocytes. The bundle of His development remains uncertain, but it may be similar to the development of the AV node. One model proposes that a primary ring of conduction tissue develops at the level of the AV border in the embryonic heart, prior to the development of the fibrous annulus. As the heart completes its anatomic development, the ring is pulled up, and parts of this ring degenerate, leaving the AV node and bundle of His. However, the bundle of His will develop a different cellular gene expression profile, which allows for the rapid conduction of the electrical signal from the AV node to the bundle branches [3–10]. Initially in the primordial heart, there is no fibrous annulus, and atrial and ventricular myocytes are continuous. By approximately 12 weeks of gestation, the fibrous annulus is mostly complete, thereby separating atrial and ventricular myocytes everywhere except for the penetrating bundle of His [3–8, 10]. The bundle of His divides into bundle branches and then more distally toward the apex of the heart into Purkinje fibers, both localized in the ventricular walls. Both the bundle branches and Purkinje fibers are derived directly from subendocardial myocytes. The protein expression profiles in the bundle branches and Purkinje cells change with developmental age and are unique from the SA node, AV node, and bundle of His. The bundle branch and Purkinje cell differentiation begins and is completed later in development

a

Fig. 1.1  Comparison of fetal and adult myocardium. (a) Fetal myocardium at 23 weeks of gestation. (b) Adult myocardium. Note the marked hypercellularity and higher nuclear to cytoplasmic ratio of the fetal

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than the SA and AV nodes [3–8, 10]. The cells in the bundle branches and Purkinje fibers conduct the depolarization signal rapidly, as these portions of the conduction system need to coordinate rapid ventricular contraction from the apex toward the base.

Histology The low-power appearance of fetal and neonatal myocardium is quite different from the appearance of adult myocardium. The striking difference is the cellularity of the myocardium (Fig. 1.1). The fetal myocardial cells are generally smaller than adult myocardial cells and have higher nuclear to cytoplasmic ratios. The traditional view has been that the number of cardiac myocytes is established at birth and there is little regenerative capacity among the population of terminally differentiated myocardial cells. This paradigm has been challenged recently, however, with the discovery of a cardiac stem cell population with regenerative capacity [11]. Nevertheless, the highly cellular appearance of the fetal myocardium is not maintained throughout life. Despite these morphologic cellular differences, the basic histological structure of the fetal heart is not different from the adult heart. The majority of the heart consists of three principal layers: epicardium, myocardium, and endocardium. Additionally, there are specialized tissues such as the fibrous skeleton, valves, and conduction system, which can be examined if they are specifically sampled for histological study.

b

myocardium compared with the adult myocardium (hematoxylin and eosin [H&E], 20×)

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Fig. 1.2  Fetal pericardium at 23 weeks of gestation. The fetal pericardium is a delicate, membranous structure with a simple squamous mesothelium and a layer of underlying dense connective tissue. In the adult, the loose connective tissue outside the pericardium can contain adipose tissue, but there is usually little adipose tissue in the fetus (H&E, 20×)

Pericardium The heart is surrounded by a fibrous sac, the pericardium (parietal pericardium), which grossly is a thin membrane in the fetus. Anatomically, the parietal pericardium is in continuity with the serous (visceral) pericardium, also known as the epicardium, at the pericardial reflections over the great vessels [12]. The fibrous pericardial sac may normally contain a small amount of serous fluid in the fetal and neonatal period, and in macerated fetuses, it may contain abundant serosanguineous fluid. Histologically, the pericardial sac consists of a core of dense connective tissue and is lined on the inner surface by a flattened layer of mesothelial cells (Fig.  1.2). In fetuses and neonates, the outer layer of the pericardium consists of loose connective tissue with occasional blood vessels and without significant adipose tissue. Heterotopic thymic or thyroid tissue can be seen in the pericardial sac [12].

Epicardium The external surface of the heart, known as epicardium, or alternatively the visceral pericardium, is composed of a simple squamous layer of mesothelial cells and variable amounts of underlying connective tissues (Fig. 1.3). Early in fetal life, the amount of connective tissue is quite sparse, but it becomes thicker with increasing gestational age (Fig. 1.4). The epicardial connective tissues consist mainly of fibrous connective

L. M. Ernst and M. K. Fritsch

Fig. 1.3  Fetal epicardium at 34 weeks of gestation. The epicardium is composed of a connective tissue layer with a flattened mesothelial layer (arrow). Note the fairly sharp demarcation between the underlying myocardium and the connective tissue of the epicardium (H&E, 20×)

tissue that contains epicardial coronary vessels. Nerves are also seen within the connective tissue. Fat is not a prominent feature of the fetal epicardium, but some adipose tissue is seen near term.

Myocardium The middle, thickest histological layer of the atrial and ventricular wall is the myocardium, which is composed of cardiac-­type muscle fibers with striated cytoplasm, centrally placed nuclei, and intercalated discs. Sections of the fetal myocardium show both longitudinal fibers and cross sections of the cardiac myocytes, arranged loosely in bundles. Early fetal myocytes frequently appear to have clear, vacuolated cytoplasm secondary to high glycogen content. This change appears most prominently in the midtrimester to early third trimester (Fig. 1.5). As the fetus approaches term, the clearing of the fetal myocyte cytoplasm is generally less prominent (Fig. 1.6). The myocytes acquire more cytoplasm as term approaches, and the cytoplasm becomes more eosinophilic. Myocardial cell nuclei are generally ovoid to slightly elongated with smooth nuclear borders. Cells with features of Anitschkow cells, also known as Anitschkow nuclear structure, have been described to occur commonly in fetal and neonatal hearts [13]. Anitschkow cells are characterized by a caterpillar-shaped nucleus in longitudinal sections, and in cross sections the nucleus shows a central condensation of chromatin with fine fibrillar chromatin extensions to the nuclear membrane. This nuclear

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a

b

c

d

Fig. 1.4  Epicardium of the heart during gestation. The epicardium of the heart is shown at gestational ages (a) 14 weeks, (b) 24 weeks, (c) 34 weeks, and (d) 40 weeks. Note the increasing thickness of the epi-

cardial connective tissue with increasing gestational age. Epicardial coronary arteries are present within the epicardium (H&E, 10×)

appearance has been classically associated with rheumatic heart disease [14, 15] (Fig. 1.7). Though Anitschkow cells frequently represent myocytes, the Anitschkow nuclear structure has been described in other cells within the heart, such as endothelium, histiocytes, and stromal cells. The significance of this particular nuclear structure is not clear, but it may represent a reactive change to stimuli such as hypoxia [13]. Physiologically, cardiac myocytes possess three major characteristics: (1) they possess sarcomeres, which mediate contraction; (2) they can spontaneously depolarize, which is regulated by membrane ion pumps and channels; and (3) they are electrically coupled to adjacent myocytes via gap junctions. The atrial and ventricular myocytes are ­fast-­conducting cells with well-developed sarcomeres that allow for strong contraction, their major function.

All atrial and ventricular contractile cardiac cells express Nkx2.5 protein as part of the standard myocardial profile. Cardiac myocytes are connected to each other via the intercalated disc composed of multiple components including gap junctions. Connexins, a family of proteins that help make up the gap junction, play a critical role in the speed of conduction of electrical coupling between cardiac myocytes [9]. Connexins allow for propagation of the electrical signal that results in sequential and coordinated contraction. There are three major connexins found in the human heart; two are fast-conducting connexins, Cx40 and Cx43, and one is a slow-conducting connexin, Cx45. Both atrial and ventricular myocardia express fast-conduction connexins. The embryonic AV canal (AV junction) cells express slow-conducting connexins, which slows conduction to allow the atria to contract before the ventricles.

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tive tissue layer in the ventricle (Fig. 1.8). Atria are also lined by endocardium, but normally the underlying connective tissue, consisting of collagen and elastic fibers, is thicker than in the ventricle (Fig. 1.9).

Specialized Structures of the Heart Papillary Muscles and Chordae Tendineae

b

Papillary muscles are specialized cones of ventricular myocardium into which the chordae tendineae of the AV valves insert. The base of the papillary muscle has an endocardial lining on all sides, surrounding the central myocardial tissue. At the tip of the papillary muscle, where the chordae tendineae insert, the myocardium is gradually replaced by fibrous tissue that merges with the fibrous tissue of the chordae tendineae (Fig. 1.10). The very tip of the papillary muscle may also lack its endocardial lining. The chordae tendineae, which tether the AV valve leaflets to the papillary muscle, have a central core of densely arranged collagen fibers with peripheral, more loosely arranged, collagen fibers.

Valve Leaflets and Cusps

c

Fig. 1.5  Myocardium in the second trimester. Myocardium is shown at gestational ages 14 weeks (a) 19 weeks (b) and 24 weeks (c) (H&E, 40×)

Endocardium The innermost layer of the heart is the endocardium, which appears as a simple squamous layer of flattened endothelial cells supported by a generally very thin, underlying connec-

Fetal valve leaflets and cusps are delicate structures composed predominantly of a fairly densely arranged central core of collagen fibers, with the free surfaces lined by endocardium (Figs.  1.11, 1.12, and 1.13). By approximately 12  weeks of development, the AV and outflow tract valve structures are complete, but they continue to undergo maturation during fetal life. Maturation involves remodeling of the extracellular matrix of the valves to form the final three layers, which are covered by endothelial cells. The atrialis layer of the AV valve leaflets and the ventricularis layer of the outflow tract cusps are the connective tissue layer closest to blood flow and are composed of extracellular matrix material and elastic fibers in order to provide motility to the valves. The second and central layer is referred to as the spongiosa, which is composed of proteoglycans and allows for compressibility of the valve leaflets. The third layer, referred to as the fibrosa, lies on the opposite side away from direct blood flow (closest to the ventricular myocardium or great vessel wall) and is mostly composed of collagen, to provide stiffness to the valves. Distinction of these three layers is difficult to discern by H&E-stained slides, especially in fetal hearts. Fetal valve stroma is frequently focally myxoid in character (Fig. 1.14), but nodular myxoid structures are considered a sign of valvular dysplasia (Fig.  1.15). Although cardiac valve tissue is not vascularized except at the base, a

1 Heart Fig. 1.6  Myocardium in the third trimester. Myocardium is shown at gestational ages (a) 34 weeks, and (b) 40 weeks (H&E, 40×)

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Cardiac Conduction System

Fig. 1.7  Anitschkow cell. This is a high-power view of the myocardium in a case of rheumatic heart disease. This perivascular cell is likely a histiocyte or stromal cell. Note the chromatin clumping in the center of the nucleus with extensions to the nuclear membrane, which give it a caterpillar appearance (H&E, 100×)

common finding in the fetal heart valve, especially the mitral and tricuspid valves, is a structure known as a congenital blood cyst. These are small, red-brown excrescences near the free edge of the leaflets on the atrial side (Fig. 1.16). Histologically, they consist of ovoid cystic spaces containing blood and are lined by endocardial-like cells (Fig. 1.17). Evidence from serial sections has shown that blood cysts result from blood being pressed into the crevices of the valve tissue and retained after the mouth of the crevice fuses shut [16, 17].

The conduction system of the heart, which consists of specialized myocardial cells, is involved in propagating the electrical signal throughout the heart that results in coordinated ventricular contraction. The origin of the electrical activation occurs within the pacemaker cells of the SA node, which is located within the right atrium at the junction with the superior vena cava, near the crista terminalis. The signal is then conducted via poorly understood atrial internodal tracts to the AV node located at the interatrial septum. The AV node has fewer cell-cell junctions and slows the electrical signal, which is then conveyed at a more rapid rate via the bundle of His (also referred to as the AV bundle). The bundle of His passes through the fibrous annulus, which separates the atrial myocytes from the ventricular myocytes. In the normal heart, the bundle of His is the only electrical connection between the atria and the ventricles. The signal then diverges into the right and left bundle branches and is rapidly conducted to the distal Purkinje fibers near the apex of the heart. The signal then spreads rapidly in the ventricular myocardium from apex to base, leading to ventricular contraction in the same sequence. The nodal cells, which make up the SA and AV nodes of the cardiac conduction system, have poorly developed sarcomeres and are poorly electrically coupled to adjacent cells. These features of nodal cells allow them to build up an electrical charge, and they have a high frequency of automaticity (pacemaker activity). Because of the poor coupling and a different protein expression profile, including Cx45, nodal myocytes are slow conductors [6, 7, 9]. The bundle of His,

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Fig. 1.8 Endocardium of the left ventricle during gestation. Endocardium of the left ventricle is shown at gestational ages (a) 14  weeks, (b) 24  weeks, (c) 34  weeks, and (d) 40  weeks. Note the

Fig. 1.9  Right atrium at 23 weeks of gestation. The full thickness of the right atrium at 23 weeks of gestation, including a slightly thicker endocardium and thinner myocardial layer, is shown (H&E, 4×)

minimal increase in thickness of the endocardial connective tissue with increasing gestational age (H&E, a 20×; b–d 10×)

bundle branches, and Purkinje fibers are primarily fast-­ conducting systems expressing Cx40 and Cx43 [6, 7, 9]. Anatomically, the SA node lies just under the epicardium at the junction of the superior vena cava and the right atrium. Histologically, the myocytes composing the SA node are paler, smaller, and more compact than adjacent cardiac myocytes. The SA node cells are embedded in a fibrous stroma with a large central artery. Use of trichrome stain accentuates the fibrous stroma and reveals that the cells of the SA node are paler than the surrounding atrial myocytes (Fig. 1.18). Large nerves and ganglion cells are often seen adjacent to the SA node [18]. Several tracts from the SA node radiate through the atrium toward the AV node/bundle of His complex. The AV node lies in the right atrial wall within the triangle of Koch, a useful anatomic landmark formed by the coronary sinus orifice (base), the anterior portion of the tricuspid valve annulus (anteriorly), and the tendon of Todaro, the tendinous structure connecting the valve of the inferior vena cava ostium to

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Fig. 1.10  Papillary muscle at 40 weeks of gestation. The tip of a papillary muscle at 40 weeks of gestation, including the transition to chordae tendineae, is shown (H&E, 4×)

Fig. 1.11 Atrioventricular (AV) valve at 23 weeks of gestation. A low-power view of the valve leaflets of the AV valve at 23 weeks of gestation is shown (a H&E, 4×; b Masson trichrome, 4×)

a

the central fibrous body (posteriorly). Histologically, the AV node is similar to the SA node, including smaller myocytes than adjacent cardiac myocytes, which are more compact and are embedded in a fibrous stroma with a large central artery. The AV node usually lies directly on the atrial side of the fibrous annulus, and the bundle of His extends directly from the AV node toward the central fibrous body (Fig. 1.19). The bundle of His runs from the AV node just superior to the endocardial cushions and eventually passes through the fibrous annulus near the central fibrous body, thereby representing the only conduction pathway through the fibrous annulus. Without a complete fibrous annulus, atrial and ventricular myocardial cells would be in direct physical contact with each other, and this could lead to reentry arrhythmias. Initially, in the primordial heart, there is no fibrous annulus; atrial and ventricular myocytes are continuous. By approximately 12 weeks of gestation, the fibrous annulus is mostly complete, thereby separating the atrial and ventricular myocytes everywhere except for the penetrating bundle of His [3–9] (Fig. 1.20). The bundle of His can often be identified microscopically penetrating the fibrous annulus and extending toward the interventricular septum (Fig. 1.21). Use of Masson trichrome stain accentuates the fibrous stroma and reveals that the cells of the AV node and bundle of His are also paler than the surrounding atrial and ventricular myocytes. This slight difference in color remains dependent on the quality of the Masson trichrome stain as well as on the presence of autolysis in fetal hearts from stillbirths with prolonged intrauterine fetal

b

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Fig. 1.12  AV valve at 23 weeks of gestation. A higher-power view of an AV valve leaflet at 23  weeks of gestation is shown, including the fibrous core and the simple squamous endothelial lining on both sides of the valve leaflet (Masson trichrome, 10×)

Fig. 1.13  AV valve at 40 weeks of gestation. A low-power view of an AV valve leaflet at 40 weeks of gestation is shown. This valve vaguely shows the three connective tissue layers (atrialis, spongiosa, and fibrosa) (H&E, 4×)

demise. As with the SA node, large nerves can occasionally be seen near the AV node [18]. The SA and AV nodes and bundle of His can be difficult to identify in the midtrimester and even in term fetal hearts because of the small size of the hearts, the small size of the elements of the conduction system, oblique or tangential sectioning, and the subtlety of the distinctive histologic features. The fibrous meshwork in which the cells of the nodes are embedded is not fully developed until postnatal life. The nodes and bundle of His are not as “tightly” formed as they will be in later postnatal life (referred to as dispersion in older literature) [19] (Fig. 1.22). In the perinatal period, the tissues of the conduction system contain loosely woven connective tissue without any evidence of active degeneration, necrosis, phagocytosis, inflammation, or replacement fibrosis [20, 21]. The bundle of His branches to form the right and left bundle branches, which travel downward along the endocardial surfaces of the interventricular septum, seen sepa-

rated from the myocardium by a thin fibrous band that is best identified by Masson trichrome stain (Fig. 1.23). The bundle branches and Purkinje fibers rapidly transmit the signal to the ventricular parenchyma that will allow for ventricular contraction occurring from the apex toward the base of the heart. The methods of removal of the cardiac conduction system have been described in many articles and texts, including a recent review on the pathologic findings in the cardiac conduction system in infants with sudden death [22]. The following is a brief description of how to dissect and examine the cardiac conduction system histologically: Begin with dissection of the SA node, which is situated in the superior part of the right atrium, where the superior vena cava enters the right atrium. To remove it, first identify the ridge of atrial tissue that runs from the tip of the right atrial appendage to the superior vena cava. Then excise a rectangular piece of right atrial tissue defined by the superior vena cava, about 1 cm of tissue on each side of the sulcus termi-

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Fig. 1.15  AV valve at 19 weeks of gestation. A tricuspid valve leaflet at 19 weeks of gestation with valvular dysplasia is shown. Note the well-defined nodular myxoid excrescence on the valve leaflet (H&E, 4×)

Fig. 1.14  AV valve at 19 weeks of gestation. This low-power view of an AV valve leaflet at 19 weeks of gestation shows the focally myxoid character of the fibrous stroma and flattened endothelial lining on both sides (H&E, 10×)

Fig. 1.16  Congenital blood cyst on the septal leaflet of the tricuspid valve, shown in a gross photograph. These cysts are typically ovoid, red-brown excrescences on the atrial side of the valve leaflet. Some can be large and bulbous, as shown here

nalis (atrial ridge), and about 2  cm distal to the superior vena cava. Keep the piece of tissue oriented with the atrial ridge in the middle, and cut four to six strips of tissue parallel to the ridge (about 3–4 mm thick) and embed them on edge (Fig. 1.24). The AV node and bundle of His are removed together en bloc and are identified by viewing the heart from the right side after opening the right atrium and ventricle. The AV node is situated in the posterior right atrial wall just superior to the septal leaflet of the tricuspid valve, below the tendon of Todaro, and adjacent to the coronary sinus ostium (within the triangle of Koch). Three anatomic landmarks should be identified when attempting to remove the AV node and bundle of His: The foramen ovale marks the superior border; the central fibrous body (identified by holding the specimen up to the light and finding the thinnest part of the superior septum) marks the medial border; and the ostium of the coronary sinus defines the lateral border. To remove these structures, a rectangular portion of tissue is excised by making four incisions: (1) a transverse incision just below the foramen ovale; (2) a medial vertical incision through the central fibrous body from the atrium down below the septal leaflet of the tricuspid valve, taking more ventricular tissue than atrial tissue; (3) a lateral vertical incision immediately adjacent to the coronary sinus; and (4) a final transverse incision through the ventricular septum (Fig. 1.25). This block of tissue containing the upper interventricular septum and lower interatrial septum is removed and sectioned longitudinally from lateral to medial at intervals of 2–3 mm; each section is embedded on edge (Fig. 1.26). It should be noted that removal of the cardiac

14 Fig. 1.17  Blood cyst on the mitral valve. Note the blood-filled spaces lined by flattened endocardial-like cells (H&E, a 2×; b 10×)

L. M. Ernst and M. K. Fritsch

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Fig. 1.18  The sinoatrial (SA) node, shown at (a) 23  weeks, (b) 27 weeks, (c) 40 weeks, and (d) in an infant at 7 months of life (d). The SA node (asterisk) lies just below the epicardium and is made up of

pale nodal cells in a fibrous meshwork surrounding a central artery. The SA node is better defined with increasing developmental age (Masson trichrome, 4×)

1 Heart

conduction system will completely disrupt the internal anatomy of the heart; in many types of congenital heart disease, the AV node and bundle of His can be displaced to alternative sites.

Special Considerations Extramedullary Hematopoiesis in the Heart Extramedullary hematopoiesis (EMH) in the heart is not commonly encountered in the fetus, but it has been described in both fetal and adult hearts. EMH in the heart typically appears as clusters of immature erythroid or myeloid cells within myocardial capillaries or interstitium. In one study that included examination of EMH in the hearts of fetuses and children, the incidence of myocardial EMH was highest among stillborn fetuses (13%) and infants less than 1  year

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old (8%) [23]. Their data showed that the incidence of EMH in the heart gradually decreased with age after the first year of life [23]. In the same study, in adult myocardium, EMH was seen in association with myocardial infarction, but not in normal or nonischemic myocardium [23]. EMH appears to reflect inflammatory or repair pathways in adult myocardium, but its etiology in fetal and neonatal hearts remains unclear.

Vacuolation of Myocytes As stated above, fetal myocytes frequently appear to have clear, vacuolated cytoplasm secondary to high glycogen content (Figs. 1.5 and 1.6). If more extensive vacuolar change is seen, however, it is likely to represent pathologic change. Vacuolar degeneration, sometimes referred to as myocytolysis or colliquative myocytolysis, has been described, particu-

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Fig. 1.19  The AV node is shown during gestation, using Masson trichrome stain. (a) 14 weeks (4×); (b) 23 weeks (4×); (c) 40 weeks (2×); (d) infant at 7 months of life (2×). The AV node (asterisk) lies on the atrial side of the fibrous annulus with a central artery and pale nodal

cells embedded in a poorly defined fibrous meshwork. The AV node is better defined with increasing developmental age. At atrial myocardium, V ventricular myocardium

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Fig. 1.20  The fibrous annulus in early and late gestation. (a) The fibrous annulus (asterisk) separating the atrium (At) and ventricle (V) at 12  weeks of gestation is irregular, thin, and nearly complete. (b) At

L. M. Ernst and M. K. Fritsch

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37 weeks of gestation, the fibrous annulus (asterisk) is better defined and completely separates the atrium (At) from the ventricle (V) (Masson trichrome, a 2×; b 4×)

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Fig. 1.21  The bundle of His is shown during gestation. (a) 14 weeks (4×). (b) 27 weeks (2×). (c) 37 weeks (2×). (d) Infant at 7 months of life (2×). The bundle of His (asterisk) initially lies on the atrial side of the

fibrous annulus and passes through it and is composed of pale cells with virtually no fibrotic meshwork. At atrial myocardium, V ventricular myocardium (Masson trichrome)

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Fig. 1.22  The bundle of His showing dispersion. (a) At 12 weeks, the bundle (asterisk) is poorly formed (4×). (b) In an infant at 7 months of life, the bundle of His (asterisk) splits into multiple components as it passes into the fibrous annulus (2×). At atrial myocardium, V ventricular myocardium (Masson trichrome)

larly in the subendocardial location; it may be a form of myocyte injury that occurs without a prominent inflammatory or granulation tissue response [24, 25]. The myocyte cytoplasm appears clear, swollen, and empty, with loss of myofibrils. The empty spaces in the cells are thought to represent water, glycogen, and/or lipids [24]. Care should be taken to exclude pathologic conditions such as ischemia, cardiomyopathy, myocarditis, or a storage disorder when extensive vacuolar change is seen.

Lymphocytes in the Heart In macerated myocardial tissues, lymphocytes can be nearly impossible to recognize, and in our experience, cells of the immune system are not a prominent feature on H&E histology of the normal fetal myocardium. However, studies have shown that T cells, B cells, and histiocytes can be seen in normal fetal heart tissue to some degree when immunohisto-

Fig. 1.23 The bundle branches are shown during gestation. (a) 27 weeks. (b) Infant at 7 months of life. The bundle branches (asterisk) lie on the ventricular side of the fibrous annulus, with pale cells separated from the myocardium by thin, fibrous bands. At atrial myocardium, V ventricular myocardium (Masson trichrome, 2×)

chemical stains are used [26, 27]. In general, the cell counts are low in normal heart tissue [27]. One study showed a bimodal distribution of both B cells and T cells during fetal life, with peaks noted at 22 and 37 weeks of gestation [26]. Furthermore, the counts were comparable in the right ventricle, left ventricle, and interventricular septum [26]. In contrast, the number of histiocytes identified by CD68 staining remained relatively constant throughout gestation [26]. A focal ventricular epicardial lymphocytic infiltrate that extends into the superficial myocardium, particularly at the AV groove, has been described in association with ­aneuploidy (trisomy 21 and triploidy) and in other fetuses with anomalies [28]. The significance of the presence of immune cells in the fetal myocardium is poorly understood, but they may play a role in host defense, homeostasis, and/or tissue repair. Large numbers of lymphocytes or clusters with or without myocyte damage may represent congenital myocarditis, which is rare but can occur.

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L. M. Ernst and M. K. Fritsch SVC

Summit of Right atrium SA node Aorta

Right atrial appendage

Fig. 1.24  Dissection of the SA node. This diagram is a view of the superior part of the right atrium where the superior vena cava (SVC) enters it. The SA node (in tan) is located near the junction of the SVC at the summit of the right atrium, identified by following the tip of the right atrial appendage toward the SVC.  The dotted lines indicate the portion of tissue that should be excised to remove the SA node. This removed tissue should be serially sectioned parallel to the ridge of the atrial appendage and entirely submitted

Fig. 1.26  Sampling of the AV node and bundle of His. This diagram shows the specimen after removal as performed in Fig. 1.25. The dotted lines represent the direction of the serial cuts needed before submitting this tissue in its entirety

Coronary sinus

Fig. 1.25  Dissection of the AV node and bundle of His. This is an image of the opened right atrium and ventricle. The opening of the coronary sinus (CS) and the patent foramen ovale (FO) are important landmarks. The AV node is situated within the triangle of Koch, the boundaries of which are the coronary sinus orifice, tendon of Todaro, and the septal leaflet of the tricuspid valve. The tendon of Todaro, which forms the hypotenuse of the triangle, connects the valve of the inferior vena cava (Eustachian valve) to the central fibrous body of the heart. The four cuts needed to remove the AV node and bundle of His are indicated by the box

Membranous septum

Tricuspid valve

References 1. Sadler TW, Langman J.  Langman’s medical embryology. 11th ed. Philadelphia: Wolters Kluwer Lippincott Williams & Wilkins; 2009. 2. Schoenwolf GC, Larsen WJ. Larsen’s human embryology. 4th ed. Philadelphia: Elsevier/Churchill Livingstone; 2009. 3. Sylva M, van den Hoff MJB, Moorman AFM. Development of the human heart. Am J Med Genet. 2014;64A:1347–71. 4. Schleich JM, Abdula T, Summers R, Houyel L.  An overview of cardiac morphogenesis. Arch Cardiovasc Dis. 2013;106:612–23.

5. Rana MS, Christoffels VM, Moorman AFM.  A molecular and genetic outline of cardiac morphogenesis. Acta Physiol. 2013;207:588–615. 6. van Weerd JH, Christoffels VM. The formation and function of the cardiac conduction system. Development. 2016;143:197–210. 7. Christoffels VM, Moorman AFM. Development of the cardiac conduction system: why are some regions of the heart more arrhythmogenic than others? Circ Arrhythm Electrophysiol. 2009;2:195–207. 8. Watanabe M, Rollins AM, Polo-Parada L, Ma P, Gu S, Jenkins MW.  Probing the electrophysiology of the developing heart. J Cardiovasc Dev Dis. 2016;3:pii:E10. https://doi.org/10.3390/ jcdd3010010.

1 Heart 9. Temple IP, Inada S, Dobrzynski H, Boyett MR. Connexins and the atrioventricular node. Heart Rhythm. 2013;10:297–304. 10. Aanhaanen WT, Moorman AF, Christoffels VM. Origin and development of the atrioventricular myocardial lineage: insight into the development of accessory pathways. Birth Defects Res A Clin Mol Teratol. 2011;91:565–77. 11. Di Nardo P, Forte G, Ahluwalia A, Minieri M. Cardiac progenitor cells: potency and control. J Cell Physiol. 2010;224:590–600. 12. Berry GJ, Billingham ME.  Normal heart. In: Mills SE, edi tor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2011. p. 563–83. 13. Favara BE, Moores H.  Anitschkow nuclear structure: a study of pediatric hearts. Pediatr Pathol. 1987;7:151–64. 14. Murphy GE, Becker CG.  Occurrence of caterpillar nuclei within normal immature and normal appearing and altered mature heart muscle cells and the evolution of Anitschkow cells from the latter. Am J Pathol. 1966;48:931–57. 15. Wagner BM, Siew S. Studies in rheumatic fever. V. Significance of the human Anitschkow cell. Hum Pathol. 1970;1:45–71. 16. Boyd TA. Blood cysts on the heart valves of infants. Am J Pathol. 1949;25:757–9. 17. Zimmerman KG, Paplanus SH, Dong S, Nagle RB.  Congenital blood cysts of the heart valves. Hum Pathol. 1983;14:699–703. 18. Titus JL. Normal anatomy of the human cardiac conduction system. Mayo Clin Proc. 1973;48:24–30. 19. Suarez-Mier MP, Gamillo C. Atrioventricular node fetal dispersion and His bundle fragmentation of the cardiac conduction system in sudden cardiac death. J Am Coll Cardiol. 1998;32:1885–90.

19 20. Valdes-Dapena MA.  Histology of the fetus and newborn. Philadelphia: Saunders; 1979. 21. Valdes-Dapena MA, Greene M, Basavanand N, Catherman R, Truex RC. The myocardial conduction system in sudden death in infancy. N Engl J Med. 1973;289:1179–80. 22. Ottaviani G, Buja LM. Anatomopathological changes of the cardiac conduction system in sudden cardiac death, particularly in infants: advances over the last 25 years. Cardiovasc Pathol. 2016;25: 489–99. 23. Hill DA, Swanson PE. Myocardial extramedullary hematopoiesis: a clinicopathologic study. Mod Pathol. 2000;13:779–87. 24. Fishbein GA, Fishbein MC, Buja LM.  Myocardial ischemia and its complications. In: Buja LM, Butany J, editors. Cardiovascular pathology. 4th ed. London: Elsevier; 2016. p. 249. 25. Rosenberg HS, Donnelly WH.  The cardiovascular system. In: Wigglesworth JS, Singer DB, editors. Textbook of fetal and perinatal pathology. 2nd ed. Malden: Blackwell Science; 1998. p. 653. 26. Nield LE, von Both I, Popel N, Strachan K, Manlhiot C, Shannon P, et al. Comparison of immune profiles in fetal hearts with idiopathic dilated cardiomyopathy, maternal autoimmune-associated dilated cardiomyopathy and the normal fetus. Pediatr Cardiol. 2016;37:353–63. 27. Fernandes NM, Taylor GP, Manlhiot C, McCrindle BW, Ho M, Miner SE, et al. The myocardium of fetuses with endocardial fibroelastosis contains fewer B and T lymphocytes than normal control myocardium. Pediatr Cardiol. 2011;32:1088–95. 28. Roberts DJ, Genest D. Cardiac histologic pathology characteristic of trisomies 13 and 21. Hum Pathol. 1992;23:1130–40.

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Blood Vessels and Lymphatic Vessels Linda M. Ernst and Michael K. Fritsch

Introduction

Embryology

The primary function of blood vessels (arteries and veins) is to distribute oxygen and nutrients to tissues throughout the body and to transmit carbon dioxide and metabolic waste material to excretory organs for removal. The lymphatic vessels are important for maintaining interstitial fluid balance and returning excess fluid to the venous system. The lymphatic vessels are also involved in presenting foreign material to the immune system via the lymph nodes. Development of the vascular tree is essential for the formation of nearly every organ during embryonic and fetal life. In the fetus, there are some unique features of the vascular tree that facilitate the fetoplacental circulation in utero, such as the umbilical vein, umbilical arteries, ductus venosus, and ductus arteriosus. Understanding the morphologic features of these specialized vascular structures is important for the interpretation of pathologic changes that may occur. In addition, the fetal and neonatal vascular tree is free of the typical aging phenomena seen in the adult vasculature, so the histologic features of many fetal blood vessels fit those of classically described “normal” vessels without superimposed pathologic changes. However, pathologic changes of the vasculature and lymphatics can occur during the fetal and neonatal period, so understanding their normal structure is helpful.

The cardiovascular system is the first functional organ system established in the developing embryo. Blood vessels begin to arise from mesoderm during gastrulation (week 3 of human development). The vascular system develops via two processes: vasculogenesis, which is the de novo establishment of blood vessels from blood islands with no prior existing blood vessels, and angiogenesis, which involves the budding, branching, or sprouting of new blood vessels from preexisting vessels [1, 2]. The first blood islands appear during the third week of development in mesoderm that will give rise to extraembryonic blood vessels, including near the umbilical vesicle (yolk sac), connecting stalk, allantois, and chorion. A few days later, blood islands form within the lateral plate mesoderm and other areas in the embryo that will give rise to intraembryonic blood vessels. Blood islands seem to arise from mesoderm adjacent to endoderm (splanchnopleuric mesoderm), not from mesoderm situated near ectoderm (somatopleuric mesoderm). Blood islands are derived from mesodermal cells that will differentiate into hemangioblasts. Hemangioblasts are partially differentiated precursor cells capable of giving rise to either hematopoietic cells or endothelial cells. Blood islands consist of an outer cell layer destined to differentiate into endothelial cells and a collection of inner cells that will give rise to hematopoietic cells, usually erythroid when associated with endothelial cells. The precursor cell committed to form an endothelial cell is referred to as an angioblast. Blood islands begin to coalesce to form lumens, thereby establishing a capillary network within the developing embryo. As the capillary network enlarges, angiogenesis (sprouting of new vessels) contributes to network expansion. There is also evidence that some endothelial cells retain their ability to give rise to hematopoietic cells; these are referred to as hemogenic endothelium. Blood vessel formation precedes hematopoiesis. Vasculogenesis is responsible for establishing the primary vascular beds,

L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected] M. K. Fritsch Department of Pathology and Laboratory Medicine, University of Wisconsin, Madison, Madison, WI, USA e-mail: [email protected]

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_2

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including the paired dorsal aortae, cardinal veins, and primary vascular plexus of the yolk sac, which join together to form a closed vascular loop. Once established, angiogenesis expands the vascular network by giving rise to additional vessels (mostly mediated via vascular endothelial growth factor (VEGF) signaling). Eventually, once the heart begins to beat at about day 21–22 and circulation is established, mechanical remodeling of the vascular beds occurs via the additional actions of mechanical shear forces acting on the vessels, with some vessels closing down and others being redirected, especially during the rotation of the heart [1–4]. The molecular regulation of early blood vessel formation is complex and involves multiple signaling pathways and molecules. Mesoderm formation is dependent on both fibroblast growth factor and transforming growth factor-beta signaling. A particular transcription factor, ETS translocation variant 2 (ETV2), a member of the ETS family of genes also known as ER71, has been identified as a transient obligatory factor regulating hemangioblast differentiation. ETV2 expression in the hemangioblast is regulated by bone morphogenic protein (BMP), Notch, and Wnt protein signaling pathways in mesoderm cells. ETV2 is only expressed transiently in the hemangioblast, but its expression is mandatory for hematopoietic and endothelial cell differentiation. ETV2, a DNA-binding transcription factor, appears to be a master regulator that initiates transcription of numerous other genes involved in endothelial differentiation, including other members of the ETS family of transcription factors and members of the VEGF signaling pathways, including both ligands and receptors. VEGF and its receptor VEGFR2 then become the most important drivers of embryonic vessel formation from the hemangioblast. VEGFR2 is expressed in the hemangioblast and endothelial cells but not in hematopoietic progeny. After the heart begins to beat and early blood flow is established, specification of arteries and veins begins. Lymphatic differentiation will not occur until later in embryonic development, at the end of the fifth week [4]. Arteries and veins are defined by the direction of blood flow, either away from the heart (arterial) or toward the heart (venous). Arterial and venous endothelial cells express specific molecular markers. Molecular markers of arterial endothelial differentiation include expression of ephrin-B2 (EFNB2), neuropilin-1 (NRP1), and members of the Notch pathway. Molecules expressed specifically in the venous system include arterial ephrin-B2 receptor and the neuropilin-2 receptor. Also in veins, Notch signaling is repressed by the expression of COUP-TFII (chicken ovalbumin upstream promoter transcription factor 2), which in humans is encoded by the nuclear receptor subfamily 2, group F, member 2 gene (NR2F2), which prevents activation of an “arterial” phenotype. The exact pathways specifying and maintaining the venous and arterial endothelial phenotypes still require further research. Data from certain model systems suggest that

L. M. Ernst and M. K. Fritsch

arterial and venous fates for endothelial cells may be predetermined from a very early developmental stage (angioblasts), but other data indicate some degree of plasticity such that artery and vein endothelium can interconvert, at least early in development. In the yolk sac blood vessels, the direction of blood flow seems to be a critical regulator of arterial-venous endothelial differentiation [2, 3]. In the early embryo, three major vascular networks—the vitelline, embryonic, and placental vasculature—join together. Each has its own arterial and venous component. The vitelline vascular system is composed of the vitelline arteries (which will fuse and give rise to the superior mesenteric artery in the adult), which arise from the dorsal aorta and pass through the vitellointestinal duct (yolk sac stalk). They empty into a capillary bed drained by the vitelline veins, which will contribute to the adult portal system and which pass back through the yolk sac stalk to enter the caudal end of the cardiac tube via the sinus venosus. The embryonic vessels will form the majority of the fetal and adult cardiovascular system. Its arterial components include the dorsal aorta, aortic arches, and umbilical arteries. The dorsal aorta is initially paired and later fuses from thoracic level 4 to lumbar level 4. The pharyngeal arch arteries connect the dorsal aorta to the ventral aorta. The caudal dorsal aorta gives rise to the umbilical arteries and laterally to intersegmental arteries. Three pairs of veins empty into the sinus venosus of the heart, including the vitelline veins, umbilical veins from the placenta (of which only the left will persist as the single umbilical vein in the fetus), and cardinal veins (anterior, common, and posterior). Placental blood vessels form initially in the connecting stalk (later to become the umbilical cord) and anastomose in the chorion. The paired umbilical arteries carry deoxygenated blood from the dorsal aorta and metabolic waste products to the placental villi, while the initially paired umbilical veins carry oxygenated blood and nutrients to the embryo via the sinus venosus. Although early establishment of the vascular network and specification of arteries and veins seem to be regulated by genetic programs and endothelial cell differentiation, later remodeling of the entire vascular network is highly dependent on blood flow patterns and hemodynamic signals, including pressure, perfusion, and flow dynamics. Blood flow patterns probably have direct mechanical effects on vessels, as well as altering genetic programs within endothelial cells. The end result of the remodeling process is the fetal vasculature, which is similar to the adult vasculature except for the presence of two vascular shunts, the ductus arteriosus and ductus venosus, and an intracardiac shunt, the foramen ovale. The umbilical vein and the two umbilical arteries are also unique to intrauterine life and regress quickly following birth and the cutting of the umbilical cord [1–4]. Lymphatic vessels begin to appear in the developing human around the end of the fifth week of embryonic

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d­ evelopment. There are two possible models for lymphatic formation. The favored model is direct budding of lymphatic endothelium from veins. The second model is the formation of multiple clefts in the mesenchyme lined by endothelial cells with a lymphatic phenotype, which coalesce to form the lymphatic complex and then join the venous vascular complex by endothelial cell sprouting and/or migration. During weeks 6–9 of development, six lymphatic sacs form, including two jugular lymph sacs near the junction of the subclavian veins with the anterior cardinal veins (the future internal jugular vein), two iliac lymph sacs near the junction of the iliac veins with the posterior cardinal veins, one retroperitoneal lymph sac in the root of the mesentery on the posterior abdominal wall, and one cisterna chyli dorsal to the retroperitoneal lymph sac, at the level of the adrenal glands. The lymphatic vessels grow along the major veins, and the jugular lymph sacs connect to the cisterna chyli via the right and left thoracic ducts. These two ducts will partially fuse in the middle portions, and the final thoracic duct is composed of a portion of the right thoracic duct caudally, the anastomosed portion of the ducts in the middle, and the cranial portion of the left thoracic duct. The final right thoracic duct represents the remaining cranial portion of the original right duct. The final main (left) thoracic duct, which is established by early fetal life, will join the venous system at the angle of the joining of the left internal jugular vein and the left subclavian vein. Molecularly, the earliest marker for the developing lymphatic endothelial cell is prospero-related homeobox protein 1 (PROX1). Absence of PROX1 results in a complete absence of lymphatic formation. Therefore, PROX1 is essential for lymphatic development and has been shown to be important for the maintenance of the lymphatic endothelial phenotype in adulthood. Other genes are also important in establishing the final lymphatic endothelial phenotype [5–7].

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a

b

Fig. 2.1  Fetal aorta at 23 weeks gestation. In this low-power view, note the bulky tunica media and the abundant elastic fibers within the tunica media (a H&E, 4×; b Verhoeff-van Gieson, 4×)

Histology Aorta The aorta of fetuses and newborns has the same anatomical structure and mural features that are present in adulthood. The innermost layer, the tunica intima, is composed of a simple squamous layer of endothelial cells with very little additional supporting intimal fibrous tissue. The presence of fibrointimal thickening or foamy macrophages within the intima is considered abnormal in the fetal or newborn period. The tunica media comprises the bulk of the aortic wall and is composed of densely packed circumferential layers of elastic tissue with intervening collagen (Figs.  2.1 and 2.2). The nuclei present in the media consist predominantly of smooth

Fig. 2.2  Fetal aorta at 23  weeks gestation. This higher-power view highlights the darkly stained elastic fibers within the tunica media (Verhoeff-van Gieson, 20×)

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a

b

Fig. 2.3  Fetal aorta at term. (a) A medium-power view of the fetal aorta at 38 weeks gestation is shown, with the thick tunica media containing abundant elastic fibers. Note the thicker and more compact tunica adventitia at this gestational age, containing vasa vasorum (H&E, 10×). (b) Elastic stain of fetal aorta at 38  weeks gestation is shown. Note the thick tunica media with abundant elastic fibers. The thick and compact tunica adventitia at this gestational age contains vasa vasorum (black arrow) and a large branch artery (white arrow) (Verhoeff-van Gieson, 20×)

muscle cells and fibroblasts. The wall is free of inflammatory cells in the normal state. The outermost layer is the tunica adventitia, which consists of loose fibrous connective tissue containing vessels of the vasa vasorum (Fig. 2.3).

Ductus Arteriosus The ductus arteriosus is the vascular structure in fetal life that shunts the relatively oxygenated blood from the right ventricle/pulmonary artery to the descending aorta, thus providing oxygenated blood to the systemic circulation and bypassing the pulmonary circuit. The ductus arteriosus is a

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a

b

Fig. 2.4  Ductus arteriosus in the midtrimester, at 21 weeks gestation. Note the features of a muscular artery, including intima, internal elastic lamina, media, and adventitia. Even at this gestational age, focal intimal thickening is present (a H&E, 10×; b Verhoeff-van Gieson, 10×)

muscular artery with an intima, media, and adventitia. The media of the ductus arteriosus lacks the tightly packed elastic fibers of the aorta, but it does have some elastic fibers, with a prominent internal elastic lamina (Fig.  2.4). The intima of the ductus arteriosus may show slight thickening during fetal life, and it has been shown that the histological changes characteristic of closure of the ductus arteriosus begin before birth [8] (Fig.  2.5). Following birth, the ductus arteriosus functionally closes within 1–2 days, but it does not structurally close for several weeks. The morphological changes seen with the closure of the ductus arteriosus (Fig.  2.6) include (1) increased thickness of the intima, with formation of intimal cushions; (2) increased thickness of the media; (3) thickening and fragmentation of the internal elastic lamina; (4) increased ground substance and connective tissue in the intima and media; and (5) the appearance of spaces in the inner part of the media [8].

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Umbilical Vein

formation of a fibrous cord which becomes the round ligament or ligamentum teres of the liver.

Though not usually seen in the examination of the adult abdomen, the umbilical vein is easily identified in the internal pathologic examination of all fetuses as the vascular structure extending from the umbilicus to the liver. In the midtrimester, the vein is a thin-walled, easily collapsible structure; it appears thicker in fetuses near term and in neonates. The vein has an inner intimal layer, a fairly thick media composed of haphazardly arranged smooth muscle fibers, and a relatively thick adventitia (Figs. 2.7 and 2.8) [9]. The umbilical vein is frequently catheterized to gain vascular access, especially in premature neonates, so it is subject to pathologic conditions such as endothelial injury, thrombosis, or even rupture. The umbilical vein involutes after birth, with obliteration of the lumen and

Fig. 2.5  Ductus arteriosus at term. A low-power view of a complete cross section of the ductus arteriosus in a term fetus. Note the thick media, smaller lumen, and areas of focal intimal thickening. Compare with Figs. 2.4 and 2.6 (H&E, 2×)

a

b

Fig. 2.6  Ductus arteriosus after birth. The ductus arteriosus of a term infant who lived for 6 days is shown. Note the features of closure of the ductus arteriosus, including increased thickness of the intima with for-

Umbilical Arteries The two umbilical arteries arise from the internal iliac arteries and can be identified in the fetus as they course along each side of the urinary bladder toward the umbilicus, where they enter the Wharton’s jelly of the umbilical cord. The umbilical arteries, therefore, transmit systemic, relatively deoxygenated blood from the fetus to the placenta. Within the fetal body, the two umbilical arteries are typically the same size and increase mildly in size over gestation. When only a single umbilical artery is present within the umbilical cord, there is usually only one umbilical artery noted during

Fig. 2.7  Umbilical vein. Low-power view of the umbilical vein from a 35-week stillborn fetus. The endothelial lining has sloughed, but note the thick media composed of collagen fibers with haphazardly arranged smooth muscle (Masson trichrome, 2×)

c

mation of intimal cushions (a); fragmentation of the internal elastic lamina (b); and increased connective tissue in the intima (c). (a H&E, 2×; b Verhoeff-van Gieson, 2×; c Masson trichrome, 2×)

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b

Fig. 2.8  Umbilical vein at term. Note the features of the umbilical vein at term, including a lining endothelium, an inner intimal layer, and a fairly thick media composed of haphazardly arranged smooth muscle. No internal elastic lamina is seen (a H&E, 10×; b Verhoeff-van Gieson, 10×)

Fig. 2.9  Umbilical artery within the fetal body at term (stillbirth at 37  weeks gestation). The umbilical artery in this section contains a prominent internal elastic lamina and two distinct muscular layers in the media (pentachrome stain, 4×)

Fig. 2.10  Umbilical artery within the fetal body at 5 days of life (born at 32 weeks gestation). The media lacks an internal elastic lamina, but degenerating elastic fibers are seen in the media. Intraluminal fibrin is present (pentachrome stain, 4×)

the fetal examination as well. The umbilical artery may be absent on either the left or the right side, and this is easily recognized when examining the umbilical artery course along the side of the urinary bladder. Aberrant umbilical artery origin from the abdominal aorta has also been described [10]. The histology of the umbilical arteries within the fetal body (i.e., between the internal iliac arteries and the umbilical cord) is not well described, but the umbilical artery within the umbilical cord is typically devoid of an internal elastic lamina. From personal observations, we have noted that some umbilical arteries within the fetal body have a well-­ developed internal elastic lamina and some do not (Figs. 2.9 and 2.10). This difference may be related to how close the sections are taken to the systemic arteries versus the umbili-

cal cord, or it may be related to autolysis, but there are no definitive studies examining the morphologic transition from systemic arteries to umbilical arteries. Within the umbilical cord proper, the umbilical arteries are characterized by two distinct muscular layers, an outer circular layer and an inner spiral longitudinal layer; they lack a well-defined internal elastic lamina (see Fig. 36.20). Like the umbilical vein, the umbilical arteries are frequently catheterized, especially to gain vascular access in premature neonates, so they are subject to pathologic conditions such as endothelial injury, thrombosis, or even rupture. The umbilical arteries involute after birth (Fig.  2.11), and their proximal portions become the hypogastric arteries. The distal portions become fibrous ligaments known as the lateral umbilical ligaments [9].

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Fig. 2.11  Umbilical artery at 3 weeks of life (born at 26 weeks gestation). The lumen of the artery (center) is closing down, and there is a discontinuous, smooth muscle layer surrounding the lumen (H&E, 4×)

Fig. 2.13  Ductus venosus at 36 weeks gestation. Higher-power view of the ductus venosus at near term. The wall is thicker and more welldefined than in Fig.  2.12. The wall is composed of thicker collagen bands. Occasional smooth muscle can be present, but none is seen in this image (H&E, 40×)

Ductus Venosus The ductus venosus is the vascular shunt present in fetal life within the liver that allows oxygenated blood from the umbilical vein to enter the inferior vena cava and then proceed to the right side of the heart. It can be identified grossly by opening the umbilical vein into the sinus intermedius of the liver (see Fig. 5.2). The ductus venosus arises from the rostral wall of the sinus intermedius and typically has a smaller diameter than the umbil-

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Fig. 2.12  Ductus venosus at 18 weeks gestation. The ductus venosus (DV) as shown here is extremely thin-walled at this gestational age. The wall is composed of loosely arranged collagen fibers, which in places merge imperceptibly with the surrounding connective tissue. Smooth muscle is not appreciated as a component of the wall in this image. The ductus venosus has a flattened endothelial lining. Note the hepatic parenchyma in the upper left (H&E, 20×)

ical vein. The inner lining of the ductus venosus is usually smooth gray-white, and it has a very thin wall. The histology is characterized by an endothelial lining, occasional smooth muscle cells, and a thin, fibrous wall [11] (Figs. 2.12 and 2.13). The inlet of the ductus venosus commonly has a grossly visible lip or rim, sometimes referred to as a sphincter. One group argued that it is not a true sphincter because in their analysis it lacked smooth muscle, but they did demonstrate a condensation of elastic fibers forming a shelf-like structure. They postulated that the elastic tissue at the inlet of the ductus venosus may accelerate blood flow from the sinus intermedius into the ductus venosus and that the shelf may also reduce back flow [11]. The ductus venosus functionally closes within a few minutes of birth and the cutting of the umbilical cord because the umbilical vein blood flow ceases, but the ductus venosus structurally closes at approximately 4–6  days after birth. It generally closes at the low end of this range in term neonates and at the higher end of the range in premature infants [12]. Agenesis of the ductus venosus can have deleterious effects on fetal blood flow and is associated with hydrops fetalis [13].

Other Fetal Veins and Arteries The general organization of most nonspecialized fetal arteries and veins is similar to that seen in the same vascular structures in adults [9]. Muscular arteries, such as the renal arteries or other arterial branches arising from the aorta (Fig. 2.14), are characterized by a very thin intimal layer consisting almost exclusively of the endothelial cell layer. Immunohistochemical studies have shown that endothelial cells mark with CD31, CD34, von Willebrand factor, and Fli-­1, but expression may vary by organ

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Fig. 2.14  Portal vein and hepatic artery at 35 weeks gestation. Fetal veins and arteries have histologic features similar to their adult counterparts. These are sections of medium- to large-sized fetal vessels, the portal vein (PV) and the hepatic artery (HA). The PV is one of the larger veins of the body and thus has recognizable smooth muscle media composed of somewhat haphazardly arranged smooth muscle, elastic fibers, and collagen. An internal elastic lamina is not present in the PV. However, the internal elastic lamina can be seen in the HA as an undulating dark layer highlighted by the elastic stain. The media of the HA is a well-organized smooth muscle layer. The adventitial collagen is yellow in this stain (pentachrome stain, 10×)

and even by organ compartment [14]. The media is the middle layer composed of the often undulating-appearing internal elastic lamina just beneath the endothelium and the muscular coat composed of smooth muscle cells. The vascular smooth muscle runs in spiral or circumferential patterns and is intermixed with collagen fibrils and elastin fibers [15]. A variable external elastic lamina may be present (Fig. 2.15). The tunica adventitia of fetal muscular arteries is typically fibrous in nature [9]. Larger arteries branch and become smaller arterioles, which ultimately terminate as capillaries. Capillaries consist of a single layer of endothelium resting on a basement membrane and lack a muscular media or elastic fibers [15]. Although capillaries have no fibrous support or adventitia, pericytes are present in and among the basement membrane [15]. Pericytes provide structural support for the capillary and can be highlighted by an immunostain for smooth muscle actin. Because they have contractile properties, they may also be involved in the regulation of blood flow [15]. In some sites, such as the liver (see Fig. 5.11) and fetal adrenal cortex (see Fig. 20.13), the equivalent of capillaries is known as sinusoids. Sinusoids are generally more dilated than capillaries, their endothelium may have prominent gaps or fenestrations, and in some organs, there is no associated basement membrane [15]. The venous structures of the fetus—even large veins such as the inferior vena cava and renal veins—are very thin-­

L. M. Ernst and M. K. Fritsch

Fig. 2.15  Small artery and vein at term. This image shows an example of a smaller artery (A) and associated veins (V). The artery demonstrates well-defined internal and external elastic laminae by the elastic stain. The media of the artery, between the two elastic laminae, is a well-organized smooth muscle layer. The adventitial collagen (pink) is fairly thick in this image. The adjacent thin-walled veins do not have recognizable smooth muscle in the media, which is composed of elastic fibers and collagen. An internal elastic lamina is not present in the veins (elastic stain, 4×)

Fig. 2.16  Inferior vena cava at 32 weeks gestation. This is the largest vein of the fetal body and thus has recognizable smooth muscle media, which is composed of somewhat loosely arranged smooth muscle fibers and collagen. An internal elastic lamina is not present (pentachrome stain, 10×)

walled structures that may be easily overlooked grossly, especially in the midtrimester. Similar to arteries, veins have three distinct layers, but the media typically lacks an internal or external elastic lamina, is thinner, and has fewer, more haphazardly arranged muscle fibers (Fig. 2.16). In some cases,

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very little muscle tissue is seen in the media, and the wall is composed mostly of fibrous tissue (see Fig. 2.15) [15].

Lymphatic Vessels Lymphatic vessels are present in most fetal tissues and typically present as thin-walled vascular structures with a flattened, endothelium-like lining that can be highlighted by immunohistochemistry for D2-40, PROX1, or LYVE-1. Small lymphatic channels closely resemble capillaries with no significant muscular media. Larger lymphatic channels have a thin, muscular media with no distinction into circular or longitudinally oriented layers. The thoracic duct and right lymphatic duct have a longitudinal muscular layer [15] (Fig. 2.17). In stillbirths, the lymphatic endothelium is often sloughed, thereby making immunohistochemistry of limited value. Lymphatic channels may contain clusters of small lymphocytes, which can sometimes be helpful in distinguishing them from capillaries.

Special Considerations Hyrtl’s Anastomosis Hyrtl’s anastomosis, defined as a connection between the two umbilical arteries, is very common in normal pregnancies [16, 17]. The types of connections include a true vessel between the two arteries, a fenestration between the two arteries, or a fusion of segments of the arteries. These anastomoses may occur

Fig. 2.17  Thoracic duct at 26 weeks gestation. The thoracic duct is the collapsed structure seen between the three arrows. Note that it has a thin, longitudinal muscular layer and a lymphatic endothelial lining that is nearly imperceptible at this power (H&E, 10×)

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within the umbilical cord at a median distance of 7–16  mm from the cord insertion, at the cord insertion itself, or on the placental surface [17]. Hemodynamic studies have shown that Hyrtl’s anastomoses function as either a safety valve or a pressure equalizer between the two umbilical arteries [16].

 ystic Hygroma and Other Lymphatic C Malformations Cystic hygroma is a clinical term used to describe a cystic neck mass in the posterior cervical space. By ultrasound, a cystic hygroma is a large, fluid-filled structure with septations in the posterior neck; it can be a sign of aneuploidy in a second-trimester ultrasound [18, 19]. The aneuploidy most commonly associated with cystic hygroma is monosomy X (Turner syndrome), but other aneuploid conditions can also present with a cystic hygroma [19]. Cystic hygroma may be present with other signs of fetal hydrops, such as subcutaneous edema and effusions. Pathologically, what is called cystic hygroma clinically arises as a result of obstruction of lymphatic flow secondary to lymphatic malformations such as hypoplasia or failure of fusion between lymphatic primordia and the jugular vein [20]. Most fetuses with cystic hygroma seen on ultrasound have a large posterior nuchal fluid-filled sac that has ill-­ defined borders and is not easily separated from the overlying (often edematous) skin. This lesion is composed of multiple irregular spaces with abundant, paucicellular intervening connective tissue (Fig. 2.18). The spaces may or may not have a lining, but some lymphatic channels can be seen

Fig. 2.18  Cystic hygroma. This is a section from a cystic hygroma in a 26-week fetus with hydrops fetalis. Note the numerous dilated lymphatic spaces (LS), three of which are indicated in this photomicrograph. Loose, fibrous connective tissue is noted between the spaces, and no obvious smooth muscle is seen. The cells lining the spaces are mostly sloughed to imperceptible at this power. Cartilaginous tissue of the neck is seen in the lower left and upper right corners (H&E, 10×)

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using D2-40 or PROX1 immunohistochemistry. Because most of the fluid is in the connective tissue and not in dilated lymphatic channels, this lesion has also been referred to as postnuchal fluid accumulation [20]. Recently, the more general term lymphatic malformation has been applied to a variety of lesions with a lymphatic component. These lesions are further subdivided descriptively as localized, macrocystic, microcystic, or combined. Lymphatic malformations contain multiple variably sized lymphovascular channels with flattened endothelium. Intervening connective tissue is fairly scanty and may contain smooth muscle and adipose tissue. The spaces may contain lymphocytes [20, 21].

L. M. Ernst and M. K. Fritsch

abdominal para-aortic bodies increase in size up to 3 years of life and then undergo degenerative changes marked by increasing stroma. The process appears to be complete by 14 years of age [23]. Paraganglia are composed of neuroendocrine cells of neural crest origin and are also known as extra-adrenal chromaffin cells. On routine H&E sections, paraganglia can be recognized as encapsulated or nonencapsulated collections of fairly uniform polygonal cells with granular cytoplasm and small, oval nuclei without nucleoli, arranged in nests and cords with prominent intervening vasculature. The chromaffin cells of paraganglia can be highlighted by typical neuroendocrine stains such as chromogranin and synaptophysin (Figs.  2.20 and 2.21). Paraganglia are often associated with sympathetic

Periaortic/Retroperitoneal Soft Tissues In routine sections of the aorta, or even of the kidney and adrenal glands, retroperitoneal soft tissue is often present. This tissue can contain specialized structures such as lymph nodes, paraganglia, and sympathetic ganglia (Fig.  2.19). Because these tissues can be especially prominent in fetuses, it is important to be aware of their histologic appearance, so as not to confuse them with a pathologic abnormality. Paraganglia are seen very frequently in the fetal retroperitoneum near the sympathetic trunk and close to sympathetic ganglia. They can sometimes attain large size and be encapsulated, especially in the periaortic region, which has led to the term para-aortic bodies. In the early 1900s, Zuckerkandl described masses of extra-adrenal chromaffin cells in close association with the inferior mesenteric artery (organs of Zuckerkandl), but similar tissue has been described at multiple sites along the aorta [22]. Studies have shown that

Fig. 2.19  Periaortic soft tissues in a term fetus. This low-power image shows some of the specialized structures present in periaortic soft tissue, including lymph node (LN), encapsulated paraganglion (P), and sympathetic ganglion (G) (H&E, 4×)

a

b

Fig. 2.20  Comparison of paraganglion and ganglion. (a) Paraganglion: Note the uniform, polygonal neuroendocrine cells with abundant cytoplasm and small oval nuclei without nucleoli arranged in nests and cords with prominent intervening vasculature. (b) Sympathetic ganglion: Note the more heterogeneous appearance, with numerous ganglion cells characterized by abundant eosinophilic cytoplasm and eccentrically placed large nuclei, many with nucleoli. The lightly eosinophilic background of ganglia is composed of wavy nerve fibers (H&E, 20×)

2  Blood Vessels and Lymphatic Vessels

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a

b Fig. 2.22  Sympathetic ganglion in the retroperitoneal tissues at 16  weeks gestation. The ganglion is composed of closely packed, immature-appearing ganglion cells at this gestational age. At this high power, the development of the eccentric eosinophilic cytoplasm and even nucleoli can be seen. The wavy stroma seen in the more mature ganglion (as in Fig.  2.20) is not as prominent (H&E, 40×)

References

Fig. 2.21  Paraganglion in the retroperitoneal tissues at 29 weeks gestation. (a) This paraganglion is slightly better preserved than the term example in Fig. 2.20. Note the uniform, polygonal neuroendocrine cells with slightly granular cytoplasm and small oval nuclei without nucleoli, arranged in nests and cords with prominent intervening vasculature. Intervening fibrous tissue is noted (H&E, 20×). (b) Chromogranin immunostain confirms the neuroendocrine origin of these cells (chromogranin immunohistochemistry, 20×)

ganglia, which provide a stark histologic contrast. Sympathetic ganglia appear more heterogeneous than paraganglia at low power and are punctuated by numerous ganglion cells with abundant eosinophilic cytoplasm and eccentrically placed large nuclei, many with nucleoli. The lightly eosinophilic background of ganglia is composed of wavy nerve fibers (see Fig. 2.20). In first- and second-­trimester fetuses, sympathetic ganglia can appear immature and should not be confused with a neuroblastic tumor (Fig. 2.22).

1. Bautch VL, Caron KM.  Blood and lymphatic vessel formation. Cold Spring Harb Perspect Biol. 2015;7:a008268. 2. Sadler TW, Langman J.  Langman’s medical embryology. 11th ed. Philadelphia: Wolters Kluwer Lippincott Williams & Wilkins; 2010. 3. Eichmann A, Yuan L, Moyon D, Lenoble F, Pardanaud L, Breant C. Vascular development: from precursor cells to branched arterial and venous networks. Int J Dev Biol. 2005;49:259–67. 4. Sumanas S, Choi K.  ETS transcription factor ETV2/ER71/Etsrp in hematopoietic and vascular development. Curr Top Dev Biol. 2016;118:77–111. 5. Escobedo N, Oliver G.  Lymphangiogenesis: origin, specifi cation, and cell fate determination. Annu Rev Cell Dev Biol. 2016;32:677–91. 6. Semo J, Nicenboim J, Yaniv K.  Development of the lymphatic system: new questions and paradigms. Development. 2016;143: 924–35. 7. Yang Y, Oliver G. Development of the mammalian lymphatic vasculature. J Clin Invest. 2014;124:888–97. 8. Mujahid BM, Gaikwad PG.  A study of histology of human ductus arteriosus: before and after birth. J Anat Soc India. 2000;49(1): 3–5. 9. Valdes-Dapena MA. Blood vessels. In: Valdes-Dapena MA, editor. Histology of the fetus and newborn. Philadelphia: Saunders; 1979. p. 31–41.

32 10. Duesterhoeft SM, Ernst LM, Siebert JR, Kapur RP. Five cases of caudal regression with an aberrant abdominal umbilical artery: further support for a caudal regression-sirenomelia spectrum. Am J Med Genet A. 2007;143A:3175–84. 11. Mavrides E, Moscoso G, Carvalho JS, Campbell S, Thilaganathan B.  The human ductus venosus between 13 and 17 weeks of gestation: histological and morphometric studies. Ultrasound Obstet Gynecol. 2002;19:39–46. 12. Kondo M, Itoh S, Kunikata T, Kusaka T, Ozaki T, Isobe K, et al. Time of closure of ductus venosus in term and preterm neonates. Arch Dis Child Fetal Neonatal Ed. 2001;85:F57–9. 13. Takeuchi M, Nakayama M, Tamura A, Kitajima H. Hydrops fetalis due to agenesis of the ductus venosus: new hepatic histological features. Pediatr Dev Pathol. 2009;12:239–43. 14. Pusztaszeri MP, Seelentag W, Bosman FT.  Immunohistochemical expression of endothelial markers CD31, CD34, von Willebrand factor, and Fli-1 in normal human tissues. J Histochem Cytochem. 2006;54:385–95. 15. Gallagher P, van der Wal A.  Blood vessels. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 233–58. 16. Gordon Z, Eytan O, Jaffa AJ, Elad D.  Hemodynamic analysis of Hyrtl anastomosis in human placenta. Am J Physiol Regul Integr Comp Physiol. 2007;292:R977–82.

L. M. Ernst and M. K. Fritsch 17. Ullberg U, Sandstedt B, Lingman G.  Hyrtl’s anastomosis, the only connection between the two umbilical arteries. A study in full term placentas from AGA infants with normal umbilical artery blood flow. Acta Obstet Gynecol Scand. 2001; 80:1–6. 18. Gaddikeri S, Vattoth S, Gaddikeri RS, Stuart R, Harrison K, Young D, et al. Congenital cystic neck masses: embryology and ­imaging appearances, with clinicopathological correlation. Curr Probl Diagn Radiol. 2014;43:55–67. 19. Sonek J, Croom C.  Second trimester ultrasound markers of fetal aneuploidy. Clin Obstet Gynecol. 2014;57: 159–81. 20. Keeling JW.  Hydrops fetalis and other forms of excess fluid accumulation in the fetus. In: Wigglesworth JS, Singer DB, editors. Textbook of fetal and perinatal pathology. 2nd ed. Malden: Blackwell Science; 1998. p. 358–83. 21. Bruder E, Alaggio R, Kozakewich HP, Jundt G, Dehner LP, Coffin CM. Vascular and perivascular lesions of skin and soft tissues in children and adolescents. Pediatr Dev Pathol. 2012;15(1 Suppl):26–61. 22. Coupland RE.  The prenatal development of the abdominal para-­ aortic bodies in man. J Anat. 1952;86:357–72. 23. Coupland RE. Post-natal fate of the abdominal para-aortic bodies in man. J Anat. 1954;88:455–64.

Part II Respiratory Tract

3

Lung Portia A. Kreiger and Jennifer E. Pogoriler

Introduction The process of fetal development is continuous, but the development of the lung can be subdivided into five morphologic stages: embryonic, pseudoglandular, canalicular, saccular, and alveolar. Based on various authors’ observations, different transition times from one stage to another have been proposed, at least in part because lung development is not completely synchronous. Specifically, lung development is more rapid in cranial regions than in caudal areas [1]. The five developmental stages can be identified by histology, and each stage of lung development is marked by a major developmental milestone (Table 3.1) [1–3].

Embryology The lung arises at approximately 3–4  weeks of gestation from the foregut endoderm as a ventral diverticulum, which is destined to ultimately become the larynx and trachea proximally and the bronchi and lung parenchyma distally. This ventral foregut diverticulum develops within a mass of mesenchyme and forms the primary lung bud by 4 2/7  weeks

postfertilization age (PFA) [4–6]. The primary lung bud bifurcates laterally to form right and left primary bronchial buds at about 4 4/7 weeks PFA and subsequently begins to branch asymmetrically, forming lobar buds (three right and two left) at 5 1/7 weeks PFA, segmental buds at 5 6/7 weeks PFA, and subsegmental buds at 6 2/7 weeks PFA [1, 4, 5]. These sequential endodermal buds carry with them coats of mesenchyme. Within this mesenchyme, islands of precartilage appear within proximal branches concurrent with segmental buds and will ultimately form the cartilage plates of bronchi [4, 5]. In addition, this mesenchyme will ultimately form the muscular and connective tissue components of the airways and the interstitial connective tissue of the alveolar parenchyma [6]. The vascular network of the lung also arises from this mesenchyme [7]. These events correspond to the embryonic stage of lung development [1, 4, 6]. During the pseudoglandular stage of lung development, subsegmental buds branch rapidly and dichotomously to form sequential generations of airways [1, 4, 6]. By 16 weeks of gestation, all pre-acinar airways (i.e., airways to the level of the terminal bronchioles) have formed [4, 6]. This process is accomplished largely through complex epithelial-­mesenchymal interactions. Also, the epithelial lining of these endodermal

Table 3.1  Five stages of lung formation Developmental stage Embryonic Pseudoglandular Canalicular Saccular Alveolar

Age range 3–4 to 6–8 weeks gestational age 6–8 to 16 weeks gestational age 16 to 26–28 weeks gestational age 26–28 to 32–36 weeks gestational age 32–36 weeks to 2–4 years postnatal age

Developmental milestone Proximal airways Distal (pre-acinar) airways Acinusa and primitive capillary network Maturing alveolar-­capillary interface Increasing acquisition of alveoli

The general time ranges and developmental milestones for the five stages tof lung formation. The embryology and histology of the lung are discussed with reference to these five morphologic stages a Acinus encompasses the respiratory bronchioles, alveolar ducts, and terminal sacs P. A. Kreiger · J. E. Pogoriler (*) Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]; [email protected] © Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_3

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buds is beginning to be organized in a proximal-­to-­distal pattern. The more proximal branches will form the airways, and the more distal branches will form the alveolar parenchyma [4]. Within the accompanying mesenchyme, generations of vessels develop alongside sequential generations of branching airways. This process of vasculogenesis forms the accompanying arteries and veins to the level of the terminal bronchiole and is complete by the end of the pseudoglandular stage [4, 7]. The subsequent venous network develops synchronously but independently from the arterial and airway network [4, 7]. Lymphatics arise initially within the hilar regions at approximately 8  weeks of gestation and develop within the lung by about 10  weeks of gestation [1]. Intrapulmonary lymphatics are distributed along the bronchovascular bundles as far as the alveolar duct, along the interlobular septa, and in the pleura. During the ensuing canalicular stage of lung development, lung growth increases substantially. The development of the acinus—i.e., respiratory bronchioles, alveolar ducts, and early airspaces (terminal sacs)—occurs during this time. However, the expansion of the mesenchymal capillary network is also greatest at this time. As this capillary network expands, the mesenchymal bed in which these capillaries grow begins to thin. By the end of the canalicular stage, the formation of primitive capillary-airspace interfaces has begun [1, 6]. During the saccular stage of lung development, increasingly complex airspace development occurs. The more primitive airspaces of the canalicular period are subdivided by the ingrowth of ridges of mesenchymal tissue containing capillaries, known as “secondary crests.” Vasculogenesis continues to increase and mature, and mesenchyme continues to thin. With these events, concomitant progressive differentiation of airspace lining epithelium results in a maturing surface required for postnatal blood-gas exchange [1, 6]. The final, alveolar stage of lung development is characterized by the formation of increasing numbers of mature alveoli with well-formed alveolar-capillary interfaces [1, 7]. The lung at birth contains a reported 24 million alveoli, but alveologenesis continues postnatally. The majority of alveoli are thought to be formed by 2 years of age, and by adulthood the lung contains 300–600 million alveoli [1–3, 6, 7].

Histology  eneral Overview of Postnatal Pulmonary G Histology The fully formed, mature lung consists of three lobes on the right and two lobes on the left. Each lobe is invested by a thin layer of visceral pleura composed of a monolayer of mesothelium and subjacent fibroconnective tissue containing small blood vessels and lymphatics (Fig. 3.1). Each lung is supplied by a primary bronchus, and each lobe is supplied by a secondary bronchus. The pulmonary lobes are further subdivided

P. A. Kreiger and J. E. Pogoriler

Fig. 3.1  Normal 5-month postnatal lung. The visceral pleural surface of the lung consists of a simple layer of mesothelium and subjacent loose fibroconnective tissue, which contains occasional small blood vessels and lymphatics (hematoxylin and eosin [H&E], 20×)

Fig. 3.2  Normal 5-month postnatal lung. This tertiary bronchus is surrounded by fibroconnective tissue, which contains cartilage plates, seromucinous glands, and smooth muscle (H&E, 2×)

into bronchopulmonary segments, each associated with a tertiary bronchus (Figs.  3.2 and 3.3). Each segment is further subdivided into pulmonary lobules, supplied by a terminal bronchiole. Each bronchiole is accompanied by an arteriole. Together, these two structures form a bronchovascular bundle (Fig. 3.4), which is invested in fibroconnective tissue that also contains lymphatics and nerves. Beyond the terminal bronchiole is the pulmonary acinus. The acinus encompasses a respiratory bronchiole and all its branches, including alveolar ducts and terminal alveoli (Figs. 3.5 and 3.6). The perimeter of each lobule is demarcated by interlobular septa (Figs. 3.7 and 3.8), which consist of bands of fibroconnective tissue containing veins or venules and lymphatics [6]. Bronchi are supported by cartilage plates (Figs.  3.9 and 3.10), which are larger proximally and become pro-

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Fig. 3.3  Normal 5-month postnatal lung. Seromucinous glands connect to the respiratory mucosal surface in this high-power image of a tertiary bronchus; smooth muscle and cartilage are also seen (H&E, 10×)

Fig. 3.4  Normal 5-month postnatal lung. This bronchovascular bundle contains both a terminal bronchiole and an accompanying arteriole. These structures are invested in a fibroconnective tissue sheath that also contains lymphatics but not venules (H&E, 10×)

Fig. 3.5  Normal 5-month postnatal lung. A pulmonary acinus consists of a respiratory bronchiole (rb) with associated alveolar ducts (ad) and alveoli (alv) (H&E, 10×)

Fig. 3.6  Normal 5-month postnatal lung. At high power, the transition from ciliated columnar bronchiolar epithelium to flattened alveolar epithelium can be seen in this pulmonary acinus (H&E, 20×)

gressively smaller proceeding distally; bronchi are also associated with seromucinous glands. In contrast, bronchioles (Fig.  3.11) contain neither a cartilage support structure nor associated seromucinous glands. However, both bronchi and bronchioles are supported by smooth muscle and connective tissue. The epithelial lining of bronchi is pseudostratified ciliated respiratory in type. Bronchioles show similar ciliated respiratory-­type epithelium, but this epithelium is simple rather than pseudostratified [6]. The respiratory epithelium of airways incorporates occasional goblet cells, which are more prevalent proximally than distally, and club cells (Clara cells/bronchiolar exocrine cells), which are present primarily within distal bronchioles. Neuroendocrine cells are also present (though normally inconspicuous) along both airways and airspaces.

Infolding of airway-lining epithelium is seen within larger bronchi and bronchioles and is likely related to the contraction of airway smooth muscle [2, 6]. Aggregates of lymphoid tissue, known as bronchus-associated lymphoid tissue (BALT), may be seen postnatally along the airways of normal infants and children [6]. Alveoli have a polygonal shape and are lined predominantly by simple, flattened epithelium, known as type I pneumocytes (Figs. 3.12 and 3.13). Type I pneumocytes provide a thin air-blood interface with the abundant capillary network of the intervening alveolar septa. Alveoli also contain interspersed type II pneumocytes, which are more cuboidal in shape. Type II pneumocytes produce surfactants and ­represent the progenitors of type I pneumocytes [1, 6]. They may become prominent in response to injury.

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Fig. 3.7  Normal 5-month postnatal lung. Interlobular septa demarcate pulmonary lobules and consist of bands of connective tissue, which intersect peripherally with the visceral pleura (H&E, 4×)

Fig. 3.8  Normal 5-month postnatal lung. Interlobular septa may contain occasional venous or lymphatic channels (H&E, 10×)

Fig. 3.9  Normal 5-month postnatal lung. Bronchi are lined by respiratory mucosa and contain subepithelial connective tissue incorporating cartilage and smooth muscle, which is more abundant proximally than distally (H&E, 20×)

Fig. 3.10  Normal 5-month postnatal lung. Bronchial mucosa consists of pseudostratified, ciliated columnar epithelium (H&E, 40×)

Fig. 3.11  Normal 5-month postnatal lung. Bronchioles are lined by mostly simple, ciliated columnar epithelium. No cartilage is present (H&E, 40×)

Fig. 3.12  Normal 5-month postnatal lung. Mature alveoli consist of polyhedral airspaces separated from one another by thin alveolar septa, which contain an abundant capillary network and occasional small, thin-walled intra-acinar arterioles (H&E, 20×)

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Fig. 3.13  Normal 5-month postnatal lung. Alveoli are lined by predominantly flattened type I pneumocytes (H&E, 40×)

Fig. 3.14  Embryonic stage: normal lung at 7  weeks of gestation. The embryonic stage of lung development is characterized by small, primitive tubules set amidst an abundant loose mesenchymal stroma (H&E, 10×)

Fig. 3.15  Embryonic stage: normal lung at 7  weeks of gestation. Mesenchyme shows some condensation around endodermal tubules (H&E, 20×)

Fig. 3.16  Embryonic stage: normal lung at 7 weeks of gestation. Tubules are lined by thick, pseudostratified endodermal epithelium (H&E, 40×)

Stages of Pulmonary Developmental Embryonic Stage During the earliest stage of lung development, the embryonic stage, the lung consists of rather primitive bronchial buds. These bronchial buds consist of small, simple tubules, which are lined by a thick layer of endodermally derived epithelium and are situated within a bed of abundant loose mesenchyme (Figs. 3.14, 3.15, and 3.16) [1]. Pseudoglandular Stage During the pseudoglandular stage, progressive branching of bronchial buds gives rise to increasing generations of airways. Early in the pseudoglandular stage (Figs.  3.17, 3.18, and 3.19), these airways remain small, simple tubules lined by pseudostratified endoderm, but by approximately 13 weeks

of gestation, these airways enlarge with progressive branching, and the endodermal lining thins. The more proximal airways become lined by ciliated respiratory epithelium with interspersed goblet cells. The more distal airways become lined by simple columnar epithelium. This epithelium is notable for prominent subnuclear vacuolization, conferring a “pseudoglandular” appearance to these simple tubules (Figs. 3.20, 3.21, and 3.22). Similar to the embryonic period, these tubules are set amid a relatively abundant loose interstitial mesenchyme, which appears somewhat condensed around bronchial buds [6, 8]. Cartilage formation around proximal airways and smooth muscle around airways and major vessels is also first appreciable during this stage [1].

Canalicular Stage The canalicular stage is marked by the initiation of airspace development and the beginnings of capillarization of

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Fig. 3.17  Early pseudoglandular stage: lung at 10–11 weeks of gestation. Early in the pseudoglandular stage, the lung consists of simple tubules, which are increased in number compared with the embryonic stage (H&E, 10×)

Fig. 3.18  Early pseudoglandular stage: lung at 10–11 weeks of gestation. These tubules remain situated amidst a relatively abundant, loose mesenchymal stroma, similar to the embryonic stage (H&E, 20×)

Fig. 3.19  Early pseudoglandular stage: lung at 10–11 weeks of gestation. Tubules are lined by pseudostratified to simple columnar endodermal epithelium (H&E, 40×)

Fig. 3.20  Late pseudoglandular stage: lung at 15 weeks of gestation. Late in the pseudoglandular stage, the lung remains composed of simple tubules amidst a loose mesenchymal stroma (H&E, 10×)

these primitive airspaces. The onset of the canalicular stage is first recognizable as capillaries becoming apposed to and even extending between airspace epithelial cells, whereby the airspaces are “canalized.” Early in the canalicular stage (Figs. 3.23, 3.24, and 3.25), the lining of the simple tubules transitions from columnar, vacuolated epithelium to cuboidal, nonvacuolated epithelium. A rather abundant mesenchyme remains. However, as the canalicular stage proceeds (Figs. 3.26, 3.27, and 3.28), these sim-

ple tubular structures are increasingly enlarged and complex. The outlines of these early airspaces change from tubular to rather wavy or undulating. The lining epithelium becomes progressively simple cuboidal in type, and a subset of epithelium may even become flattened as early as 20–22 weeks of gestation. This epithelial transition represents the initiation of mature pneumocyte differentiation. Ultimately, type II pneumocytes will remain cuboidal, and type I pneumocytes will be flattened. In

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Fig. 3.21  Late pseudoglandular stage: lung at 15 weeks of gestation. Tubules are lined by simple columnar epithelium (H&E, 20×)

Fig. 3.22  Late pseudoglandular stage: lung at 15 weeks of gestation. Epithelium is notable for prominent subnuclear vacuoles (H&E, 40×)

Fig. 3.23  Early canalicular stage: lung at 18 weeks of gestation. Early in the canalicular stage, the lung is characterized by increasingly branched tubules (H&E, 10×)

Fig. 3.24  Early canalicular stage: lung at 18 weeks of gestation. Tubules are increasingly branched, but abundant mesenchyme remains (H&E, 20×)

addition, the amount of interstitial mesenchyme begins to decrease. The beginning of pneumocyte differentiation, in combination with the canalization and decreasing interstitial mesenchyme of this stage, results in a primitive aircapillary membrane, also as early as 20–22  weeks of gestation. Airways show condensation of surrounding elastic tissue, and by the end of this stage, the most distal airways show differentiation of lining epithelium to form ciliated respiratory epithelium and club cells [1, 6, 8].

Fig. 3.25  Early canalicular stage: lung at 18 weeks of gestation. The lining epithelium is transitioning from columnar and vacuolated to cuboidal and nonvacuolated. Capillaries are just beginning to protrude between these epithelial cells (H&E, 40×)

Saccular Stage The saccular stage is heralded by the formation of secondary crests. These ridges of epithelial-surfaced mesenchyme contain a double-layered capillary network and protrude into airspaces, subdividing them into increasingly more complex structures known as saccules. With the ingrowth of secondary crests and expansion of the capillary network, the saccules become vascularized, initially with two layers of

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P. A. Kreiger and J. E. Pogoriler

Fig. 3.26  Late canalicular stage: lung at 23 weeks of gestation. Late in the canalicular stage, the lung is characterized by more complex airspaces. The amount of interstitial mesenchyme is beginning to decrease, compared with earlier gestations. Distal airways are readily recognizable among airspaces (H&E, 10×)

Fig. 3.27  Late canalicular stage: lung at 23  weeks of gestation. Airspaces show slightly undulating outlines (H&E, 20×)

Fig. 3.28  Late canalicular stage: lung at 23  weeks of gestation. A primitive capillary network is beginning to emerge. Airspaces are lined by predominantly cuboidal epithelium, with flattened, nonvacuolated epithelium in some areas. Many capillaries are apposed to and protrude between airspace epithelial cells (H&E, 40×)

Fig. 3.29  Early saccular stage: lung at 28  weeks of gestation. The early saccular stage is characterized by more complex airspaces with decreasing amounts of interstitial mesenchyme and an expanding capillary network (H&E, 10×)

capillaries (Figs.  3.29, 3.30, and 3.31). The saccular stage is also marked by progressive thinning of the interstitial mesenchyme and an increasing framework of elastic tissue, not only around airways but also between airspaces. By the late saccular stage (Figs. 3.32, 3.33, and 3.34), the double-­ layered capillary network begins to coapt into a single capillary layer. The formation of a single-layered capillary network in concert with the differentiation of airspace epi-

thelium to flattened pneumocytes provides for the maturing ­capillary-­airspace interface necessary for eventual postnatal respiration [1, 6, 8].

Alveolar Stage During the alveolar stage, the amount of interstitial mesenchyme decreases substantially to form thin interalveolar septa. Alveoli now show their mature back-to-back configuration and are lined by flattened mature epithelium. Alveoli

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Fig. 3.30  Early saccular stage: lung at 28 weeks of gestation. Airspaces show a wavy outline and an expanding capillary network (H&E, 20×)

Fig. 3.31  Early saccular stage: lung at 28 weeks of gestation. Airspaces are now becoming subdivided by “secondary crests” (arrow), to form “saccules.” These saccules are increasingly lined by flattened epithelium. Secondary crests initially form a double capillary network within the interstitium (H&E, 40×)

Fig. 3.32  Late saccular stage: lung at 31 2/7 weeks of gestation. The late saccular stage is characterized by increasing numbers of saccules (H&E, 10×)

Fig. 3.33  Late saccular stage: lung at 31 2/7 weeks of gestation. The interstitial capillary network is expanding and is also transitioning from a double-layered network to a single layer of capillaries (H&E, 20×)

attain a more polyhedral configuration, compared with the rounded sacs of the saccular stage; this process is thought to be related to increasing elastin deposition within the interstitium of the lung (Figs.  3.35, 3.36, and 3.37). Capillaries appear as thin single layers within alveolar septa and directly coapt with airspace-lining epithelium [1, 6, 7]. By the completion of the alveolar stage of lung development, the lung has attained its full complement of structures, including lymphatics, vasculature, airways, and airspaces, and a fully mature alveolar-capillary surface for postnatal respiration has been achieved [1].

Special Considerations Pulmonary Arteries and Arterioles Pulmonary arteries and arterioles can be described by the relative thickness of the muscular media and by the airway level to which smooth muscle extends. In the healthy adult, pulmonary arteries and arterioles are relatively thin-walled, and muscular arterioles extend to the level of the alveoli and

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Fig. 3.34  Late saccular stage: lung at 31 2/7  weeks of gestation. Saccules are lined by flattened epithelium with an increasingly thin interstitium (H&E, 40×)

Fig. 3.35  Alveolar stage: lung at 36 3/7 weeks of gestation. The beginning of the alveolar stage is characterized by the attainment of thin interalveolar septa separating now back-to-back alveoli (H&E, 4×)

Fig. 3.36  Alveolar stage: lung at 36 3/7 weeks of gestation. Alveolar septa contain an abundant single-layered interalveolar capillary network (H&E, 10×)

Fig. 3.37  Alveolar stage: lung at 36 3/7 weeks of gestation. Alveoli are lined by flattened epithelium, and the abundant capillaries bulge into the alveoli (H&E, 40×)

contain a single external elastic lamina. In the fetus, the muscular wall is thicker as a fraction of the total arterial circumference, and the muscle layer in smaller arterioles is incomplete (at the level of the alveolar ducts) or absent (at the level of the saccules/alveoli) [9]. Thinning of the arterial walls occurs within the first weeks of life, and extension of a more complete muscular coat into the arterioles at the level of the alveolar duct and the alveoli is present by approximately 6  months of age (Figs.  3.38 and 3.39) [10]. In pathologic states either in utero or postnatally, alveolar (intra-acinar) arterioles may appear thick-walled from muscularization and may develop a partial internal elastic lamina. This may reflect an aberration of the normal remodeling process of the pulmonary arterial tree, but applying these findings to an individual case is often problematic without extensive quantification, as

the extent of muscularization is highly variable between patients, and the presence of a subset of muscularized arterioles is normal even in neonates (Fig. 3.40) [9].

Intrapulmonary Lymphatics Intrapulmonary lymphatics are distributed within the connective tissue along the bronchovascular bundles as far as the alveolar duct, along the interlobular septa, and in the pleura. They are lined by specialized endothelial cells without smooth muscle. Given this very thin wall, they may be relatively inapparent when collapsed. They can be highlighted by immunohistochemistry for D2-40, and, in contrast to arterioles and veins, they should be negative for smooth muscle actin. They

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Fig. 3.38 Pulmonary arterioles, normal 5-month postnatal lung. Normal intra-acinar arterioles are present within the alveolar parenchyma beyond the level of the bronchovascular bundle (H&E, 10×)

Fig. 3.39  Pulmonary arterioles, normal 5-month postnatal lung. Intra-­ acinar arterioles normally appear to be progressively thin-walled as they extend through the parenchyma toward the pleural surface (H&E, 20×)

Fig. 3.40  Immunohistochemistry for smooth muscle actin in this 1-day-old infant shows incomplete muscularization of an arteriole (asterisk) (smooth muscle actin immunohistochemistry, 60×)

Fig. 3.41  Intrapulmonary lymphatics: Lymphatics (asterisk) in this lung from an infant at 26 weeks of gestation are seen in the bronchovascular bundle and in the pleura (H&E, 10×)

are present in a similar distribution throughout development at the earliest stages at which they have been specifically evaluated in humans (Fig. 3.41) [11]. Lymphatics may appear more or less prominent depending on the degree of distension, which can be artifactually induced by perfusion.

monia [14]. In contrast, BALT is often seen in healthy infants and young children, increasing in frequency in the first several years of life [15]. It is not a normal constituent of healthy adult human lungs, although it can be seen transiently following exposure to new antigens (Figs. 3.42 and 3.43).

Intrapulmonary Lymphocytes

Pulmonary Neuroendocrine Cells

Bronchus-associated lymphoid tissue (BALT) consists of peribronchial lymphoid aggregates with follicular B cells, parafollicular B and T cells, and scattered intraepithelial T cells. In the absence of in utero infection, BALT is generally absent in fetuses [12, 13], but either BALT or interstitial collections of mononuclear cells can be seen in congenital pneu-

Single pulmonary neuroendocrine cells and tight clusters (neuroendocrine bodies) are present along the epithelium, most prominently at airway bifurcations and in respiratory bronchioles at the junction of airway and alveolar epithelium (Fig. 3.44) [16]. They are recognizable from approximately 8 to 10 weeks of gestation, but they do not become numerous

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Fig. 3.42  Normal 5-month postnatal lung. Bronchus-associated lymphoid tissue (BALT) may be seen as small aggregates of lymphocytes along the airways in the normal postnatal lung, but it is not typically prominent in the normal fetal lung (H&E, 10×)

P. A. Kreiger and J. E. Pogoriler

Fig. 3.43  Bronchus-associated lymphoid tissue (BALT). In this lung from a 12-month-old infant, a well-developed germinal center is present adjacent to the bronchiole, and lymphocytes infiltrate into the overlying epithelium (H&E, 10×)

Intrapulmonary Karyorrhectic Cells The normal fetal lung, particularly in the midtrimester, may contain prominent karyorrhectic material. This material may be the result of apoptosis, which is likely integral to the normal development of the lung [18, 19]. Karyorrhectic material within fetal airways and airspaces may mimic neutrophils, an important distinction (Fig. 3.45).

Intrapulmonary Squamous Cells

Fig. 3.44  Pulmonary neuroendocrine cells. Immunohistochemistry for bombesin in this lung from a 2-month-old shows a respiratory bronchiole with a few positive cells at the junction between the airway and alveolar epithelium. Other markers that are more often clinically available (such as chromogranin and synaptophysin) stain fewer cells (bombesin immunohistochemistry, 20×)

until late in the second trimester [11]. Their numbers remain higher in fetal and neonatal life than in healthy adult lungs. Perinatally, their numbers are decreased in patients with acute hyaline membrane disease and are increased in chronic lung disease of prematurity, consistent with the fact that hyperplasia of neuroendocrine bodies occurs in hypoxia [11, 17]. Immunohistochemistry for common neuroendocrine markers such as chromogranin and synaptophysin may identify fewer cells than bombesin.

A few scattered squamous cells within the fetal airways and airspaces of later gestations are a normal finding. These squamous cells enter the lung via fetal inspiration of amnionic fluid. Large numbers of squamous cells in the fetal lung are considered to be pathologic, however, and a sign of fetal hypoxia (Fig. 3.46) [6].

 valuation of Alveolar Development E in the Perinatal Period One frequently proposed measure of lung alveolar development is the radial alveolar count (RAC) described by Emery and Mithal [20]. In this method, a line is drawn from respiratory bronchioles to the nearest pleura or interlobar septum, counting the number of alveolar spaces that are crossed (Fig. 3.47). In published reference ranges, full-term newborns have an average of 4–6 alveoli, increasing to 8–12 by 1 year of age. The degree of inflation may significantly alter counts,

3 Lung

a

Fig. 3.45  Karyorrhectic material: lung at 16 weeks (a) and 14 weeks (b) gestation. Karyorrhectic material (a) is present within the lung of a midtrimester fetus. This material should not be mistaken for neutro-

47

b

phils, which have better-defined nuclear lobes, as seen in (b), a case of amniotic fluid infection (H&E, 60×)

a

b

Fig. 3.46  Squamous cells. Lung at 41 weeks of gestation. Occasional squamous cells are present within this late-gestation lung (a), a finding generally considered to be within normal limits. However, large numbers of squames, as seen in this late-gestation lung (b), are considered to be pathologic (H&E, a 40×; b 20×)

Fig. 3.47  Radial alveolar count. In this normal 1-day-old lung, a line is drawn from the center of the respiratory bronchiole to the pleura. The number of alveolar spaces that the line crosses through is counted (H&E, 10×)

however, particularly postnatally, so a large numbers of counts should be performed. Moreover, marked i­nterobserver variability has been noted [21, 22]. Therefore, if an RAC is performed, the results should be considered definitively abnormal only if they are markedly outside of the expected range. In a typical postmortem case, identification of respiratory bronchioles may be challenging because of detachment of the bronchiolar epithelium and extension of smooth muscle into the alveolar walls of ventilated infants. In general practice, therefore, evaluation is usually more subjective. A more widely applicable rule would suggest that bronchioles should not extend all the way to the pleura, but should be several airspaces away. Other measures of pulmonary hypoplasia include lung weight, lung volume, and lung–body weight ratio.

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References 1. dePaepe M. Lung growth and development. In: Churg AM, Myers JL, Tazelaar HD, Wright JL, editors. Thurlbeck’s pathology of the lung. 3rd ed. New York: Thieme; 2005. p. 39–71. 2. Colby TV, Leslie KO, Yousem SA.  Lungs. In: Mills S, editor. Histology for pathologists. 3rd ed. Philadelphia: Lippincott Williams & Wilkins; 2007. 3. Langston C.  Prenatal lung growth and pulmonary hypoplasia. In: Stocker JT, editor. Pediatric pulmonary disease. New  York: Hemisphere; 1989. p. 1–13. 4. Kimura J, Deutsch GH.  Key mechanisms of early lung development. Pediatr Dev Pathol. 2007;10:335–47. 5. O’Rahilly R, Müller F. Human embryology and teratology. 3rd ed. New York: Wiley-Liss; 2001. 6. Valdés-Dapena MA. The lower respiratory tract. Histology of the fetus and newborn. Philadelphia: WB Saunders; 1979. p. 309–41. 7. Hislop A.  Developmental biology of the pulmonary circulation. Paediatr Respir Rev. 2005;6:35–43. 8. Wigglesworth J. Perinatal pathology. Philadelphia: WB Saunders; 1996. 9. Haworth SG, Hislop AA.  Pulmonary vascular development: normal values of peripheral vascular structure. Am J Cardiol. 1983;52:578–83. 10. Hislop A, Reid L.  Pulmonary arterial development during childhood: branching pattern and structure. Thorax. 1973;28:129–35. 11. McNellis EM, Mabry SM, Taboada E, Ekekezie II. Altered pulmonary lymphatic development in infants with chronic lung disease. Biomed Res Int. 2014;2014:109891. 12. Gould SJ, Isaacson PG.  Bronchus-associated lymphoid tissue (BALT) in human fetal and infant lung. J Pathol. 1993;169:229–34.

P. A. Kreiger and J. E. Pogoriler 13. Ersch J, Tschernig T, Stallmach T. Frequency and potential cause of bronchus-associated lymphoid tissue in fetal lungs. Pediatr Allergy Immunol. 2005;16:295–8. 14. Barter R. The histopathology of congenital pneumonia: a clinical and experimental study. J Pathol Bacteriol. 1953;66:407–15. 15. Tschernig T, Kleemann WJ, Pabst R.  Bronchus-associated lymphoid tissue (BALT) in the lungs of children who had died from sudden infant death syndrome and other causes. Thorax. 1995;50: 658–60. 16. Watanabe H. Pathological studies of neuroendocrine cells in human embryonic and fetal lung. Light microscopical, immunohistochemical and electron microscopical approaches. Acta Pathol Jpn. 1988;38:59–74. 17. Johnson DE, Kulik TJ, Lock JE, Elde RP, Thompson TR. Bombesin-, calcitonin-, and serotonin-immunoreactive pulmonary neuroendocrine cells in acute and chronic neonatal lung disease. Pediatr Pulmonol. 1985;1:S13–20. 18. Kresch MJ, Christian C, Wu F, Hussain N. Ontogeny of apoptosis during lung development. Pediatr Res. 1998;43:426–31. 19. Scavo LM, Ertsey R, Chapin CJ, Allen L, Kitterman JA. Apoptosis in the development of rat and human fetal lungs. Am J Respir Cell Mol Biol. 1998;18:21–31. 20. Emery JL, Mithal A. The number of alveoli in the terminal respiratory unit of man during late intrauterine life and childhood. Arch Dis Child. 1960;35:544–7. 21. Cooney TP, Thurlbeck WM. The radial alveolar count method of Emery and Mithal: a reappraisal. 2 – intrauterine and early postnatal lung growth. Thorax. 1982;37:580–3. 22. Cooney TP, Thurlbeck WM. The radial alveolar count method of Emery and Mithal: a reappraisal. 1 – postnatal lung growth. Thorax. 1982;37:572–9.

Part III Digestive System

4

Gastrointestinal Tract Dale S. Huff and Chrystalle Katte Carreon

Introduction The gastrointestinal tract is a tube that extends from the end of the pharynx to the anus. Its functions are the propulsion and digestion of food and the elimination of waste products. It is divided into primary segments that are, from rostral to caudal, the esophagus, stomach, small intestine, and large intestine. The primary segments are divided into secondary segments (from rostral to caudal): the proximal, middle, and distal parts of the esophagus; the cardia, fundus, body, and antrum or pylorus of the stomach; the duodenum (with its first, second, third, and fourth parts), jejunum and ileum of the small intestine; and the cecum, appendix, ascending, transverse, descending, sigmoid, rectum, and anal canal segments of the large intestine. The histologic junctions between the primary segments may be abrupt, but those between the secondary segments are gradual transition zones. Sphincters and valves at junctions of primary segments regulate forward propulsion and prevent retrograde flow of the contents. Although the general architecture of the gastrointestinal tract is established in embryonic life, the layers of the wall continue to develop during fetal life. Thus, the histology of the gastrointestinal tract is characterized by numerous changes in the fetal period, which are highlighted in this chapter.

Embryology The gastrointestinal tract is composed of an epithelial lining derived from the endoderm. The epithelial tube is surrounded by concentric layers of mesenchyme derived from the

D. S. Huff ∙ C. K. Carreon (*) Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]; [email protected]

splanchnic mesoderm. The development of the tube is controlled by rostral-to-caudal, right-to-left, and inside-to-­ outside (radial) molecular genetic morphogenetic gradients. The rostral-to-caudal division of the tube into its primary and secondary segments is completed by the end of the seventh postfertilization week. The right-to-left orientation is established by the tenth postfertilization week, when rotation and fixation of the stomach and intestines are completed [1–3]. There is one exception: The “descent” of the cecum, the vertical orientation of the ascending colon, and the formation of the hepatic flexure are not completed until the third trimester. The radial gradient is controlled by epithelial-mesenchymal interactions and results in the orderly development of the concentric layers of the tube and in the radial organization of the villi and crypts; it is completed by the tenth week. The histogenesis of these layers is described below.

Histology General Considerations  rganization of the Wall of the Gastrointestinal O Tract The wall of the gastrointestinal tract is organized into five concentric layers. From inside out, these are the mucosa, muscularis mucosae, submucosa, muscularis propria, and subserosa. The histology of the mucosa of each segment is unique to that segment and is specific for the function of that segment. The muscularis mucosae, submucosa, muscularis propria, and subserosa or adventitia vary little from segment to segment. The mucosa is composed of the epithelium and the lamina propria. The epithelium lies on a basement membrane. In the esophagus, it is nonkeratinizing, stratified squamous epithelium. In the stomach, it is simple, columnar, mucus-­producing surface epithelium with gastric pits, which give rise to simple mucous cardiac and pyloric glands, as well as serous

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_4

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g­ astric glands or crypts. In the small intestine, it is villous, with serous crypts between the villi. In the large intestine, it is simple, columnar, mucus-producing surface epithelium with mucous crypts. Numerous neuroendocrine cells are scattered throughout the epithelium of the gastrointestinal tract (primarily in the base of the crypts) and are seen beginning at the 11th week [4]. At least 16 subtypes have been identified, each of which produces a different gastrointestinal peptide hormone [5]. The frequency and topographic distribution of neuroendocrine cells have been reviewed [6]. Although some can be identified in routine sections by the presence of subnuclear eosinophilic granules or, in the case of Paneth cells, by supranuclear eosinophilic granules, they are difficult to see in routine sections and are best studied using immunohistochemical stains. The lamina propria is composed of loose stroma containing nerves, blood vessels, lymphatics, and the scattered lymphoid and other inflammatory cells of the diffuse gut-associated lymphoid tissue. Eosinophils and mast cells are seen in the small and large intestines beginning by the 16th–20th weeks. The number and cell types of the diffuse gut-associated lymphoid tissue vary somewhat, but otherwise, the lamina propria varies little from segment to segment. The muscularis mucosae is a thin double layer of smooth muscle that separates the mucosa from the submucosa and is present throughout the gastrointestinal tract from the esophagus to rectum. It consists of an inner circular and outer longitudinal layer in all segments of the gastrointestinal tract—except in the esophagus, where it consists of bundles of longitudinal muscle fiber, making it more conspicuous in this area. In the stomach and intestines, usually only the inner circular layer is visible in H&E-stained sections; the outer longitudinal layer is usually attenuated and is hardly visible except in the setting of intestinal obstruction, where it becomes more prominent proximal to the obstructed segment. The muscularis mucosae is composed of smooth muscle throughout the gastrointestinal tract, even in the upper one third of the esophagus, where the muscularis propria consists of striated muscle. Wisps of smooth muscle extend from the muscularis mucosae into the lamina propria and attach to the basement membrane. Small nerve fibers are present. The submucosa is composed of loose connective tissue containing nerves, ganglia, blood vessels, scattered smooth muscle fibers, lymphatics, and the solitary and aggregated lymphoid follicles of the gut-associated lymphoid tissue, which are most well-developed at the junctions of the primary segments, such as the duodenum, terminal ileum, appendix, and anorectal region. The submucosal neural plexuses are divided into the submucosal plexus of Meissner, located immediately below the muscularis mucosae, and the deep submucosal plexus of Henle, located along the inner surface of the inner circular layer of the muscularis propria.

D. S. Huff and C. K. Carreon

The submucosa of the esophagus and duodenum contain specialized glands: the esophageal submucosal mucous glands and Brunner’s glands, respectively. The muscularis propria consists of two layers of muscle, the inner circular layer and the outer longitudinal layer. These layers are separated by a thin, stromal, intermuscular septum containing the myenteric or Auerbach’s plexus of nerves and ganglion cells. The muscularis propria is composed of smooth muscle except in the upper one third of the esophagus, where it is skeletal muscle. In the proximal transitional region of the esophagus, both types of muscle may be observed. Recent studies have described a strong and diffuse expression of smoothelin in both layers of the muscularis propria, whereas the muscularis mucosae typically demonstrates absent to patchy, weak staining by immunohistochemistry. This difference between the two muscular layers in expression and immunohistochemical staining pattern of smoothelin allows for possible differentiation of the two layers, especially in suboptimally sampled and/or poorly oriented specimens [7, 8]. The subserosa or adventitia is loose areolar tissue containing nerves, blood vessels, lymphatics, and lymph nodes. In segments that are intraperitoneal, it is continuous with the mesentery and is covered by the mesothelium. The esophagus, duodenum (except for its first part), rectum, and anus are retroperitoneal and have no mesothelial covering. The dorsal walls of the ascending and descending colon also are retroperitoneal, so only the ventral walls are covered by the mesothelium.

 iming and Sequence of Developmental Events T The gestational ages at which developmental events occur vary considerably as cited by different authors. The gestational ages given below are averages of different ages given for each event by several authors [1–4, 9–14] and should be considered approximations. It is convenient to divide the development of the gastrointestinal tract into three arbitrary periods. During the first period, up to the 14th postfertilization week, the unique definitive histology of each segment is established. During the second period, between 14 and 30  weeks, a few additional histological structures appear. The third period, from 30 to 40 weeks, is one of growth and continued maturation of structures and cells that developed in the previous periods. Histogenesis in general proceeds from rostral to caudal. Less than 14 weeks  The differentiation of the mucosa occurs in an orderly sequence that is similar in all segments. First, the simple endodermal epithelium, which is present since the formation of the bilaminar disc, develops into a stratified, columnar primordial epithelium before the sixth postfertilization week. This primitive stratified epithelium is similar throughout the gastrointestinal tract. Second, during the

4  Gastrointestinal Tract

sixth week, the epithelium enters into a proliferative phase, also referred to as vacuolization, in which the epithelium of the entire tract becomes multilayered and thick. Third, during the eighth week, the thick proliferative epithelium differentiates into tall, simple, or pseudostratified columnar epithelium, which then immediately differentiates by 8–10  weeks into the epithelium specific for each segment. The development of the unique mucosa of each segment is described below. The development of the muscular coats and the enteric neural system are intertwined. The muscularis propria appears first in the esophagus, beginning with the appearance of the inner circular layer during the sixth week, followed by the outer longitudinal layer during the seventh week. Differentiation spreads from rostral to caudal and reaches the anal canal by as early as the eighth week. The outer longitudinal layer is very thin until approximately the third trimester. The muscularis mucosae is the last layer to develop. It begins in the esophagus during the 11th week, in the stomach during the 14th week, in the anal canal in the 16th week, and in the duodenum in the 21st week. It spreads from the duodenum caudally and from the anal canal rostrally to meet at the ileocecal junction during the last trimester. However, recent studies utilizing immunohistochemical stains suggest that the onset of differentiation of the muscularis propria and muscularis mucosae in the upper esophagus occur earlier in gestation and the rostral-to-caudal wave of differentiation of the smooth muscle coats to the rectum is more rapid than the times indicated above, which are based on routine stains [15, 16]. The rostral neural crest cells, which will form the nerves and ganglion cells of the neurenteric plexuses, diffusely invade the upper esophagus during the fourth week and migrate caudally to reach the anal canal by the seventh week. The neural crest cells gather into loose bands in the connective tissue layer of the future muscularis propria to form the primordium of Auerbach’s plexus in the esophagus during the fifth week. During the sixth week, the neural crest cells migrate centripetally through the developing inner circular muscle layer into the submucosa to form the primordia of the submucosal plexuses. During the seventh week, the bands of neural primordia in the esophagus form discrete nests or ganglia composed of neuroblasts. In the ninth week, interstitial cells of Cajal enter the rostral esophagus and gather around Auerbach’s plexuses, then spread centripetally into the submucosa, and surround the submucosal plexuses of Henle and Meissner. CD117 is a useful immunohistochemical marker for identification of interstitial cells of Cajal [17, 18], and its expression, particularly in the fetal small intestine, has been demonstrated by immunohistochemistry as early as 13 weeks of gestation [17]. All these events begin first in the muscularis propria of the upper esophagus, spread into submucosa a week or two later, and spread caudally so that the entire

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enteric neural system is completed by approximately 14  weeks [15, 16]. Neural crest cells from the sacral area invade the anorectal area during the 10th–12th weeks. In the meantime, the neuroblasts of the plexuses gradually differentiate into mature ganglion cells from rostral to caudal, a process that is not complete until the neonatal period. Beginning at 11  weeks, neuroendocrine cells can be seen throughout the gastrointestinal tract by careful examination and can be particularly conspicuous in the appendix. These cells are thought to help maintain homeostasis by secreting various regulatory peptides [19]. In the appendix, the subepithelial neuroendocrine cells have been postulated as the precursor of neuroendocrine tumors (formerly “carcinoid tumor”), the most common neoplasm in this site [20]. 14–30 weeks  Scattered lymphoid cells appear in the lamina propria during the seventh week. Beginning in the 11th week, immunohistochemical stains identify precursors of lymphoid tissue in sites where lymphoid aggregates will develop in the future. After 14  weeks, the gut-associated lymphoid tissue is visible in routine histological sections. Tiny aggregates of lymphoid and inflammatory cells in the lamina propria of the terminal ileum are infrequent at 14  weeks and more consistently seen at 16  weeks. By 19–20 weeks, more organized primary follicles are present, and these become easily recognizable in the terminal ileum, appendix, duodenum, anorectal area, and other sites after 20  weeks postmenstrual age. By 24  weeks, large Peyer’s patches are present in the submucosa of the terminal ileum, and prominent submucosal solitary primary follicles are seen in the duodenum, jejunum, appendix, and distal colon [21]. The development of primary follicles and the diffuse lymphoid tissue in the lamina propria and submucosa during this period is antigen-independent. Germinal centers and mature plasma cells, the development of which are dependent upon exposure to foreign antigens, are not found in normal fetuses or in neonates prior to 2 weeks of age. The presence of easily identifiable submucosal lymphoid aggregates and primary follicles prior to 20 weeks, or the presence of germinal centers and mature plasma cells at any time during fetal life, may indicate fetal infection or other abnormal antigenic stimulus. 30–40 weeks  From 30  weeks to term, the gastrointestinal tract continues to grow and mature. Some of the major changes include maturation of the esophageal epithelium, development of the muscularis mucosae of the small and large intestines, and continued maturation of the ganglion cells of the plexuses of Auerbach, Henle, and Meissner. Maturation of some of the histologic components of the gastrointestinal tract continues into the neonatal period and infancy; this is especially true of the gut-associated lymphoid tissue and the ganglion cells of the enteric neural system.

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D. S. Huff and C. K. Carreon

Esophagus Less Than 14 Weeks During the seventh week, the primitive esophageal mucosa proliferates and forms multiple irregular bridges that traverse and partially occlude the lumen, leaving multiple small, round lumens called vacuoles. At this time, the inner circular layer of the muscularis propria has formed, the outer longitudinal layer is beginning to appear, and primitive Auerbach’s plexuses populated by neuroblasts are present. The muscularis mucosae and submucosal plexuses have not yet appeared (Fig. 4.1). The proliferative phase recedes; the epithelium differentiates into a primitive, pseudostratified columnar type (Figs.  4.2, 4.3, and 4.4); and the muscularis mucosae appears (see Fig.  4.4). The epithelium then differentiates into poorly organized, stratified squamous epithelium. Columnar ciliated cells cover the surface of the middle of the esophagus and spread to cover the entire surface by the 11th week (Figs.  4.5 and 4.6). The muscularis mucosae becomes thicker, and the outer longitudinal layer of the muscularis propria remains very thin. Some of the neuroblasts in submucosal and myenteric plexuses begin to mature toward ganglion cells (see Fig. 4.5).

Fig. 4.1  Esophagus at 7  weeks postfertilization. The proliferative phase of esophageal mucosal development is shown. The lumen is nearly occluded by irregular bridges of proliferative epithelium (EP), leaving several round lumens. This appearance has been referred to as vacuolization. The submucosa (SM) is prominent, but the muscularis mucosae and submucosal neural plexuses of Meissner and Henle are not present. The inner circular (IC) layer of the muscularis propria is well formed. The outer longitudinal layer (arrow) is discontinuous and one or two cells deep. Auerbach’s plexus (AP) is composed of a nearly continuous strip of undifferentiated neuroblasts (hematoxylin and eosin [H&E], 10×)

14–30 Weeks Beginning at approximately 16  weeks, squamous epithelium gradually replaces the ciliated columnar epithelium on the surface of the mid esophagus (Figs.  4.7 and 4.8). Esophageal cardiac glands, which are similar to gastric cardiac or pyloric glands, are mucus-producing glands in the lamina propria that are connected to the lumen by a duct that passes through the epithelium. Papillae of the lamina propria, which are finger-like processes of lamina propria that penetrate up into the epithelium during postnatal life and are important in the evaluation of esophagitis, are not present during fetal life (see Fig. 4.7). The esophageal muscularis mucosae rapidly increases in thickness and becomes the thickest of all the segments of the gastrointestinal tract. The primordia of esophageal submucosal mucous glands appear at 27 weeks and are thought to be analogous to the submucosal glands of the pharynx. The outer longitudinal layer of the muscularis propria becomes slightly thicker, and the deep submucosal plexuses of Henle become more numerous and conspicuous (see Figs.  4.7 and 4.9). Ectopic gastric mucosa of the cardiac, oxyntic, or pyloric variety is often

Fig. 4.2  Esophagus at 11 weeks postfertilization. The receding proliferative phase is shown. The lumen is re-established, and the epithelium is heaped up and multilayered in some areas and tall pseudostratified in others. The inner circular layer is conspicuous, but the outer longitudinal layer is discontinuous and one or two cells deep. Auerbach’s plexus is composed of neuroblasts (H&E, 10×)

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Fig. 4.3  Esophagus at 11  weeks postfertilization. This figure is a higher magnification of Fig.  4.2. The tall, pseudostratified columnar epithelium (EP) is undifferentiated. The submucosa (SM) is present, and the muscularis mucosae is just starting to develop. The submucosal plexuses are not yet visible. The inner circular layer (IC), outer longitudinal layer (arrow), and Auerbach’s plexus (outlined by asterisks) are similar to those in Fig. 4.2 (H&E, 20×)

Fig. 4.4  Esophagus at 11  weeks postfertilization. Undifferentiated, pseudostratified columnar epithelium is shown. The epithelium (EP) is no longer proliferative. The muscularis mucosae (MM) is better developed than in Fig. 4.3 despite the similar gestational age, and it separates the lamina propria (LP) from the submucosa (SM) (H&E, 20×)

Fig. 4.5  Esophagus at 12 weeks postfertilization. The primitive stratified squamous epithelium (EP) with ciliated columnar epithelium on the surface is shown. The epithelium is specific for the esophagus. The thickening of the muscularis mucosae (MM) and the inner circular layer (IC) continues to increase. The outer longitudinal layer (arrow) is still only one or two cells deep. Some of the neuroblasts of Auerbach’s plexus (AP) have begun to differentiate into ganglion cells (H&E, 20×)

Fig. 4.6  Esophagus at 12 weeks postfertilization. This higher magnification of Fig. 4.5 shows how the stratified squamous epithelium is disorganized, and the surface cells are ciliated (H&E, 60×)

seen in the mucosa of the upper esophagus throughout fetal life (Fig. 4.10) and may be confused with esophageal cardiac glands. Upward of 20 weeks of gestation, the lining epithelium is almost complete, and the muscularis mucosae and propria are readily identified. In addition, interstitial

cells of Cajal, which may be detected as early as 13 weeks, can be highlighted using CD117 immunohistochemistry (Figs. 4.11 and 4.12).

30–40 Weeks Figures 4.13, 4.14, 4.15, 4.16, and 4.17 illustrate the histology of the distal and proximal esophagus near term. The squamous epithelium spreads rostrally and caudally to replace

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D. S. Huff and C. K. Carreon

Fig. 4.7  Esophagus at 19  weeks postfertilization. Squamous cells are beginning to replace the ciliated columnar surface epithelium. The deep surface of esophageal epithelium is flat, and papillae of the lamina propria are not present. The deep submucosal plexus of Henle (asterisks) is present. The outer longitudinal layer of the muscularis propria (arrow) is only a few cells deep and is much thinner than the inner circular layer. Auerbach’s plexuses are discrete nests, as opposed to the nearly continuous ribbons that are characteristic of the plexus prior to 14 weeks (H&E, 4×)

Fig. 4.8  Esophagus at 19 weeks postfertilization. This higher magnification of Fig. 4.7 shows how squamous epithelium (arrows) begins to replace the ciliated columnar epithelium of the surface in focal areas (H&E, 10×)

Fig. 4.9  Esophagus at 19 weeks postfertilization. Partially differentiated submucosal and myenteric plexuses are shown. The deep submucosal plexus of Henle (asterisk) and the myenteric plexus of Auerbach (arrows) are composed of a mixture of undifferentiated and partially differentiated neuroblasts. IC inner circular layer of muscularis propria, IMS intermuscular septum, OL outer longitudinal layer of muscularis propria, SM submucosa (H&E, 20×)

Fig. 4.10  Ectopic gastric cardiac or pyloric mucosa in the esophagus at 19 weeks. The ectopic gastric mucosa or gastric heterotopia is composed of tall, mucus-producing simple columnar epithelium, which replaces the stratified squamous epithelium of the upper esophagus (H&E, 10×)

the ciliated cells and covers the surface of the entire esophagus by the 40th postmenstrual week (see Fig. 4.13). The submucosal mucous glands, the primordia of which appear by approximately 27 weeks, become well-­differentiated during this period. The ducts arise from the squamous mucosa and penetrate through the muscularis mucosae into the submucosa. The proximal ends of the ducts in the lamina propria are

lined by squamous cells. After they penetrate through the muscularis mucosae, the epithelium is composed of two layers of cuboidal epithelium, a luminal layer and a basal layer of myoepithelial cells (see Fig. 4.13). These ducts give rise to the mucous glands located in the submucosa (see Fig. 4.14). Some of the ganglion cells are fully mature, but partially differentiated neuroblasts remain (see Fig. 4.15). Some patches of ciliated epithelium may remain at birth (see Fig. 4.17), but those disappear within a few days after birth.

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Fig. 4.11  Esophagus at 21 weeks postfertilization. The lining epithelium is almost completely replaced by squamous epithelium. The muscularis mucosae and layers of the muscularis propria are also readily identifiable. Although quite inconspicuous on H&E stain, pale, elongated cells, the interstitial cells of Cajal, may be seen interspersed between the smooth muscle cells on careful examination (H&E, 10×)

Fig. 4.12  Esophagus at 21 weeks postfertilization. Immunohistochemical staining for CD117 highlights numerous interstitial cells of Cajal within the muscularis propria. These cells can be detected as early as 13 weeks of gestation by immunohistochemistry (CD117 Immunohistochemistry, 10×)

Fig. 4.13  Esophagus at 34 weeks postfertilization. Mature stratified squamous surface epithelium and ducts of esophageal submucosal mucous glands are present near the gastroesophageal junction. The epithelium is well-­ differentiated, stratified squamous epithelium without ciliated columnar cells on the surface. The duct of the submucosal mucous gland penetrates through the muscularis mucosae into the submucosa. The bilayer cuboidal ductal epithelium is composed of a luminal layer and a basal layer. The ducts are coursing caudally toward the gastroesophageal junction, which is out of the picture to the left (H&E, 10×)

Fig. 4.14  Esophagus at 34  weeks postfertilization. The submucosal glands of the ducts, shown in Fig. 4.13, are demonstrated. The esophageal epithelium and muscularis mucosae are in the upper left corner. The glands are immature and small (H&E, 10×)

Stomach Less Than 14 Weeks The unique histology of the stomach is established before the 14th postfertilization week. The multilayered, pseudostratified primordial gastric epithelium (Fig. 4.18) enters the pro-

liferative or vacuolization phase during the sixth to seventh postfertilization week. The proliferating epithelium partially occludes the lumen of the cardia and pylorus. The unique gastric epithelium, characterized by foveolar epithelium and gastric pits, replaces the proliferative epithelium in the body during the seventh to eighth week [22] and lines the entire stomach by the 10th to 12th week (Fig. 4.19). The first gastric glands with mucous neck cells, parietal cells, chief cells, and endocrine cells appear by the ninth to tenth week (Figs. 4.20 and 4.21). Cardiac mucosa becomes distinguishable from oxyntic mucosa, although the existence of a distinct cardiac

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Fig. 4.15  Auerbach’s plexus in the esophagus at 34 weeks postfertilization. The cells are a mixture of neuroblasts, partially differentiated neuroblasts, and mature ganglion cells. Compare with Figs. 4.3 and 4.9 (H&E, 60×)

D. S. Huff and C. K. Carreon

Fig. 4.16  Midesophagus at 35  weeks postfertilization. There are no papillae of the lamina propria. Ducts of esophageal submucosal mucous glands are seen in the lamina propria and submucosa, and the muscularis mucosae is very thick. The muscularis propria also is very thick, and although the outer longitudinal layer has grown considerably in thickness, it is still approximately half the thickness of the inner circular layer. The intermuscular septum is distinct (H&E, 2×)

Fig. 4.17  Esophagus at 35  weeks postfertilization. This figure is a higher magnification of the esophageal mucosa shown in Fig. 4.16. A few tufts of cilia remain on the surface (H&E, 20×)

mucosa has been questioned. The embryology of the mucosa of the esophagogastric junction is the subject of continuous debate [23, 24]. Specific cardiac and pyloric mucosae appear by week 13 (Figs. 4.22, 4.23, and 4.24), and the mucosa of the cardia and pylorus becomes villous. The gastric muscularis propria is composed of three layers: the inner circular and outer longitudinal layers, which are characteristic of the entire gastrointestinal tract, and an additional oblique layer internal to the inner circular layer (Fig. 4.25). The inner circular and oblique layers appear in the cardia during the seventh week, and the outer longitudinal layer appears during the eighth

Fig. 4.18  Stomach at 7 weeks postfertilization. The primordial multilayered pseudostratified epithelium is on the top. The inner circular layer (IC) of the muscularis propria is apparent, but the outer longitudinal layer has not yet differentiated. The serosa is at the bottom, and the gastric mesentery (M) is included (H&E, 20×)

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Fig. 4.19  Formation of gastric pits in stomach at 8 weeks postfertilization. The primordial epithelium has differentiated into distinctive gastric epithelium with the formation of gastric pits (asterisks) (H&E, 20×)

Fig. 4.20  Formation of gastric glands in stomach at 12 weeks postfertilization. The primitive gastric pits have differentiated into gastric glands with eosinophilic parietal cells, and the surface epithelium is differentiating into foveolar epithelium. The muscularis mucosae has not appeared. The muscularis propria is composed of the innermost oblique layer and the inner circular layer, which are nearly indistinguishable from each other, and the outer longitudinal layer, which is two or three cells thick. The nests of neuroblasts of Auerbach’s myenteric plexus are visible in the intermuscular septum (H&E, 10×)

Fig. 4.21  Stomach at 12 weeks postfertilization. This higher magnification is of the gastric glands in Fig. 4.20. The parietal cells and surface epithelium are distinct, and the muscularis mucosae is not present (H&E, 20×)

Fig. 4.22  Esophagogastric junction at 12 weeks postfertilization. The distal esophagus occupies the right half of the picture, and the cardiac mucosa is to the left. The squamous-columnar junction is indicated by the two asterisks. The arrows indicate the junction between the cardiac mucosa and the transition to oxyntic mucosa with gastric pits and parietal cells. The distance between the squamous-columnar junction and the transitional mucosa in this specimen is less than 1 mm. The thick muscularis mucosae (MMe) of the esophagus abruptly transitions into the much thinner muscularis mucosae of the stomach (MMs) (H&E, 4×)

week. All layers are present throughout the entire stomach by the ninth week, but the three layers may be indistinct. The circular layer is the most distinct of the three: it enlarges at the pylorus to form the pyloric sphincter during the ninth postfertilization week. The submucosal and myenteric plexuses, as well as the interstitial cells of Cajal, are formed at the same time as the m ­ uscularis propria, but the neuroblasts within the plexuses remain undifferentiated. The muscularis mucosae appears in the cardia during the 10th week and spreads throughout the entire stomach by the 14th week.

14–30 Weeks The cardiac and pyloric villi become prominent at this time (Fig. 4.26). They often disappear by 30 weeks but may persist beyond this point in some cases. The gastric glands ­elongate, and the specific cell types of the glands continue to differentiate (Figs.  4.27, 4.28, and 4.29). The diffuse gut-­

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Fig. 4.23  Stomach at 12  weeks postfertilization. This is a higher-­ magnification view of the squamous-columnar junction in Fig.  4.22, indicated by the lower asterisk. The esophageal stratified squamous epithelium with ciliated columnar epithelium on its surface is to the right. The distinctive pseudostratified cardiac epithelium is to the left (H&E, 20×)

D. S. Huff and C. K. Carreon

Fig. 4.24  Gastric cardiac mucosa at 12 weeks postfertilization. This is a higher-magnification view of the cardiac mucosa in Fig.  4.22. Primitive cardiac pits (asterisks) are present in the pseudostratified, columnar cardiac epithelium. Gastric pits (arrows) with primitive parietal cells indicate the junction between the cardiac and transitional epithelia (H&E, 20×)

Fig. 4.25  Muscularis propria of the stomach at 11 weeks postfertilization. The submucosa occupies the top half of the picture. The three layers of the gastric muscularis propria can be identified: the thin, innermost oblique layer; the thick, inner circular layer; and the thin, outer longitudinal layer. Auerbach’s myenteric plexuses with undifferentiated neuroblasts are prominent. Neuroblasts are migrating from Auerbach’s plexuses up into the submucosa (arrow) to form the deep submucosal plexuses of Henle (asterisks) (H&E, 20×)

associated lymphoid tissue develops in the lamina propria. The neuroblasts of the submucosal and myenteric plexuses continue to differentiate toward ganglion cells.

30–40 Weeks The cardiac and pyloric villi disappear (Fig. 4.30). Gastric glands change little during this period (Fig. 4.31). The diffuse gut-associated lymphoid tissue becomes more abundant, but aggregated lymphoid tissue is uncommon. Maturation of neuroblasts into ganglion cells is nearly complete.

Fig. 4.26  Gastric pyloric mucosa at 24  weeks postfertilization. The pyloric mucosa is villous, and its mucous glands are distinct from the gastric glands. The pyloric sphincter is formed by the inner circular layer of the muscularis propria (between the two arrows). The outer longitudinal layer is below, and Auerbach’s plexuses are visible between the two. The innermost oblique layer is not present in the pyloric sphincter (H&E, 4×)

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Fig. 4.28  Stomach at 19 weeks postfertilization. The full thickness of the wall of the gastric body is shown. The muscularis mucosae is welldeveloped, and the lamina propria is distinctly separated from the submucosa (H&E, 4×)

Fig. 4.27  Stomach at 16 weeks. The full thickness of the wall of the gastric body is shown. The oxyntic mucosa has increased in thickness, and the gastric glands have elongated; compare this image with Fig. 4.20 from age 12 weeks postfertilization, taken at the same magnification. Foveolar epithelium, mucous cells, and parietal cells continue to differentiate. The muscularis mucosae (arrow) is beginning to differentiate. The muscularis propria is very thick, and the outer longitudinal layer, though thicker than previously, is still much thinner than the other layers (H&E, 10×)

Fig. 4.29  Stomach at 19 weeks postfertilization. This image is a higher magnification of the oxyntic mucosa shown in Fig. 4.28. Foveolar epithelium, mucous neck cells, chief cells, and parietal cells all are well-differentiated. The basophilic cytoplasm of the chief cells is distinct from the eosinophilic cytoplasm of the parietal cells. The lamina propria contains scattered lymphoid cells. The muscularis mucosae is distinct (H&E, 20×)

Fig. 4.30 Gastric pyloric mucosa at 35  weeks postfertilization. The pyloric mucosa is no longer villous; compare with Fig. 4.26 (H&E, 10×)

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Fig. 4.31  Oxyntic mucosa at 34 weeks postfertilization. The histology of the oxyntic mucosa has not changed significantly in the last half of pregnancy, as shown; compare with Fig. 4.29 (H&E, 10×)

Small Intestine Less Than 14 Weeks The unique histology of the small intestine is established before the 14th week postfertilization. The multilayered, primitive, pseudostratified columnar epithelium enters the proliferative or vacuolization phase during the sixth week, and the proliferative epithelium transiently occludes the duodenal lumen during the seventh week. The proliferative epithelium regresses, the duodenal lumen is re-established, and villi appear in the duodenum in the eighth week (Fig. 4.32). Crypts of Lieberkühn appear in the duodenum during the ninth week (Fig. 4.33). The villi and crypts spread from rostral to caudal and arrive at the terminal ileum in the 12th to 14th weeks (Figs. 4.34, 4.35, 4.36, and 4.37). Some subsets of mucosal cells, such as enterocytes, goblet cells, gastrin cells, and somatostatin cells, differentiate prior to 14  weeks. Brunner’s glands appear in the first part of the duodenum as early as the 14th week. The inner circular layer of the muscularis propria appears in the duodenum in the seventh week, and the outer longitudinal layer appears in the eighth week; they differentiate from rostral to caudal and arrive at the terminal ileum in the ninth to tenth week. The submucosal and myenteric plexuses differentiate in synchrony with the muscle coats; the myenteric plexus appears first, followed by the submucosal plexuses of Henle and Meissner. 14–30 Weeks Brunner’s glands continue to differentiate from rostral to caudal throughout the rest of the duodenum during the 14th to 16th weeks (Figs. 4.38, 4.39, and 4.40). Brunner’s

D. S. Huff and C. K. Carreon

Fig. 4.32  Duodenum at 11 weeks postfertilization. The primitive villi are covered by differentiating enterocytes. The epithelium between the villi is multilayered, pseudostratified primordial epithelium, which is forming primitive crypts in some areas and will differentiate into single-­ layered surface epithelium in the future. The muscularis mucosae and Brunner’s glands have not yet differentiated. Although the inner circular layer of the muscularis propria is well-differentiated, the external longitudinal layer is only one or two cells thick. Auerbach’s plexus, with its undifferentiated neuroblasts, forms an almost continuous ring in the intermuscular septum. Pancreatic acini are in the left lower corner (H&E, 10×)

Fig. 4.33  Duodenum at 12 weeks postfertilization. The enterocytes are more well-differentiated than they were previously, and goblet cells have appeared. Crypts of Lieberkühn are becoming recognizable. The primordial epithelium seen previously is differentiating into the crypts and surface epithelium. Brunner’s glands have not yet differentiated. The muscularis propria is on the left, and Auerbach’s plexus contains sheets of neuroblasts (H&E, 20×)

glands are sparse distal to the papilla of Vater. The transition from pyloric to duodenal mucosa is gradual in some cases, and duodenal-like villi may be found in the distal

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Fig. 4.34  Small intestine (jejunum or ileum) at 11 weeks postfertilization. The jejunum and ileum cannot be differentiated histologically from the duodenum at this gestational age, and the duodenum can be identified only by its attachment to the head of the pancreas. Notice the villi, intervillous primordial epithelium, primitive crypts of Lieberkühn, the two layers of muscularis propria, and the ring of Auerbach’s plexus (H&E, 4×)

Fig. 4.35  Small intestine at 11 weeks postfertilization. This image is a higher magnification of Fig. 4.34. Note the villi, primitive crypts, muscularis propria (lower right), and Auerbach’s plexus. The muscularis mucosae has not yet differentiated (H&E, 20×)

Fig. 4.36  Small intestine (not the duodenum) at 12 weeks postfertilization. The villi, enterocytes, and crypts of Lieberkühn are better differentiated than in the previous figures. The muscularis mucosae is absent, and the muscularis propria and Auerbach’s plexus have had minimal changes (H&E, 10×)

Fig. 4.37  Small intestine at 12  weeks postfertilization. This is a higher-magnification view of the small intestinal mucosa in Fig. 4.36 (H&E, 20×)

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pylorus, and pyloric-like epithelium may be found in the duodenum. The villi of the small intestine grow longer (Fig.  4.41) and are covered by enterocytes with distinct brush borders, goblet cells, and a few endocrine cells. The stem cells of the intestinal mucosa reside in the base of the crypts of Lieberkühn. They proliferate and give rise to the enterocytes, goblet cells, and endocrine cells, which migrate centripetally up the crypts and to the tips of the villi, where they may undergo apoptosis and slough into the lumen. Sembra et  al. [4] state that “inclusions” are detected in the intestinal villous epithelium in the 16th week, but these authors do not describe or illustrate the inclusions. Inclusions are seen in small intestinal epithelium of some of our cases between 14 and 20 weeks. Most are brightly eosinophilic, but some are orange or brown (Figs.  4.42, 4.43, and 4.44). They resemble the hyaline globules or thanatosomes thought to be related to apoptosis described in adult intestinal epithelium in some conditions [25]. The intestinal mucosa is said to closely resemble that of infants by 20  weeks (Fig.  4.45). The villi may appear shortened, blunted, or bent in segments distended by meconium (Fig.  4.46). Clusters of basally granulated cells called Segi’s caps are seen at the tips of the villi after 20 weeks; these are thought to be precursors of endocrine cells, but little has been written about them [26]. Perfectly preserved tissue and special stains are needed to see them. Paneth cells are easily recognized in the bases of the crypts because of their brightly eosinophilic, coarse supranuclear granules (Fig.  4.47). The neuroendocrine cells, on the other hand, are identified by the presence of red subnuclear granules (Fig.  4.48). The muscularis mucosae appears in the duodenum as early as the 21st week and spreads caudally. It may arrive at the terminal ileum by 30 weeks in some cases, but not until later in others. The neuroblasts in the myenteric and submucosal plexuses continue to differentiate toward ganglion cells (Fig. 4.49). The development of the gut-associated lymphoid tissue is ongoing (Figs. 4.50 and 4.51).

30–40 Weeks The subtypes of mucosal epithelial cells continue to differentiate. Brunner’s glands enlarge and spread caudally, but few are seen in the third and fourth portions of the duodenum. The glands are largest near the pyloric sphincter and are progressively smaller caudally (Figs. 4.52 and 4.53). The development of the muscularis mucosae is completed with its arrival at the ileocecal junction from the duodenum. Neuroblasts of the submucosal and myenteric plexuses continue to differentiate toward mature ganglion cells.

D. S. Huff and C. K. Carreon

Fig. 4.38  Duodenum at 17 weeks postfertilization. Brunner’s glands are budding off the crypts of Lieberkühn. The arrow at the lower right points to the junction of a crypt and a Brunner’s gland. The cells of the crypts are basophilic, whereas those of Brunner’s glands are foamy and light pink. The two layers of the muscularis propria and Auerbach’s plexus are at the bottom of the picture. The neuroblasts show little evidence of differentiating toward ganglion cells. The subserosa and serosa are visible below the muscularis propria (H&E, 10×)

Large Intestine, Appendix, and Anal Canal Less Than 14 Weeks The primordial multilayered epithelium of the large intestine and appendix enters a minor proliferative phase during the seventh postfertilization week. By the 10th to 12th week, the proliferative epithelium differentiates into colonic epithelium characterized by simple columnar mucous surface epithelial cells called colonocytes, primitive villi, and crypts (Fig. 4.54). The colonic mucosa is villous from this point in development until the 30th week or later. Differentiating sections of colon from small intestine is difficult, especially prior to 16 weeks. Later in gestation, the colonic epithelium is predominantly composed of goblet cells, so differentiating it from the small intestine may be easier. The muscularis propria is completely developed by the eighth or ninth week. The mucosa of the appendix is similar to that of the colon except that it lacks villi. By the eighth week, the structures of the anal canal are complete. The transition from colonic mucosa to the anal transitional zone epithelium to the distal, stratified squamous epithelium is distinct, and the anal columns and sphincters are formed. 14–30 Weeks The villi become more distinct and diffuse and are covered by cells similar to colonocytes. The crypts deepen,

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Fig. 4.39  Duodenum (first portion) at 20  weeks postfertilization. Brunner’s glands are becoming more complex and larger tubuloalveolar glands, as shown here (H&E, 10×)

and the villi elongate (Figs. 4.55 and 4.56). As in the small intestine, the villi may not be apparent in segments distended by meconium (Fig.  4.57). The base of the crypts contains the stem cells and proliferating cells, which differentiate into colonocytes, goblet cells, endocrine cells, and Paneth cells. Colonocytes are tall, columnar mucusproducing cells with basal nuclei and a brush border. The colonocytes and goblet cells migrate centripetally to the tips of the villi and undergo apoptosis or are sloughed into the lumen. The endocrine cells migrate to the midregion of the crypts; Paneth cells, which are confined to the right colon, remain in the base of the crypts. Gut-associated lymphoid tissue develops as described above and is most prominent in the cecum and rectum. The muscularis mucosae appears at the anorectal junction and grows rostrally toward the ileocecal junction. Tenia coli appear by 16 weeks. The mucosal villi gradually disappear and are usually not seen after the 30th week. The neuroblasts in the submucosal and myenteric plexuses begin to differentiate toward gan-

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Fig. 4.40 Duodenum (second portion) at 24  weeks postfertilization. Brunner’s glands are less well-developed in this second portion of the duodenum than the Brunner’s glands in the first part of the duodenum shown in Fig. 4.39, from a fetus that was 4 weeks younger. This difference is because the glands first develop near the pyloric sphincter and progress from rostral to caudal. The junctions between two crypts and two Brunner’s glands are indicated by asterisks in the lumen of the crypts. Nests of neuroblasts are visible in the deep submucosal plexus of Henle. Partially differentiated ganglion cells with a small amount of eosinophilic cytoplasm are present in Auerbach’s plexus. The muscularis propria, especially the outer longitudinal layer, has grown thicker (H&E, 10×)

glion cells, but this maturation lags behind that of the more proximal segments, owing to the rostral-to-caudal developmental gradient. Prior to 20 weeks, the histology of the appendix is similar to that of the large intestine, except that villi are inconspicuous and the inner circular and outer longitudinal layers of the muscularis propria are indistinct (Fig. 4.58). Gut-associated lymphoid tissue is inconspicuous and composed of scattered lymphocytes and occasional small lymphoid aggregates in the lamina propria. Primary follicles occupying the submucosa are not common (Fig.  4.59). Beginning at 20  weeks, larger primary follicles become common. In some cases, the appendiceal crypts become

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Fig. 4.41  Small intestine at 17 weeks postfertilization. Compare this image with that of the small intestine mucosa at 12  weeks shown in Figs. 4.36 and 4.37; the villi and crypts are more mature. The nuclei of all the enterocytes are located at the basal aspect of the cells, and goblet cells are numerous. Note the continued absence of the muscularis mucosae (H&E, 10×)

microcystically dilated and push the muscularis mucosae downward, forming microdiverticula, so that the deep border of the mucosa becomes irregular and disorganized (Fig. 4.60). In 1912, Lewis [12] described these changes in the fetal appendix. The lumens of the cystic crypts often contain apoptotic debris and eosinophils (Figs. 4.61, 4.62, and 4.63); polygonal and spindled myoid-­like cells, eosinophils, and aggregated lymphoid follicles may be seen surrounding the cysts. The cystic crypts begin to regress and disappear; they may leave behind a tiny clump of calcified debris or even a small granuloma, occasionally with giant cells (Figs. 4.63 and 4.64). This process results in a disorganized mucosa with extensive areas of crypt dropout, and a “well-organized” mucosa may not be regained until a few weeks after birth. Endocrine cells may be numerous

D. S. Huff and C. K. Carreon

Fig. 4.42  Small intestine at 17 weeks postfertilization. Intracytoplasmic hyaline globules are shown in this image. The globules are brightly eosinophilic, round to oval, and of various sizes, often very large. They seem to be most numerous at the tips of the villi and apparently do not occur in the epithelium of the crypts. Note the other histological features of the small intestine wall (H&E, 10×)

(Figs. 4.65 and 4.66). Based on our experience at our institution, these cystic crypts begin to appear at approximately 17  weeks gestation (15 weeks postfertilization), peak at about 20–25  weeks gestation (18-23 weeks postfertilization), and start to disappear after 28  weeks gestation (26 weeks postfertilization). Also, the previously described histologic features may be observed simultaneously occurring in different crypts within this time frame. The function of these structures is unclear; they do not seem to be affected by the presence or absence of amniotic fluid or fetal systemic infection, which favors their being a part of the normal gut development process and not some sort of

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Fig. 4.43  Small intestine at 17 weeks postfertilization. In this higherpower view, the hyaline globules are seen in more detail (H&E, 20×)

inflammatory response. Perhaps they may have a role in the assembly of mucosa-associated lymphoid tissue, as aggregated gut-associated lymphoid tissue is also prominent at about this period. Because of the temporal occurrence and transient nature of the cystic crypts, we believe that their presence can be used as an additional histologic marker for midtrimester gestation [27]. The anal canal, which is completely formed by the eighth postfertilization week, continues to mature (Fig. 4.67). The anal transitional zone epithelium is stratified cuboidal and columnar epithelium with peg cells on the surface and goblet cells scattered within the epithelial membrane and on its surface. It is distinct from both the

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Fig. 4.44  Small intestine at 19  weeks postfertilization. This highpower image shows hyaline globules in another case. The epithelium is slightly macerated, and the tips of the villi are at the top of the picture. Some globules are bright red, some are orange, and some are brown. Goblet cells are bright blue (H&E, 60×)

squamous epithelium of the anus and the colonic mucosa (Figs.  4.68 and 4.69). Anal glands and anogenital mammary-like glands (previously called anogenital sweat glands) described in adults have not been fully studied in fetuses, but they are thought to be a potential origin of neoplastic processes in this location in adults [28]. Neuroblasts and differentiating ganglion cells of the myenteric and submucosal plexuses are present at the squamous-columnar junction and become numerous within a few millimeters of the junction. The hypoganglionic segment is less than 1 cm in most fetuses (see Fig. 4.67).

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Fig. 4.46  Small intestine at 19 weeks postfertilization. This segment is distended with meconium, which flattens the mucosal surface and nearly obliterates the villi (H&E, 10×)

Fig. 4.45  Small intestine at 19 weeks postfertilization. The histology of the mucosa is nearly identical to that of a full-term infant, and the mouths of the crypts of Lieberkühn are well illustrated. The muscularis mucosae is not present, although the muscularis propria and Auerbach’s plexus are well-­developed (H&E, 10×)

Fig. 4.47  Small intestine (terminal ileum) at 14  days postnatal age. Paneth cells in the base of the crypts are characterized by coarse, brightly eosinophilic, supranuclear granules (H&E, 60×)

Fig. 4.48  Small intestine at 21 weeks postfertilization. Two neuroendocrine cells are seen toward the base of the crypt (arrows) and are identified by their red subnuclear granules. For comparison, a Paneth cell is present halfway between the two neuroendocrine cells (H&E, 40×)

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Fig. 4.49  Auerbach’s plexus in the duodenum at 19 weeks postfertilization. The neuronal cells are a mixture of neuroblasts (the small, round blue cells with little cytoplasm) and partially differentiated ganglion cells (the larger cells with finely granular chromatin, a small nucleolus, and eosinophilic cytoplasm, one of which is indicated by the arrow) (H&E, 20×)

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Fig. 4.50  Gut-associated lymphoid tissue in the small intestine at 19 weeks postfertilization. Prior to 20 weeks, the gut-associated lymphoid tissue is usually characterized by inconspicuous nodules of lymphoid cells in the upper lamina propria, such as the one marked by the arrows along its deep surface. Lymphoid cells are also diffusely scattered throughout the lamina propria (H&E, 20×)

Fig. 4.51  Gut-associated lymphoid tissue in the small intestine (terminal ileum) at 20 weeks postfertilization. Peyer’s patches are aggregates of multiple, submucosal lymphoid follicles fused into large sheets; they are more common at 24  weeks or later than at 20  weeks, as in this instance. This slightly precocious appearance of Peyer’s patches might raise the question of fetal infection or incorrect dates (H&E, 4×)

30–40 Weeks The villi have disappeared, and the crypts elongate and acquire the well-organized, parallel test tube arrangement typical of postnatal colonic epithelium (Fig. 4.70). The neuroblasts continue to differentiate, but only a few completely mature ganglion cells are seen (Figs.  4.71 and 4.72). The hypoganglionic segment remains less than 1 cm long.

Fig. 4.52  Duodenum (first portion) at 34  weeks postfertilization. Brunner’s glands are complex and large, as shown. Most of the glands are in the submucosa beneath the muscularis mucosae (MM), but a few of the glands are in the lamina propria above the muscularis mucosae among the crypts of Lieberkühn, which are the basophilic glands in the upper left-hand corner of the picture (H&E, 10×)

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D. S. Huff and C. K. Carreon

Fig. 4.54  Large intestine at 12 weeks postfertilization. Primitive villi are covered by differentiating colonocytes, and the surface epithelium between the villi is primordial, multilayered, and pseudostratified. Primitive crypts of Lieberkühn are beginning to evolve from the primordial epithelium. The muscularis mucosae is not yet present. The inner circular layer of the muscularis propria is well-developed, but the outer longitudinal layer is only one or two cells thick. Auerbach’s plexus is a continuous ring of neuroblasts in the intermuscular septum. At this gestational age, the colon is difficult to distinguish histologically from the duodenum (see Fig. 4.32) or the small intestine (see Figs. 4.34 and 4.35) (H&E, 20×)

Fig. 4.53  Duodenum (second portion) at 35  weeks postfertilization. Brunner’s glands are shown; compare with Fig.  4.52. The gestational ages are nearly identical, but the Brunner’s glands in the second portion of the duodenum in this picture are smaller and less numerous than in the first portion of the duodenum in the previous picture. The mucosa is at the top and the muscularis propria is at the bottom. Note the solitary primary lymphoid follicle of gut-associated lymphoid tissue (H&E, 10×)

Fig. 4.55  Large intestine at 15 weeks postfertilization. The villi are longer, and most of the nuclei of the colonocytes are located basally. Goblet cells are present. The crypts of Lieberkühn are more definite. The outer longitudinal layer of the muscularis propria is very thin (H&E, 10×)

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Fig. 4.56  Large intestine at 19  weeks postfertilization. The mucosal villi are well formed and long, and goblet cells are numerous. The crypts of Lieberkühn are apparent. The villous colonic epithelium may be difficult to distinguish from small intestine mucosa. The muscularis mucosae is beginning to develop and is seen here as a thin, discontinuous longitudinal layer of spindle cells below the lamina propria. The muscularis propria is artifactually absent (H&E, 10×)

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Fig. 4.57  Large intestine at 19 weeks postfertilization, from the same fetus as in Fig. 4.56. This segment is distended by meconium, which flattens the mucosa and obliterates the villi, similar to the situation in the small intestine in Fig. 4.46 (H&E, 4×)

Fig. 4.58  Appendix at 19  weeks postfertilization. The histology is similar to the large intestine except that villi are shorter and less frequent. Diffuse and small aggregates of gut-associated lymphoid tissue are present (H&E, 4×)

Fig. 4.59  Gut-associated lymphoid tissue in the appendix at 19 weeks postfertilization. Solitary lymphoid follicles covered by follicle-associated epithelium begin to form in the submucosa. Eosinophils are seen in the lamina propria in the upper left. Two endocrine cells with subnuclear eosinophilic granules are seen in the crypt on the left (H&E, 20×)

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Fig. 4.60 Appendix at 24  weeks postfertilization. Microcystically dilated crypts forming microdiverticula are seen. The dilated crypts are filled with apoptotic debris and inflammatory cells, especially eosinophils, and are surrounded by polygonal and myoid cells. The cystic changes begin to appear at about 17 weeks gestation (15 weeks postfertilization). Follicular gut-associated lymphoid tissue is abundant (H&E, 4×)

Fig. 4.61  Appendix at 24 weeks postfertilization. This higher-power view is of the cystically dilated crypt seen in Fig. 4.60. The cyst is filled with apoptotic and inflammatory debris, but there is neither appendicitis nor neutrophils in the lumen of the appendix (H&E, 10×)

Fig. 4.62  Appendix at 21 weeks postfertilization. A prominent cuff of myoid-like spindle cells surrounds the cystic crypts. Adjacent aggregates of lymphocytes admixed with eosinophils are also present (H&E, 20×)

Fig. 4.63  Appendix at 20 weeks postfertilization (22 weeks gestation). A small cluster of giant cells (arrow) is present and is surrounded by spindle cells and mixed inflammatory cells. A cystically dilated crypt is present in the right upper corner (H&E, 20×)

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Fig. 4.64  Appendix at 20 weeks postfertilization (22 weeks gestation). This is a higher-power view of the giant cell cluster in Fig. 4.63. Note the surrounding spindle cells and inflammatory cells consisting predominantly of lymphocytes and eosinophils (H&E, 40×)

Fig. 4.65  Appendix at 24 weeks postfertilization. Endocrine cells may be common in the appendix at this gestational age (H&E, 20×)

Fig. 4.66  Appendix at 24 weeks postfertilization. This image is a higherpower version of Fig. 4.65. The endocrine cells are characterized by brightly eosinophilic, fine subnuclear granules, in contrast to the coarse supranuclear granules seen in Paneth cells (Compare with Fig. 4.47) (H&E, 60×)

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Fig. 4.67  Anal canal at 21 weeks postfertilization. The first ganglion cells of Auerbach’s plexus are marked by the asterisk on the left. The asterisk on the right marks the level where the number of ganglion cells is similar to that in the rest of the rectum. Between the two asterisks is the hypoganglionic segment, which is 5 mm long in this case. The internal sphincter is formed by the inner circular layer. The arrow indicates the squamous-­columnar junction. AS anal sinus, EXT external sphincter, IC inner circular layer of the muscularis propria, INT internal sphincter, OL outer longitudinal layer of the muscularis propria (H&E, 1×)

Fig. 4.68  Anal canal at 21 weeks postfertilization. This higher-power view is of the squamous-columnar junction indicated by the arrow in Fig. 4.67. The stratified squamous epithelium changes abruptly into the stratified columnar epithelium, with intraepithelial goblet cells characteristic of the anal transitional zone epithelium (H&E, 10×)

Fig. 4.69  Anal canal at 21 weeks postfertilization. This higher-power view shows the anal transitional zone epithelium near the colonic mucosa, demonstrating the details of this unique epithelium (H&E, 20×)

Fig. 4.70  Large bowel at 35 weeks postfertilization. The large intestine mucosa is no longer villous. The crypts of Lieberkühn are straight, parallel, and of equal length, imparting the classic test tube appearance of the large intestinal crypts. The base of the crypts reaches the muscularis mucosae (MM). Lu lumen containing meconium (H&E, 20×)

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Fig. 4.71  Auerbach’s plexus in the rectum at 35 weeks postfertilization. Most of the neuroblasts are undifferentiated small, blue cells with little cytoplasm. A few partially differentiated ganglion cells with larger amounts of eosinophilic cytoplasm are present. Identifying these poorly differentiated cells as ganglion cells in sections of rectal mucosal biopsies to rule out Hirschsprung’s disease is difficult. The inner circular layer is at the top and the outer longitudinal layer is at the bottom (H&E, 20×)

Fig. 4.72  Auerbach’s plexus in the rectum of a 3-week-old neonate. It can be seen that not all ganglion cells are completely mature even in the immediate postnatal period. Several mature ganglion cells are present. They exhibit more abundant wispy cytoplasm, finely granular chromatin, and a small, dark nucleolus. Familiarity with the appearance of both immature and mature ganglion cells is paramount in the evaluation of rectal biopsies for Hirschsprung’s disease (H&E, 40×)

Acknowledgment The authors thank Dr. Tricia R.  Bhatti from The Children’s Hospital of Philadelphia, Philadelphia, PA, for invaluable assistance in selecting microscopic slides for photography, taking photographs, and editing the original manuscript.

10. Lewis F. Development of the stomach. In: Keibel F, Mall FP, editors. Human embryology. Philadelphia: J.B.  Lippincott; 1912. p. 368–81. 11. Lewis F. Development of the small intestine. In: Keibel F, Mall FP, editors. Human embryology. Philadelphia: J.B.  Lippincott; 1912. p. 381–93. 12. Lewis F. Development of the large intestine. In: Keibel F, Mall FP, editors. Human embryology. Philadelphia: J.B.  Lippincott; 1912. p. 393–9. 13. Huff D.  Developmental anatomy and anomalies of the gastrointestinal tract, with involvement in major malformative syndromes. In: Russo P, Ruchelli E, Piccoli DA, editors. Pathology of pediatric gastrointestinal and liver disease. New  York: Springer; 2004. p. 3–36. 14. O’Rahilly R, Muller F. Human embryology & teratology. 3rd ed. New York: Wiley-Liss; 2001. 15. Fu M, Tam PK, Sham MH, Lui VC. Embryonic development of the ganglion plexuses and the concentric layer structure of human gut: a topographical study. Anat Embryol (Berl). 2004;208:33–41. 16. Wallace AS, Burns AJ. Development of the enteric nervous system, smooth muscle and interstitial cells of Cajal in the human gastrointestinal tract. Cell Tissue Res. 2005;319:367–82. 17. Wester T, Eriksson L, Olsson Y, Olsen L. Interstitial cells of Cajal in the human fetal small bowel as shown by c-kit immunohistochemistry. Gut. 1999;44:65–71. 18. Min KW, Sook Seo I. Interstitial cells of Cajal in the human small intestine: immunochemical and ultrastructural study. Ultrastruct Pathol. 2003;27:67–78. 19. Sundler F, Böttcher G, Ekblad E, Håkanson R. The neuroendocrine system of the gut. Acta Oncol. 1989;28:303–14. 20. Carr NJ, Sobin LH. Neuroendocrine tumors of the appendix. Semin Diagn Pathol. 2004;21:108–19. 21. Spencer J, MacDonald TT, Finn T, Isaacson PG. The development of gut associated lymphoid tissue in the terminal ileum of fetal human intestine. Clin Exp Immunol. 1986;64:536–43.

References 1. O’Rahilly R. The timing and sequence of events in the development of the human digestive system and associated structures during the embryonic period proper. Anat Embryol (Berl). 1978;153:123–36. 2. Lewis F.  The early development of the entodermal tract and the formation of its subdivisions. In: Keibel F, Mall FP, editors. Human embryology. Philadelphia: J.B. Lippincott; 1912. p. 295–334. 3. O’Rahilly R, Muller F. Developmental stages in human embryos. Washington, DC: The Carnegie Institution of Washington; 1987. 4. Sembra R, Tanaka O, Tanimura T. The digestive tract. In: Nishimura H, editor. Atlas of prenatal histology. Tokyo: Igaku-Shoin; 1983. p. 193–225. 5. Moxey PC, Trier JS. Endocrine cells in the human fetal small intestine. Cell Tissue Res. 1977;183:33–50. 6. Sjolund K, Sanden G, Hakanson R, Sundler F. Endocrine cells in human intestine: an immunocytochemical study. Gastroenterology. 1983;85:1120–30. 7. Chan OTM, Chiles L, Levy M, Zhai J, Yerian LM, Xu H, et  al. Smoothelin expression in the gastrointestinal tract: implication in colonic inertia. Appl Immunohistochem Mol Morphol. 2013;21:452–9. 8. Montani M, Thiesler T, Kristiansen G.  Smoothelin is a specific and robust marker for distinction of muscularis propria and muscularis mucosae in the gastrointestinal tract. Histopathology. 2010;57:244–9. 9. Lewis F. The development of the esophagus. In: Keibel F, Mall FP, editors. Human embryology. Philadelphia: J.B.  Lippincott; 1912. p. 355–68.

76 22. Otani H, Yoneyama T, Hashimoto R, Hatta T, Tanaka O. Ultrastructure of the developing stomach in human embryos. Anat Embryol (Berl). 1993;187:145–51. 23. Zhou H, Greco MA, Daum F, Kahn E. Origin of cardiac mucosa: ontogenic consideration. Pediatr Dev Pathol. 2001;4:358–63. 24. De Hertogh G, Van Eyken P, Ectors N, Geboes K.  On the origin of cardiac mucosa: a histological and immunohistochemical study of cytokeratin expression patterns in the developing esophagogastric junction region and stomach. World J Gastroenterol. 2005;11:4490–6. 25. Dikov DI, Auriault ML, Boivin JF, Sarafian VS, Papadimitriou JC.  Hyaline globules (thanatosomes) in gastrointestinal epithe-

D. S. Huff and C. K. Carreon lium: pathophysiologic correlations. Am J Clin Pathol. 2007;127: 792–9. 26. Kobayashi S, Iwanaga T, Fujita T. Segi’s cap: huge aggregation of basal-granulated cells discovered by Segi (1935) on the intestinal villi of the human fetus. Arch Histol Jpn. 1980;43:79–83. 27. Carreon CK, Ruchelli ED, Mihok C, Huff DS. Cystic crypt changes in midgestational human vermiform appendix: an unrecognized transient histologic feature. Pediatr Dev Pathol. 2019 May 24:1093526619853180. https://doi.org/10.1177/1093526619853180. [Epub ahead of print] PubMed PMID: 31126217. 28. Mills SE. Histology for pathologists. 4th ed. Philadelphia: Wolters Kluwer Health/Lippincott Williams & Wilkins; 2012.

5

Liver Pierre Russo

Introduction The fetal liver, an important organ with synthetic and hematopoietic functions, undergoes dramatic histological changes during the second and third trimesters of gestation. This chapter reviews the histological changes in the fetal liver evident in hepatocytes, the sinusoidal lining cells, the intrahepatic biliary tree, and the hematopoietic elements and discusses the development of the hepatic vasculature and extrahepatic biliary tree.

Embryology Mechanisms of Development of the Liver Hepatobiliary morphogenesis in humans begins in the third to fourth week of development and arises from two primordia. The hepatocytes, which ultimately account for 70% of the cells in the liver, are derived from the cranial portion of the hepatic diverticulum, an outgrowth of the foregut endoderm that invades the septum transversum, the mesodermal plate that separates the embryonic thoracic and abdominal cavities. The caudal part of the hepatic diverticulum (the portion that does not invade the septum transversum) forms the extrahepatic biliary tree and ventral pancreatic anlage (Fig. 5.1). The stromal cells of the connective tissue elements, Kupffer’s cells, stellate cells, and blood vessels are mesenchymal in origin and develop from the septum transversum and from cells lining the coelomic cavity. The prehepatocytes or hepatoblasts proliferate within the septum transversum, organized in cords around developing sinusoids derived from branches of the P. Russo (*) Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]

Fig. 5.1  Hepatic morphogenesis. A 30-day embryo shows the cephalic portion of the hepatic diverticulum (HD) penetrating the septum transversum and forming cords where the endodermal cells (yellow) will differentiate into hepatoblasts. The caudal portion of the diverticulum will form the gallbladder (GB) and biliary tree

vitelline veins that penetrate the septum transversum [1– 3]. Development of the hepatic primordium is dependent on continuous interactions with the adjacent mesoderm and on the concerted development of the hepatic vasculature, as revealed by studies of Klk1−/− mouse embryos, which lack mature endothelial cells, with consequent inhibition of expansion of the liver bud [4]. The molecular mechanisms that control liver development are the subject of several excellent reviews [5, 6]. In summary, these mechanisms appear to be well conserved among various organisms and depend on the coordinated expression of transcription and signaling factors from the cardiac mesoderm, the septum transversum, and the endoderm to commit the endodermal cells to a hepatic fate and to induce and regulate hepatic gene expression, the most important of which include fibroblast growth factor (FGF) from the cardiac mesoderm, bone morphogenetic protein (BMP) from the septum transversum mesenchyme, and WNT sig-

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naling, all of which induce the adjacent endoderm region to adopt a hepatic fate. Growth of the liver bud is very rapid between the fourh and tenth weeks of gestation in humans; by 50 days, 700 portal branches from the second to the sixth order can be counted [7].

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Development of the Hepatic Vasculature The vascular architecture of the liver is unique in that the organ receives both a venous blood supply via the portal vein from the intestines, pancreas, and spleen, which is responsible for its functional circulation, and an arterial supply conveying nutrition and oxygenated blood from the hepatic artery. The development of the liver is closely associated with changes in the omphalomesenteric (vitelline) veins and in the umbilical veins. The definitive venous pattern within the liver is established by the seventh week [8]. The portal vein arises from anastomotic channels derived from the paired vitelline veins in the hepatic primordium. The intrahepatic segments of the portal vein and its branches become surrounded by mesenchyme, from which portal spaces will develop and within which arterial segments will grow. The proximal end of the right vitelline vein forms the terminal part of the inferior vena cava (IVC); its remnants in the liver become the hepatic venous system, and its distal portion will form the splenic and superior mesenteric veins. The left vitelline vein forms part of the IVC. By the sixth week, the right umbilical vein disappears, and all oxygenated blood from the placenta enters the liver from the left umbilical vein. After the umbilical vein enters the liver, it gives off numerous branches to the left lobe and connects to the portal sinus, derived from the subhepatic anastomosis of the disappearing vitelline veins. This connection allows for a substantial portion of the oxygenated blood from the umbilical vein to be conveyed to the hepatocytes via the sinusoidal network [7]. The left angle of the portal sinus is connected to the IVC by the ductus venosus, an oblique channel formed by the coalescence of some of the hepatic sinusoids, which connects the umbilical vein to the IVC, diverting a portion (probably less than 20% at 30 weeks gestation) of the oxygenated blood directly to the heart [7] (Fig. 5.2). Hemodynamic studies performed during intrauterine life have confirmed that the umbilical vein is the predominant liver vessel until birth and that the left lobe is supplied mainly by oxygenated blood from the umbilical vein, whereas the right lobe of the liver receives blood predominantly from the portal vein [7]. This vascular asymmetry is believed to account for differences observed between the right and left lobes of the liver in stillbirths and neonates [9, 10]. At birth, the left lobe is comparably larger than in later life. Birth results in the closure of the umbilical circulation, starting

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Fig. 5.2  The ductus venosus in the fetal liver. (a) A catheter in the umbilical vein with its tip in the sinus intermedius (or portal sinus) points toward the ductus venosus. A portion of oxygenated blood from the umbilical vein drains directly into the ductus venosus and hence into the inferior vena cava during fetal life. (b) Cut section from a 29-week fetus further illustrates the relationship between the ductus venosus, portal sinus, umbilical vein, and portal vein

with collapse of the umbilical arteries and followed by closure of the umbilical venous flow, which is completed by 15–20 days. This decrease in blood flow results in collapse of the ductus venosus and its obliteration by fibrous tissue. The umbilical vein becomes the ligamentum teres, and the ductus venosus transforms into the ligamentum venosum. Fetal blood flow through the hepatic artery is insignificant compared with that delivered by the umbilical and portal veins. The development of the hepatic arterial system parallels that of the bile ducts and reaches maturity at approximately 15 years of age [8].

Histology Hepatic Lobule Hematopoiesis is one of the defining histological characteristics of the developing liver, beginning between the sixth and seventh

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Fig. 5.3  Fetal liver at 16 weeks gestation. (a) Marked hematopoiesis is present in the sinusoids and the portal tracts. (b) Hematopoiesis tends to segregate in the fetal liver, with erythropoiesis present in the sinusoids; myelopoiesis is seen in the portal tracts (hematoxylin and eosin (H&E): a 4×; b 40×.)

Fig. 5.4  Fetal liver at 19 weeks gestation. (a) Kupffer cells highlighted by an immunostain for CD163. (b) A higher-power view illustrates the “Bessis island,” a specialized microenvironment in which erythroblasts proliferate, consisting of a central macrophage with cytoplasmic protrusions extending around surrounding erythroblasts (CD163 immunohistochemistry: a 10×; b 60×)

weeks of gestation. Intense hematopoietic activity is present through the second trimester (Figs. 5.3, 5.4, 5.5, 5.6, 5.7, and 5.8), as the liver is the main hematopoietic organ in the body at that time. Additionally, hematopoietic cells appear essential for hepatocyte maturation, because they secrete oncostatin M, which activates a signaling cascade and is responsible for the expression of enzymes such as glucose-6-phosphatase [5]. Hematopoiesis has a distinct zonal distribution (see Fig. 5.3b). The production of erythroid cells occurs within the sinusoids, partly in “Bessis islands,” which consist of a central macrophage surrounded by erythroid cells (see Fig. 5.4b). These macrophages are believed to play an essential role in erythroid maturation [11]. Myelopoiesis is located mainly around the vascular structures within the portal triads. Hematopoietic activity begins to decrease between 24 and 32 weeks of gestation, as the bone marrow becomes hematopoietic [12]. By 36 weeks, hematopoiesis remains only as scattered islands in the hepatic paren-

chyma (Fig. 5.9), and it normally ceases within a few weeks after birth [13]. Increased erythropoiesis may be seen in cases of chronic fetal hypoxia or anemia and in infants of diabetic mothers [14]. Erythropoiesis has been reported to decrease as a result of preeclampsia, suggesting that erythropoiesis in utero may also depend on factors other than just oxygen supply [15]. Fetal myelopoiesis has been observed to be elevated as a result of intrauterine infection, and this observation may help confirm a diagnosis of chorioamnionitis when the placenta is unavailable for examination [16–18]. The early hepatocyte cords that form around the developing sinusoids remain thick throughout intrauterine life and the first few months after birth, thinning to two-cell trabeculae by 5  months of life. By 5  years of age, they adopt the single-cell cord architecture of adult life (Fig.  5.10). Glycogen becomes plentiful in fetal hepatocytes and remains so until birth (see Fig.  5.5b); lipofuscin is usually scanty.

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Fig. 5.5  Fetal liver at 22 weeks gestation. (a) A bile duct emerges from the limiting plate. (b) The fetal hepatocytes feature an alternating light and dark pattern resulting from cytoplasmic glycogen content. These hepatocytes are similar to those noted in “fetal”-type hepatoblastomas (H&E: a 10×; b 20×)

Ultrastructurally, rough endoplasmic reticulum is prominent by the seventh week and peroxisomes by the eighth week; the Golgi apparatus moves from the perinuclear to the pericanalicular zone during the third month, as bile formation begins [19]. The lateral domains have tight junctions separating canaliculi from sinusoids. The canalicular and basolateral domains of hepatocytes contain specific transporters for organic and inorganic ions and macromolecules [20]. The canaliculus, the smallest component of the biliary system, develops by 7–8 weeks of gestational age [21]. Enzymes of the ornithine cycle are expressed early in the human liver. It appears that the liver changes from a primarily glycolytic role during the first trimesters to a gluconeogenic role before birth [22]. Metabolic zonation, whereby periportal hepatocytes differ from pericentral hepatocytes on the basis of differential expression of metabolism-regulating genes, begins shortly after birth and appears to be under the control of Wnt signaling and the β-catenin pathway [6].

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Fig. 5.6  Fetal liver at 23 weeks gestation. Prominent sinusoidal hematopoiesis (a) is noted. A large portal tract shows several well-formed bile ducts. Bile ducts are mostly still connected to the limiting plate in the smaller portal tracts (b) and in the more peripheral portal tracts (c); the liver capsule is noted at the upper left (H&E: a 4×; b 20×; c 10×)

The endothelial cells that line the sinusoids are believed to originate either from the septum transversum or from the disrupted vitelline veins and play a key role in hematopoiesis during fetal life. They undergo significant remodeling, with

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Fig. 5.7  Fetal liver at 28 weeks gestation. (a) Sinusoidal hematopoiesis has decreased in this low-power view. (b) Small clusters of hematopoiesis are noted in the sinusoids. Bile ducts can be seen in most portal tracts; the ductal plate is less apparent except in the smallest portal tracts (H&E: a 4×; b 20×)

Fig. 5.8  Fetal liver at 30 weeks gestation. (a) Hematopoiesis is present mainly as scattered clusters in the lobule. (b) Clusters of erythropoiesis around a portal tract; most portal tracts by now show no evidence of hematopoiesis (H&E: a 4×; b 20×)

development of fenestrations and changes in immunophenotypic features during growth of the liver, progressively losing typical endothelial markers such as CD34 and acquiring distinctive markers such as CD4 and ICAM-1, as expressed by the postnatal liver (Fig. 5.11) [7]. This fenestrated endothelium also allows fluid passing through channels between hepatocytes to enter the space of Disse and thence to drain into portal lymphatic spaces [23]. Kupffer cells are hepatic macrophages that belong to the mononuclear phagocyte system. They are derived from the yolk sac and, later, from the bone marrow. Large numbers of macrophages are present in the sinusoids after 30 days gestation, identifiable immunohistochemically by markers such as CD68 and CD163 (see Fig.  5.4a); they are characterized ultrastructurally by abundant cytoplasm, numerous filopodia, dense bodies, and prominent hemophagocytosis [24]. Hemosiderin is normally present mainly within periportal hepatocytes and in the ductal plate in the fetal liver (Fig. 5.12), though there is great variability in the amount of hemosiderin

normally present [21, 25]. When hepatocyte siderosis is abundant, it is also present within Kupffer cells. Measured iron content appears to normally increase with gestational age [25]. The distribution of the iron stores may shift from a primarily periportal to a perivenular location in cases of amniotic fluid infection [18]. Hemosiderin accumulation may increase with perinatal liver damage and also in conditions such as hemolysis without evidence of liver damage [25].

Biliary Tract  he Intrahepatic Biliary Tree T The development of the intrahepatic biliary tree begins between the fifth and ninth week postfertilization. Biliary epithelium is believed to arise from precursor bipotential hepatoblasts that can differentiate into either hepatocytes or biliary epithelial cells [26]. Hepatoblasts at the outer boundary of the portal tracts acquire a biliary phenotype,

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Fig. 5.9  Fetal liver at 37 weeks gestation. (a) Hematopoiesis is much diminished. (b) Bile ducts can be observed in most portal tracts (H&E: a 4×; b 10×)

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Fig. 5.10  Reticulin staining of the hepatic cords at various ages. (a) At 16 weeks gestation, cords are poorly formed and many cell layers in thickness. (b) At 23  weeks gestation, reticulin surrounds most of the cords, which remain several layers in thickness. A portal tract is seen at the left edge of the image and a central vein to the right. (c) At 36 weeks

gestation, the reticulin framework is more developed, and most hepatic cords are two to three cell layers in thickness. (d) By contrast, a single-­ cell layer arrangement prevails in the cords of a 5-year-old boy (reticulin stain: a–d 20×)

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Fig. 5.11  Endothelial sinusoids stained with CD34. (a) In a 16-week fetus, immunohistochemical staining with CD34 highlights the vasculature of the portal tracts and the sinusoids of the periportal areas. (b) In

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a 36-week fetus, CD34 staining has mostly disappeared from the sinusoids and is essentially limited to the portal vasculature and central veins (CD34 immunohistochemistry: a, b 20×)

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Fig. 5.12  Iron accumulation. (a) At 16 weeks gestation, hemosiderin accumulation in the fetal liver is primarily within the ductal plate and periportal hepatocytes. (b) At 24  weeks gestation, hemosiderin accumulation is still prominent, outlining the limiting plate. (c) At 37 weeks

gestation, hemosiderin accumulation is less prominent. (d) Increased hemosiderin accumulation is noted in the periportal areas and in surrounding hepatocytes in this neonate born at 36 weeks gestation with severe hypoxia (iron stain: a 20×; b 4×; c 20×; d 20×)

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forming the ductal plate (Fig. 5.13) [27, 28]. A combination of periportal signals, including Notch, transforming growth factor beta (TGF-β), WNT, and FGF, along with transcription factors such as SOX4 and SOX9, is responsible for differentiation of hepatoblasts toward a biliary fate [6, 29]. The downstream transcription factor HNF-1β is required for the development of interlobular bile ducts and arteries in mice, and HNF-1β −/− mutant mice have a paucity of intrahepatic bile ducts and an absence of hepatic arteries [30, 31]. Remodeling of the ductal plate, starting around week 12, involves formation of discrete tubular spaces, incorporation of the tubules more centrally into the portal tract mesenchyme, and elimination of excess epithelial elements, as evidenced by staining with cytokeratin-7 and cytokeratin-19 (CK19) (Figs.  5.14, 5.15, and 5.16). This process extends Fig. 5.13  Biliary development. A portal tract from a 14-week fetus is centrifugally from the hepatic hilum toward the periphery of delimited by a prominent and focally bilayered ductal plate. There is a the liver and continues throughout fetal life, but with a “slow-­ central portal vein, and a small hepatic artery is noted (H&E, 10×) a

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Fig. 5.14  Biliary development in a 16-week fetus. (a) In a larger portal tract, a bile duct lined by a cuboidal epithelium (arrow) is distinguishable from the ductal plate. The bilayered ductal plate is apparent in the superior portion of this portal space, as well as in an adjacent portal space to the right (arrowheads) (b) Immunostaining with cytokeratin 19

outlines the ductal plate, as well as early interlobular bile ducts being incorporated more centrally by the portal mesenchyme. (c) By contrast, bile ducts are not apparent in a smaller, more peripheral portal tract in the same liver. (d) Immunostaining with cytokeratin 19 outlines the ductal plate (H&E: a, c 10×; CK19 immunohistochemistry: b, d 10×)

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Fig. 5.15  Biliary development in a 24-week fetus. (a) More mature bile ducts separate from the ductal plate and begin to be incorporated more centrally in this larger portal tract. The ductal plate is recognizable at the periphery of the uppermost portion of the portal tract. (b) Immunostaining with CK19 shows a more discontinuous ductal plate (a H&E, 10×; b CK19 immunohistochemistry, 10×)

Fig. 5.16  Biliary development in a 36-week fetus. (a) Most portal tracts have assumed the configuration that will prevail throughout life, with a well-formed bile duct comparable in diameter to the adjacent hepatic artery. (b) Immunostaining with CK19 highlights a mature bile duct and canals of Hering in the immediate periportal area. The ductal plate has become essentially unapparent (a H&E, 10×; b CK19 immunohistochemistry, 10×)

down” period between the 20th and 32nd weeks, when portal myelopoiesis is active [32]. Histologic examination of the fetal liver thus reveals more mature ducts in the hilar region, while the ducts in the periphery are still in ductal plate configuration [28] (see Fig. 5.6c). This remodeling is not complete at birth but continues into the first month of life ex utero [33]. Thus, the “normal” bile duct/portal tract ratio is likely lower in the premature infant than in the term neonate [27, 32]. The ductal plate malformation as observed in disorders such as congenital hepatic fibrosis and other “ciliopathies” may represent a form of “maturation arrest” of the fetal ductal plate (Fig. 5.17).

 he Extrahepatic Biliary Tree T The extrahepatic biliary tree is visible as a bud on the hepatic diverticulum by the fourth week postfertilization. By the fifth week, the gallbladder, cystic duct, hepatic ducts, common bile duct, and pancreatic duct are well demarcated. The anlage of the gallbladder and biliary tree have a continuously

Fig. 5.17  Congenital hepatic fibrosis in a 34-week fetus. Note the presence of a prominent residual double-layered ductal plate around almost the entire circumference of the portal tract, more similar to the appearance of a portal tract in a 16-week fetus (H&E, 10×)

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Fig. 5.18  Gallbladder and porta hepatis at 23  weeks gestation. (a) Gallbladder with prominent mucosal infoldings. (b) The muscle layer is well developed. (c) Hepatic duct at the liver hilum. The shortest transverse diameter of this hepatic duct is about 100 μm. (d) Nearest to the

liver edge, branches of the hepatic ducts close to the liver parenchyma connect to peripheral ductal plate elements. (e) Section adjacent to (d) with CK19 immunostaining. (H&E: a 2×; b 10×; c 4×; d 10×. e CK19 immunohistochemistry, 10×)

patent lumen throughout gestation, visible as early as 29 days postfertilization, in direct continuity with converging ductules at the hepatic hilum and with the developing intrahepatic ducts; it does not have a solid stage, as was previously suggested [34, 35]. The major bile ducts at the porta hepatis

are established by the eighth week of gestation (Figs. 5.18, 5.19, and 5.20) [36]. The distal and middle extrahepatic ducts take shape by 9 weeks gestational age (see Fig. 5.19). The extrahepatic bile duct epithelium contains neutral and acidic mucins by 10–12  weeks gestational age (see

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Fig. 5.19  Distal common bile duct and ampulla in a 23-week fetus. (a) The duct is surrounded by pancreatic tissue. Part of the duodenum is seen at the right side. (b) The mucosa has infoldings, and the epithelium is columnar. (c) The duct is partly surrounded by thin wisps of smooth

muscle. (d) The columnar epithelium is coated by a periodic acid-Schiff (PAS) stain-positive glycocalyx (H&E: a 2×; b 10×. (c) Gomori-­ trichrome stain, 20×. (d) PAS stain, 40×)

Fig.  5.19d). Peribiliary glands and a peribiliary capillary plexus appear to be formed by 30 weeks gestational age (see Fig. 5.20) [36]. Figure 5.21 illustrates several developmental aberrations. Exocrine pancreatic tissue may be seen in all portions of the extrahepatic tree, even near the liver hilum (Fig.  5.21a).

Hyaline cartilage may be found near extrahepatic ducts and may be observed in cases of biliary atresia (Fig. 5.21b) [37]. Ectopic liver is a rare entity and can be found in various locations such as the diaphragm, omentum, stomach, and retroperitoneum or a hernial sac (as in Fig.  5.21c, d); the most commonly reported site is the gallbladder [38].

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Fig. 5.20  Porta hepatis in a 41-week fetus. (a) Hepatic duct at the hilum has an irregular contour. The epithelium dips into the fibrous stroma to form pits or sacculi of Beale. Liver parenchyma is at the upper right of the frame. (b) The diameter of the duct is about 400 μm. The

duct is surrounded by paler-staining, mucin-filled peribiliary glands. (c) The hilar hepatic duct has a fibrous wall ringed by a network of small capillaries. (d) The peribiliary glands are filled with mucin (H&E: a 4×; b 10×. (c) Gomori-trichrome stain, 10×. (d) PAS stain, 20×)

5 Liver

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Fig. 5.21  Selected developmental aberrations. (a) Ectopic pancreatic tissue in the hepatic hilum. A longitudinal section of the bile duct is present toward the top of the frame; liver parenchyma is at the lower left. (b) Nodules of cartilage are noted around the common bile duct near the ampulla of Vater in a newborn infant. The duct (center) is sur-

rounded by pancreas. (c) Cords of pale-staining hepatocytes and bile ducts (lower right) can be appreciated in this inflamed inguinal hernia sac from a 3-year-old boy. (d) Hep Par-1 immunostain outlining the cords of hepatocytes in (c) (H&E: a–c 4×. (d) Hep Par-1 immunohistochemistry, 4×)

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References 1. MacSween RN, Desmet V, Roskams T, Scathorne RJ. Developmental anatomy and normal structure. In: MacSween RN, Burt AD, Portman B, Ishak KG, Scheuer PJ, Anthony PP, editors. Pathology of the liver. 4th ed. London: Churchill Livingstone; 2002. p. 1–67. 2. O’Rahilly R, Müller F. Human embryology and teratology. 3rd ed. New York: Wiley-Liss; 2001. 3. Ruchelli E.  Developmental anatomy and congenital anomalies of the liver, gallbladder, and extrahepatic biliary tree. In: Russo P, Ruchelli E, Piccoli D, editors. Pathology of pediatric gastrointestinal and liver disease. New York: Springer; 2004. p. 191–202. 4. Zhao R, Duncan SA.  Embryonic development of the liver. Hepatology. 2005;41:956–67. 5. Lemaigre FP.  Mechanisms of liver development: concepts for understanding liver disorders and design of novel therapies. Gastroenterology. 2009;137:62–79. 6. Zong Y, Stanger BZ. Molecular mechanisms of liver and bile duct development. Wiley Interdiscip Rev Dev Biol. 2012;1:643–55. 7. Collardeau-Frachon S, Scoazec JY.  Vascular development and differentiation during human liver organogenesis. Anat Rec (Hoboken). 2008;291:614–27. 8. Saxena R, Zucker SD, Crawford JM.  Anatomy and physiology of the liver. In: Zakim D, Boyer TD, editors. Hepatology. 4th ed. Philadelphia: Saunders; 2003. p. 3–30. 9. Ghosh ML, Emery JL.  Hypoxia and asymmetrical fibrosis of the liver in children. Gut. 1973;14:209–12. 10. Emery JL. Asymmetrical liver disease in infancy. J Pathol Bacteriol. 1955;69:219–24. 11. Timens W, Kamps WA. Hemopoiesis in human fetal and embryonic liver. Microsc Res Tech. 1991;39:387–97. 12. Naus GJ, Amann GR, Macpherson TA. Estimation of hepatic hematopoiesis in second and third trimester singleton gestations using flow cytometric light scatter analysis of archival autopsy tissue. Early Hum Dev. 1992;30:101–7. 13. Roskams T, Desmet V, Verslype C.  Development, structure and function of the liver. In: Burt AD, Portman B, Ferrell L, editors. MacSween’s pathology of the liver. 5th ed. Philadelphia: Churchill Livingstone Elsevier; 2007. p. 1–74. 14. Singer DB. Hepatic erythropoiesis in infants of diabetic mothers: a morphometric study. Pediatr Pathol. 1986;5:471–9. 15. Stallmach T, Karolyi L, Lichtlen P, Maurer M, Hebisch G, Joller H, et al. Fetuses from preeclamptic mothers show reduced hepatic erythropoiesis. Pediatr Res. 1998;43:349–54. 16. Bendon RW, Coventry S. Non-iatrogenic pathology of the preterm infant. Semin Neonatol. 2004;9:281–7. 17. Miranda RN, Omurtag K, Castellani WJ, De las Casas LE, Quintanilla NM, Kaabipour E.  Myelopoiesis in the liver of stillborns with evidence of intrauterine infection. Arch Pathol Lab Med. 2006;130:1786–91. 18. Pfisterer C, Faber R, Horn LC. Chorioamnionitis-induced changes of fetal extramedullar hematopoiesis in the second trimester of gestation. Is diagnosis from fetal autopsy possible? Virchows Arch. 2005;446:150–6.

P. Russo 19. Dimmick J, Jevon G.  Liver disease in the perinatal infant. In: Wigglesworth JS, Singer DB, editors. Textbook of fetal and perinatal pathology. 2nd ed. Oxford: Blackwell Science; 1998. p. 865–99. 20. Hutchins GF, Gollan JL. Recent developments in the pathophysiology of cholestasis. Clin Liver Dis. 2004;8:1–26. 21. Dimmick J.  Hepatobiliary system. In: Dimmick J, Kalousek D, editors. Developmental pathology of the embryo and fetus. Philadelphia: JB Lippincott; 1992. p. 545–78. 22. Sadava D, Frykman P, Harris E, Majerus D, Mustard J, Bernard B. Development of enzymes of glycolysis and gluconeogenesis in human fetal liver. Biol Neonate. 1992;62:89–95. 23. Ohtani O, Ohtani Y.  Lymph circulation in the liver. Anat Rec (Hoboken). 2008;291:643–52. 24. Naito M, Hasegawa G, Takahashi K.  Development, differen tiation, and maturation of Kupffer cells. Microsc Res Tech. 1997;39:350–64. 25. Silver MM, Valberg LS, Cutz E, Lines LD, Phillips MJ.  Hepatic morphology and iron quantitation in perinatal hemochromatosis. Comparison with a large perinatal control population, including cases with chronic liver disease. Am J Pathol. 1993;143:1312–25. 26. Spagnoli FM, Amicone L, Tripodi M, Weiss MC.  Identification of a bipotential precursor cell in hepatic cell lines derived from transgenic mice expressing cyto-met in the liver. J Cell Biol. 1998;143:1101–12. 27. Van Eyken P, Sciot R, Desmet V.  Intrahepatic bile duct development in the rat: a cytokeratin-immunohistochemical study. Lab Investig. 1988;59:52–9. 28. Desmet VJ.  Ludwig symposium on biliary disorders  – part I.  Pathogenesis of ductal plate abnormalities. Mayo Clin Proc. 1998;73:80–9. 29. Poncy A, Antoniou A, Cordi S, Pierreux CE, Jacquemin P, Lemaigre FP.  Transcription factors SOX4 and SOX9 cooperatively control development of bile ducts. Dev Biol. 2015;404:136–48. 30. Coffinier C, Gresh L, Fiette L, Tronche F, Schütz G, Babinet C, et al. Bile system morphogenesis defects and liver dysfunction upon targeted deletion of HNF1beta. Development. 2002;129:1829–38. 31. Lemaigre FP.  Development of the biliary tract. Mech Dev. 2003;120:81–7. 32. Sergi C, Adam S, Kahl P, Otto HF. The remodeling of the primitive human biliary system. Early Hum Dev. 2000;58:167–78. 33. Crawford JM. Development of the intrahepatic biliary tree. Semin Liver Dis. 2002;22:213–26. 34. Nakanuma Y, Hoso M, Sanzen T, Sasaki M.  Microstructure and development of the normal and pathologic biliary tract in humans, including blood supply. Microsc Res Tech. 1997;38:552–70. 35. Tan CE, Vijayan V.  New clues for the developing human biliary system at the porta hepatis. J Hepato-Biliary-Pancreat Surg. 2001;8:295–302. 36. Terada T.  Development of extrahepatic bile duct excluding gall bladder in human fetuses: histological, histochemical, and immunohistochemical analysis. Microsc Res Tech. 2014;77:832–40. 37. Mirkin LD, Knisely AS. Hyaline cartilage at porta hepatis in extrahepatic biliary atresia. Pediatr Pathol Lab Med. 1997;17:587–91. 38. Mani VR, Farooq MS, Soni U, Kalabin A, Rajabalan AS, Ahmed L.  Case report of ectopic liver on gallbladder serosa with a brief review of the literature. Case Rep Surg. 2016;2016:7273801.

6

Pancreas Eduardo D. Ruchelli and Tricia R. Bhatti

Introduction In contrast to the adult pancreas, the fetal pancreas does not undergo rapid postmortem autolysis and usually is relatively well preserved at autopsy, except for stillborn fetuses with prolonged retention in utero. Samples should be obtained from the head (including the duodenum), body, and tail. Evaluation of the fetal and neonatal pancreas, particularly its endocrine component, requires familiarity with the normal changes that the stromal, exocrine, and endocrine components undergo throughout gestation. The unique features of the endocrine tissue of the fetal and neonatal pancreas must be considered before drawing conclusions as to their possible causative role in either presumed or clinically established hyperinsulinism.

Embryology The pancreas develops from two primordia, a dorsal larger bud and a smaller ventral bud [1, 2]. The pancreas begins to develop on day 26, when the dorsal pancreatic bud starts to grow out of the duodenum into the dorsal mesentery just opposite the hepatic diverticulum. Over the next few days, the ventral pancreatic bud, adjacent to the hepatic diverticulum, grows into the ventral mesentery. Because of the close proximity of the ventral bud and the developing extrahepatic bile ducts, the main duct of the ventral pancreatic bud becomes connected to the proximal end of the

E. D. Ruchelli (*) ∙ T. R. Bhatti Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]; [email protected]

common bile duct by day 32. When the duodenum rotates to the right, the mouth of the common bile duct and the ventral pancreatic bud migrate posteriorly toward the dorsal mesentery. Late in the sixth week, the two pancreatic buds fuse to form the definitive pancreas. The dorsal pancreatic bud gives rise to a small part of the head of the pancreas and to its entire body and tail, whereas the ventral pancreatic bud gives rise to most of the head and the uncinate process. The duct systems of the pancreatic buds fuse to form the main pancreatic duct in 90–95% of the population. Branching of the pancreatic buds occurs differently from the classic branching of other organs, such as the developing lung [3]. Rather than expanding and folding the epithelium, the originally solid epithelial clusters of the pancreatic buds develop central lumina. These soon coalesce to generate a continuous lumen, forming an epithelial tree that drains exocrine products into the duodenum. The pancreatic ductal cells, exocrine cells, and endocrine cells in the islets of Langerhans all differentiate from the endoderm of the pancreatic buds. The acinar cells first appear as small groups of cells along the sides and tips of the ducts. They are penetrated by duct cells that become the centroacinar cells. Other duct cells elongate to form the intercalated ducts, which serve as the connection between the acini and the larger ducts. The endocrine cells are seen first during the seventh week as small clusters of cells on the outer surface of the ducts [4, 5]. They then detach from the ducts and lie in clusters in the adjacent fibrous septa. The term nesidioblastosis was initially used to describe this normal developmental process [6]. Unfortunately, over the past few decades, this term has been used erroneously in the literature to describe the pathologic changes of congenital hyperinsulinism, creating much confusion regarding the distinction between the normal features of pancreatic development and the true histologic abnormalities of congenital hyperinsulinism. A second generation of endocrine

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cells arises from centroacinar cells in the fourth month of gestation, and these cells lie individually or in small clusters in the center of the lobules. Both generations of endocrine cells continue to be present throughout gestation, and both α- and β-cells can be recognized even in the earliest developing islets.

within the lobules is proportionally decreased. At term, the connective tissue is markedly diminished, and in the normal adult pancreas, there is little connective tissue between the lobules and almost none within them. The epithelial component of the pancreas includes exocrine elements (acini and ducts) and endocrine elements (islets of Langerhans).

Histology

Exocrine Pancreas

Microscopically, the pancreas is arranged in lobules. The lobular architecture is particularly prominent during the midtrimester, because of the relatively abundant connective tissue present between the lobules (Fig.  6.1). This normal feature should not be misdiagnosed as fibrosis. As the pancreas matures, the amount of connective tissue between and

Acinar cells are arranged in a single layer surrounding a minute central lumen. They have basally situated round nuclei and apical granular eosinophilic cytoplasm. In contrast, endocrine cells have central round nuclei and pale amphophilic cytoplasm; they do not form acini (Fig. 6.2). Although these cytologic features can be appreciated at any gestational

a

b

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Fig. 6.1  Pancreatic stroma. (a) Pancreas at 16 weeks. Note the abundant connective tissue within and between lobules, which is characteristic of the midtrimester pancreas. (b) Pancreas at 32  weeks. The relative amount of connective tissue has diminished as the glandular elements increase. Intralobular connective tissue still remains visible, in contrast to the minimal amount present in the fully mature pancreas. (c)

Pancreas at 38 weeks. Lobules are more compact, with little intervening intralobular connective tissue. (d) Pancreas at 2 months postnatal age. The acini are more compact, with minimal intervening connective tissue, and the islets of Langerhans are becoming better defined on routine stains. All figures are shown at the same magnification for comparison (hematoxylin and eosin (H&E), 10×)

6 Pancreas

Fig. 6.2  Exocrine and endocrine cells. Acinar cells (AC) are clearly distinguished by their basally located nuclei and apical bright, eosinophilic cytoplasm surrounding a central small lumen, in contrast to the paler endocrine cells (EC) with central nuclei and cord-like arrangement within the islets. Pancreas at 35 weeks is shown (H&E, 20×)

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Fig. 6.3  Centroacinar cells. Prominent centroacinar cells are seen within acini on the left (arrows), characterized by their pale cytoplasm and centrally placed nuclei. They may be difficult to distinguish from endocrine cells. A distinct islet on the right (EC) is composed of endocrine cells with slightly more eosinophilic cytoplasm, but otherwise the cytologic features are similar to the centroacinar cells. Pancreas at 23 weeks is shown (H&E, 40×)

ducts (Fig. 6.4). As the ducts leave the lobules, their epithelial lining cells become slightly taller, and they become enveloped by a thicker coat of collagenous tissue. These interlobular ducts gradually enlarge as they approach the main excretory ducts.

Endocrine Pancreas

Fig. 6.4  Ducts. The intralobular ducts (center) are surrounded by little connective tissue and receive innumerable intercalated ducts (arrows) draining the acini. Pancreas at 32 weeks is shown (H&E, 20×)

age, many endocrine cells in the fetal pancreas are arranged individually or in small clusters and can be difficult to ­distinguish from exocrine cells on routine stains. Centroacinar cells are flat to cuboidal cells with pale or lightly eosinophilic cytoplasm and central oval nuclei, located between the acinar cell and the intercalated duct. Centroacinar cells usually are inconspicuous, but occasionally they can be prominent and can be mistakenly interpreted as endocrine cells (Fig. 6.3). If necessary, an immunostain for cytokeratins can be used to distinguish them from endocrine cells, which generally are not reactive for keratins. The lumen surrounded by acinar and centroacinar cells drains into the intercalated ducts, which in turn join together to form the intralobular

Although endocrine cells constitute only 1–2% of the volume of the adult pancreas, about 10% of the neonatal pancreas is composed of endocrine cells [7]. In spite of this relatively large proportion of endocrine tissue in the fetal pancreas, endocrine cells can be difficult to distinguish with certainty in routinely stained sections because so many of them occur as single cells or tiny nests rather than as the traditionally compact, larger islets (Fig.  6.5). Endocrine cells arising in the ducts are particularly difficult to identify on routine stains, and immunohistochemistry for various neuroendocrine markers is necessary in order to outline their pattern of growth (Fig. 6.6). The extrainsular endocrine cells become much less abundant later in life and constitute less than 10% of the total endocrine cell population in the adult. The quantification of endocrine tissue is complicated by the considerable variation in the concentration of islets from one lobule to the next (Figs. 6.7, 6.8, and 6.9). The size and shape of the islets can also vary significantly, as seen in Fig. 6.8. Fetal islets generally are not as compact and well circumscribed as islets in individuals who are several months old. Although most islets are sur-

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Fig. 6.5  Endocrine pancreas. (a) Endocrine cells are not readily recognized on routine H&E stain. (b) The immunostain for synaptophysin on the same section demonstrates innumerable small clusters and single

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endocrine cells. Pancreas at 19 weeks is shown (a H&E, 4×; b synaptophysin immunostain, 4×)

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Fig. 6.6  Endocrine pancreas. (a) The endocrine cells are first seen as small clusters of cells on the outer surface of the ducts. Budding and the subsequent detachment of islets from ducts can be seen in this field (nesidioblastosis). (b) The process is highlighted by immunostain for

synaptophysin. After detaching from the ducts, the endocrine cells lie in clusters in the adjacent fibrous septa (see Fig.  6.10). Pancreas at 16 weeks is shown (a H&E, 10×; b synaptophysin immunostain, 10×)

rounded by acinar elements and tend to have a centrilobular location, septal islets are not uncommon in the fetus and newborn (Fig. 6.10). These normal features of the fetal and neonatal pancreas should not be mistaken for evidence of presumed or clinically established hyperinsulinism. Islet cells with nucleomegaly (nuclei three to four times the size of their neighbors) are the most reliable evidence for congenital hyperinsulinism (Fig.  6.11). Similar cells may be seen in the pancreases of infants of diabetic mothers, Beckwith-Wiedemann syndrome, trisomy 13, and other

anomalies, but they are rarely seen in otherwise normal fetuses and newborns. They are occasionally seen in asymptomatic adults.

Other Histologic Features Small and large lymphocytic infiltrates are frequently encountered within the fetal pancreas. These infiltrates lack follicle formation and are characterized by small, mature

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Fig. 6.7  Endocrine pancreas. Note the variable density of endocrine tissue within neighboring lobules. Pancreas at 19 weeks is shown (synaptophysin immunostain, 10×)

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Fig. 6.8  Endocrine pancreas. Note the variable size and shape of the developing islets. Pancreas at 24 weeks is shown (synaptophysin immunostain, 10×)

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Fig. 6.9  Endocrine pancreas. The fetal and neonatal pancreas can contain areas with a large proportion of endocrine tissue, but this finding by itself should not be considered pathologic. Pancreas at 35  weeks is shown (synaptophysin immunostain, 4×)

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lymphocytes intermingled with acini and ducts (Fig. 6.12). Most lymphoid aggregates are seen in otherwise normal individuals and are thought to represent lymphoid heterotopias, but if such infiltrates are accompanied by a significant infiltrate of plasma cells or the destruction of parenchyma or ducts, infections such as cytomegalovirus or immune dysregulation disorders should be considered. Extramedullary hematopoiesis is also commonly encountered in the fetal pancreas, particularly during the midtrimester (Fig. 6.13). Fig. 6.10  Septal islets. (a) Some islets originating from large ducts (see Fig.  6.6) can remain in a septal location. (b) Well-defined islet within background fibrous septa. Pancreas at 36  weeks is shown (a H&E, 20×; b synaptophysin immunostain, 20×)

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Fig. 6.11  Endocrine cell nucleomegaly. β-Cells with enlarged nuclei (arrows) are frequently seen in various types of congenital hyperinsulinism, but they are extremely rare among normal fetuses and newborns (H&E, 40×)

Fig. 6.13  Hematopoiesis. Although hematopoiesis is never as prominent in the pancreas as in the liver, it is a fairly constant feature during the second and early third trimesters. Pancreas at 24 weeks is shown (H&E, 20×)

E. D. Ruchelli and T. R. Bhatti

Fig. 6.12  Lymphoid tissue. Lymphocytic aggregates around ducts and also extending between acini are common in the preterm pancreas and have no pathologic significance. They become less frequent toward term. Pancreas at 23 weeks is shown (H&E, 10×)

Fig. 6.14  Nodular splenic heterotopia. A splenic nodule (Spl) is completely surrounded by pancreatic parenchyma in the tail of the pancreas (H&E, 2×)

Nodular splenic heterotopias are occasionally seen in the tail of the pancreas (Fig. 6.14). They are usually incidental findings, but if they are extensive or accompanied by splenopancreatic fusion, they may be part of multiple congenital anomalies such as trisomy 13. Pancreatic heterotopias are found in locations such as the site of esophageal atresia, in the wall of the stomach, on the antimesenteric border of the small intestine, or in the wall of a Meckel’s diverticulum (Fig. 6.15).

Fig. 6.15  Small-bowel pancreatic heterotopia. A heterotopic pancreatic nodule within the wall of the small bowel is shown. This is one of the most common locations of heterotopic pancreatic tissue (H&E, 4×)

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References 1. Moore KL, Persaud TV, Torchia MG. The developing human: clinically oriented embryology. 10th ed. Philadelphia: Saunders; 2015. 2. Bock P, Abdel-Moneim M, Egerbacher M.  Development of pancreas. Microsc Res Tech. 1997;37:374–83. 3. Schoenwolf GC, Bleyl SB, Brauer PR, Francis-West PH. Larsen’s human embryology. 4th ed. Philadelphia: Churchill Livingstone Elsevier; 2009.

97 4. Peters J, Jurgensen A, Kloppel G.  Ontogeny, differentiation and growth of the endocrine pancreas. Virchows Arch. 2000;436:527–38. 5. Hill DJ. Development of the endocrine pancreas. Rev Endocr Metab Disord. 2005;6:229–38. 6. Jaffe R, Hashida Y, Yunis EJ. The endocrine pancreas of the neonate and infant. Perspect Pediatr Pathol. 1982;7:137–65. 7. Rahier J, Wallon J, Henquin JC.  Cell populations in the endocrine pancreas of human neonates and infants. Diabetologia. 1981;20:540–6.

7

Salivary Glands Eduardo D. Ruchelli

Introduction The three major salivary glands (parotid, sublingual, and submandibular) can be examined at autopsy during dissection of the neck to evaluate the presence of developmental anomalies or cytomegalovirus infection. The submandibular glands are easier to dissect at autopsy than the other two glands and can be routinely sampled before removal of the neck block by exposing the tissues between the angle of the mandible and the base of the tongue. Thus, this chapter focuses on the development and histology of the submandibular gland.

Embryology During the sixth and seventh weeks, the salivary glands begin as solid epithelial buds from the primordial oral cavity [1]. Five stages of development have been described in the mouse [2]. In the first stage, known as the pre-bud stage, the oral epithelium thickens; it then protrudes into the underlying mesenchyme (bud stage). The bud then undergoes branching (pseudoglandular stage). These branches are initially solid but later develop a lumen and become ducts (canalicular stage). Subsequently, the terminal buds are also hollowed out to form the acini (terminal bud stage). The parotid glands are the first to appear, early in the sixth week; they are derived from the ectoderm of the primitive oral cavity. The submandibular glands appear late in the sixth week and probably develop from endodermal buds in the floor of the stomodeum, although their exact derivation remains somewhat obscure [3]. Acini begin to form at 12 weeks, and secretory activity begins at 16 weeks. The sublingual glands E. D. Ruchelli (*) Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]

appear in the eighth week, approximately 2 weeks later than the other salivary glands. Unlike rat or mouse salivary glands, which differentiate predominantly in postnatal life, human salivary glands appear to achieve complete maturation by the end of gestation [3]. However, the full details of the development of human salivary glands have yet to be elucidated.

Histology The mature salivary glands have a secretory lobular component (acini, intercalated ducts, and striated ducts) and an excretory component (interlobular ducts) [4]. The acini of the submandibular glands are mixed, including both mucous cells and serous cells. Mucous cells have abundant clear cytoplasm, whereas serous cells have basophilic cytoplasm and are arranged in a crescent-shaped formation around the mucous cells. The intercalated duct lies between the acinus and the striated duct. It is lined by a single layer of cuboidal epithelium, and it is typically very short in the submandibular gland. In contrast, the striated duct is longer, and its single layer of columnar epithelium with eosinophilic cytoplasm and characteristic striations stands out on routine sections. The interlobular ducts are lined by pseudostratified epithelium with sparse goblet cells. A layer of myoepithelial cells surrounds the acini and intercalated ducts, but these cells are difficult to identify on routine sections. The histology of the acini and ducts changes during fetal life, as described in the following section. During the early midtrimester, the submandibular gland architecture is already lobular, but abundant loose mesenchyme separates the epithelial elements and constitutes the main tissue component (Fig. 7.1). The epithelial component consists of tubules and terminal buds with a two-cell layer structure (Figs. 7.2, 7.3, and 7.4). Acini are primitive at this time, but myoepithelial cell differentiation can be demonstrated by immunohistochemistry during the early midtrimester (see Figs.  7.2b and 7.4b). Ducts and terminal units typically contain secretions within the lumens in the second

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Fig. 7.1  Submandibular gland at 16  weeks. Abundant loose mesenchyme constitutes the major tissue component at this age. The branching epithelial cords have already developed lumina throughout, forming tubular structures. Larger vessels are associated with the more proximal ducts (hematoxylin and eosin (H&E), 4×)

E. D. Ruchelli

Fig. 7.3  Submandibular gland at 19 weeks. The gland has a distinct lobular organization. Interlobular and intralobular tubular structures continue to have a two-cell epithelial layer, and secretions are present in their lumens (H&E, 10×)

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a

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Fig. 7.2  Submandibular gland at 16  weeks. (a) Proximal and distal tubular structures are all lined by a two-cell epithelial layer. (b) The distal tubular structures, which are the precursors of the acini, demonstrate myoepithelial cell differentiation, as evidenced by expression of smooth muscle actin along the basal cell layer (a H&E, 10×; b smooth muscle actin immunostain, 10×)

Fig. 7.4  Submandibular gland at 19  weeks. (a) The proximal ducts have a two-cell structure with low columnar inner cells and cuboidal outer cells (arrows). The distal tubular structures have a cuboidal inner cell layer and an outer cuboidal or flattened cell layer (arrowheads). (b) Smooth muscle expression indicative of myoepithelial cell differentiation is seen in the outer layer of the distal tubular structures but not in the proximal ducts (a H&E, 20×; b smooth muscle actin immunostain, 20×)

7  Salivary Glands

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trimester (see Figs. 7.3 and 7.4). Near the end of the midtrimester, the striated ducts become more distinct as the cytoplasm grows larger and eosinophilic, whereas the intercalated ducts and primitive acini remain smaller, one-cell to two-cell layered tubular structures (Fig. 7.5). During the third trimester, as the gland matures the epithelial structures become more numerous and crowded, with a relative decrease in intervening stroma (Fig. 7.6). The interlobular ducts (excretory ducts) thicken and adopt a multilayer-­ cell structure (Fig.  7.7). At the same time, striated ducts elongate, becoming more prominent, and mucous differentia-

tion becomes evident in the acini (Figs. 7.8 and 7.9). Although we have observed mucous acini during the third trimester, others consider the development of mucous acini a postnatal event [5, 6]. In contrast to the parotid and sublingual salivary glands, which have short striated ducts, the submandibular gland has long striated ducts that become very distinct during the third trimester (Fig. 7.10). The two-cell layer structure of these ducts during the second trimester gradually changes to a single layer of columnar cells with occasional cuboidal cells in the basal area (Fig. 7.11). At term, the acini display complete differentiation (Fig. 7.12) [3].

Fig. 7.5  Submandibular gland at 23 weeks. The intralobular striated ducts (SD) become more distinct as the cytoplasm becomes more abundant and more eosinophilic. The columnar cells of the interlobular ducts (ILD) become taller, whereas the intercalated ducts and primitive acini (AC) remain smaller, one-cell to two-cell layered tubular structures (H&E, 10×)

Fig. 7.7  Submandibular gland at 27  weeks. The interlobular ducts thicken and develop a multilayer-cell structure with scattered goblet cells (H&E, 40×)

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b

Fig. 7.6 (a) Submandibular gland at 27 weeks. The lobular architecture becomes more pronounced as the density of the interlobular connective tissue increases. Loose mesenchyme is still focally present within the lobules, separating the epithelial elements (compare with Fig. 7.1). (b) Submandibular gland at 32 weeks. During the third tri-

c

mester, the gland becomes progressively more compact as the result of an increase in the number of acini and a relative decrease in stroma. (c) Submandibular gland at 41 weeks. The lobules have a compact appearance, whereas the interlobular connective tissue is dense and fibrous (a–c H&E, 4×)

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Fig. 7.8  Submandibular gland at 27 weeks. Striated ducts (SD) elongate and the outer cell layer becomes discontinuous. Long striated ducts are characteristic of the submandibular gland, in contrast to the shorter striated ducts of the other major salivary glands. Note that relatively abundant, loose mesenchyme is still present within the lobules at this age (H&E, 20×)

Fig. 7.9  Submandibular gland at 27  weeks. Acini show evidence of mucous differentiation (MC), which continues to progress during the third trimester (H&E, 40×)

Fig. 7.10  Submandibular gland at 35  weeks. Acini contain mucous cells (MC). The striated ducts (SD) are long and very distinct, whereas the intercalated ducts are short and difficult to visualize in the submandibular gland (H&E, 20×)

Fig. 7.11  Submandibular gland at 41 weeks. The striated ducts (SD) lose the two-cell layer structure seen earlier in gestation, and the mature wall consists of a single layer of columnar cells with eosinophilic cytoplasm and barely visible basal striations. The intercalated duct (ICD) is very short as it enters the acinus (H&E, 40×)

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References 1. Moore KL, Persaud TV, Torchia MG. The developing human: clinically oriented embryology. 10th ed. Philadelphia: Saunders; 2015. 2. Tucker AS.  Salivary gland development. Semin Cell Dev Biol. 2007;18:237–44. 3. Chi JG.  Prenatal development of human major salivary glands. Histological and immunohistochemical characteristics with reference to adult and neoplastic salivary glands. J Korean Med Sci. 1996;11:203–16. 4. Martinez-Madrigal F, Bosq J, Casiraghi O. Major salivary glands. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 477–502. 5. el-Mohandes EA, Botros KG, Bondok AA.  Prenatal develop ment of the human submandibular gland. Acta Anat (Basel). 1987;130:213–8. 6. Scott J. Qualitative and quantitative changes in the histology of the human submandibular salivary gland during post natal growth. J Biol Buccale. 1979;7:341–52. Fig. 7.12  Submandibular gland at 41  weeks. Mucous cells (MC) become focally more abundant in the acini and are surrounded by serous cells (SC) (H&E, 20×)

Part IV Genitourinary Tract

Early Embryology of the Genitourinary Tract Dale S. Huff The embryology of the genital and urinary tracts are inextricably interrelated and will therefore be described together. The development of the genitourinary tract can be divided into three phases. First, the primitive genitourinary tract develops in the early part of the fifth postfertilization week. Second, in the middle of the fifth week even before the first phase is completed, the definitive urinary tract and the bipotential genital tract begin to be superimposed upon the primitive genitourinary tract. Before the second phase is completed, the third phase, sexual differentiation of the bipotential genitourinary tract, begins when the bipotential genital tract begins to differentiate into either a male or female definitive genital tract beginning in the sixth week, a process that is not completed until the 12th week. The postfertilization gestational ages are assigned following the revised data of O’Rahilly and Muller [1] (Appendix 2).

Primitive Genitourinary Tract General The primitive genitourinary tract is composed of the paired mesonephric bodies, the paired mesonephric ducts, and the midline cloaca. The cloaca first appears in the middle of the fourth week when the caudal curl of the embryo and the caudal intestinal portal define the hindgut from which the cloaca develops. The urorectal septum will divide the cloaca into a ventral half, which will become the urogenital sinus, and a

dorsal half, which will become the rectum. The urogenital sinus will become the bladder and urethra, and, in the female, the introitus. The mesonephric bodies are composed of multiple mesonephric nephrons that are arranged in single file from rostral to caudal. Each mesonephric nephron connects to the mesonephric duct, also called the primary excretory duct, which connects to the ventral or urogenital part of the cloaca. The mesonephric nephrons produce urine, which drains through the collecting ducts into the mesonephric duct and into the cloaca. A mesonephric body is also called the mesonephros. The mesonephros is the functional part of the primitive genitourinary tract, and its development is described next.

Mesonephros and Mesonephric Duct The mesonephros develops from the intermediate mesoderm. Its development is simultaneous with the rostral to caudal wave of differentiation of the somites from the paraxial mesoderm [2] and the differentiation of the overlying ectodermal ring within the ectoderm. In the middle of the fourth week, the first somite appears in the paraxial mesoderm of the occipital region. Every few hours another pair of somites differentiates caudally to the previous pair. Simultaneously, the adjacent intermediate mesoderm differentiates into the nephrogenic cord. By the end of the fourth week, the occipital and upper cervical somites appear, and the nephrogenic cord adjacent to these somites differentiates into primitive nephric structures called nephrotomes, which rapidly disappear. The nephrotomes are somewhat analogous to the

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pronephros of lower vertebrates, but a true pronephros does not exist in humans [3]. By the beginning of the fifth week, the middle and lower cervical somites appear. The mesonephric duct arises from the dorsolateral border of the nephrogenic cord adjacent to somite C4 at the level of the upper limb bud. The first mesonephric nephron immediately differentiates within the nephrogenic cord adjacent to C6. Mesonephric nephrons are analogous to metanephric nephrons, and each is attached to the mesonephric duct by its connecting tubule [4]. The mesonephric duct separates from the nephrogenic cord and grows caudally between the dorsolateral border of the nephrogenic cord and the ectoderm by elongation of its proliferating blind tip. Every few hours a new somite differentiates caudal to its predecessor [5], and new mesonephric nephrons differentiate caudally to their predecessors and attach to the independently elongating mesonephric duct. The caudal growth of the mesonephric duct is faster than the rostral to caudal differentiation of mesonephric nephrons. As the duct elongates caudally, it is attached to the ectoderm at multiple sites. It follows the caudal curl of the embryo to the midlumbar region at approximately the level of L2–L3, where its most caudal attachment to the ectoderm is located. From this point of attachment, it curves ventromedially away from the ectoderm and attaches to the urogenital portion of the cloaca, at the level of L4–L5, before the caudal growth of the mesonephros is completed. The caudal end of the mesonephros reaches

Nasal disc Optic cup

only to the level of L2–L3, the site of the curve of the mesonephric duct and its attachment to the ectoderm. The mesonephric nephrons are not metameric, and there are more nephrons than somites. As new nephrons are added caudally, the rostral nephrons and mesonephric duct degenerate. A total of over 80 nephrons develop, but, due to the simultaneous degeneration of the rostral nephrons, when the last nephron differentiates and attaches to the curve of the mesonephric duct at the level of L2–L3, only approximately 32 nephrons remain. The mesonephric duct is attached to the mesonephros by a common investing mesenchyme and the attachments of the connecting tubules of the mesonephric nephrons to the mesonephric duct. The attachment of the caudal end of the mesonephros to the curve of the mesonephric duct and the attachment of the curve of the mesonephric duct to the ectoderm are the beginnings of the development of the inguinal canal and the caudal gonadal ligaments. This coordinated rostral to caudal differentiation of somites, mesonephros, and mesonephric duct is so rapid that within 1–2  days, early in the middle of the fifth week, all of the thoracic and lumbar somites have been added [6], the development of the mesonephric bodies and mesonephric duct is complete, the mesonephric duct has attached to the cloaca, and the development of the inguinal canal and caudal gonadal ligaments has begun [2] (see Figs. IV.1 and IV.2). The continued development of the inguinal region and the caudal gonadal ligaments is described below.

Otocyst Pharyngeal arches Ectodermal ring (shaded)

Anterior neuropore

Upper limb

Allantois Cloacal membrane Midlumbar level

Level of C4 Mesonephric duct Mesonephric vesicles Mammary crest

Lower limb

Fig. IV.1  Left lateral view of the embryo at the beginning of the fifth postfertilization week. The mesonephros extends from the midcervical to the midlumbar area and follows the caudal curl of the embryo. The caudal development of the mesonephros is complete, and the caudal end of the mesonephros is at the midlumbar area. The mesonephric duct curves ventromedially around the distal end of the mesonephros at the midlumbar area where it is attached to the ectodermal ring at the future site of the internal inguinal ring and canal. This is the first curve of the mesonephric duct. The

common mesenchyme, which invests the mesonephros and mesonephric duct, is not shown in this figure so that the individual mesonephric nephrons can be seen. The lower limb bud has appeared within the ectodermal ring at the midlumbar area in continuity with the cloacal membrane. The precursors of the caudal attachments of the gonads and gonadal ducts to the inguinal canal at the midlumbar area are already established. Several developmental fields within the ectodermal ring are depicted (shaded area)

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107

a

b

Midlumbar level (most caudal area of attachment between mesonephric duct and ectodermal ring

* *

Mesonephric duct

Tail gut

Cloaca

Cloacal membrane

Lower limb

Cloaca Allantois

Hind gut

Mesonephric duct

*

*

Caudal end of mesonephric vesicles

Mesonephric vesicles Most caudal area of attachment between mesonephric duct and ectodermal ring

Fig. IV.2  Closer views of the caudal end of the primitive genitourinary tract early in the fifth week. (a) Left lateral view. The mesonephros follows the caudal curl of the embryo. At the midlumbar area, the first curve of the mesonephric duct is attached to the overlying ectodermal ring (not shown) between the asterisks at the future site of the internal inguinal ring and

canal. (b) Dorsal view of the caudal curl. The first curve of the mesonephric seen on end is attached the ectodermal ring at the asterisks at the future site of the internal inguinal ring and canal at the origin of the lower limb. The ureteric bud will arise from the segment of the mesonephric duct between its attachment to the ectodermal ring and its entrance into the cloaca

Ectodermal Ring

The mesonephric duct is in contact with the ectodermal ring at various points from the midcervical to the midlumbar regions. Its attachment to the ectodermal ring at the level of C4 through C8, the site of the upper limb bud, is so close that some believe the duct arises in whole or in part from the epithelium of that part of the ectodermal ring rather than entirely from the nephrogenic cord [9]. The most caudal attachment is the one described above, between the first curve of the mesonephric duct (the future tail of the epididymis) and the ectodermal ring in the midlumbar region at the site of the future inguinal ring. It is possible that the epithelial–mesenchymal interactions between the ectodermal ring and the first curve of the mesonephric duct at this site induce the formation of a developmental field within which the gubernaculum, inguinal canal, and gonadal ligament develop.

The ectodermal ring is a placodal strip of thickened ectoderm that encircles the embryo in the boundary between the dorsal and ventral halves of the body. It first appears during the end of the fourth postfertilization week in the face and extends caudally on both sides of the embryo like an inverted U to cover the lateral head, neck, thorax, abdomen, and the inguinal areas at the same time as the somites, mesonephros, and mesonephric duct are differentiating [7]. The two limbs of the inverted U meet caudally in the ventral midline over the cloacal membrane to complete the ring on the second day of the fifth week at the same time that the development of the somites, mesonephros, and mesonephric duct reaches the midlumbar region. The ectodermal ring is in intimate contact with the underlying mesenchyme of the intermediate mesoderm, mesonephros, and especially the mesonephric duct. The rich epithelial–mesenchymal connections between the ectodermal ring and the underlying mesoderm are the histologic basis for molecular epithelial–mesenchymal interactions, which are features characteristic of developmental fields. The ectodermal ring is thought to play an important role in directing the growth and development of the entire embryo of all vertebrates, both nonmammalian and mammalian [8]. Developmental fields arise within the ectodermal ring and the underlying specialized mesoderm for the nose, eyes, ears, pharyngeal apparatus, upper limbs, mammary glands, lower limbs, inguinal region, external genitalia, and anus (see Fig. IV.1).

I nguinal Region and Caudal Gonadal Ligaments The inguinal region is defined as the junction between the lower limb and the abdomen. The apical ectodermal ridge of the lower limb bud arises within the ectodermal ring in the midlumbar area early in the fifth week. The caudal end of the lower limb bud is adjacent to the cloacal membrane. The appearance of the lower limb bud establishes the embryonic inguinal region. The internal structures of the inguinal area are the inguinal canal and the bipotential caudal gonadal ligaments. These structures develop rapidly beginning in

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the middle of the fifth week when the mesoderm of the body wall grows between the ectodermal ring and the mesonephric duct, breaking the epithelial–mesenchymal attachments between the ectodermal ring and the mesonephric duct, beginning rostrally and progressing caudally. The last attachment to be broken is the one between the first curve of the mesonephric duct and the ectodermal ring at the level of L2–L3. At the site of this broken attachment, a placode of primitive mesenchyme forms a pyramidal nodule within the mesoderm of the anterior abdominal wall [10]. The tip of the nodule protrudes into the peritoneal cavity. This mesenchymal nodule is the inguinal crest, and it is located at the site of the inguinal canal. The mesenchyme around the first curve of the mesonephric duct attaches to the tip of the inguinal crest (see Fig. IV.3). The caudal end of the mesonephros, as described above, is attached to the first curve of the mesonephric duct by the common mesenchyme investing the mesonephros and mesonephric duct. The mesenchymal attachment of the caudal end of the mesonephros to the first curve of the mesonephric duct is the bipotential gonadal ligament. The inguinal crest becomes the gubernaculum. The abdominal wall develops around the gubernaculum forming the inguinal canal. The first curve of the mesonephric duct is therefore attached to the gubernaculum in the inguinal canal and to the caudal end of the mesonephros by the gonadal ligament. The gonad will replace the mesonephros in the near future (see Fig. IV.4), and, during this process, the attachment of the gonadal ligament to the caudal end of the mesonephros gradually shifts to the caudal end of the gonad. Tail gut

a

Mesonephric body Gonad

Epigenitalis

Paragenitalis

*

First curve L3

Fig. IV.4  Left lateral view of the mesonephros and gonad at approximately the middle of the sixth postfertilization week. The individual mesonephric nephrons are hidden by the investing mesenchyme of the mesonephros and mesonephric duct. The enlarging gonad gradually replaces the regressing mesonephros. The mesonephros extends from approximately T1 to L3. The gonad extends from approximately T3 to T12. The caudal ends of the mesonephros and gonad are not separated, and the caudal end of the gonad is indirectly attached to the first curve of the mesonephric duct through its continuity with the caudal end of the mesonephros. The mesonephros is divided into three parts: the part cranial to the gonad, the pregenitalis; the part adjacent to the gonad, the epigenitalis; and the part caudal to the gonad, the paragenitalis. The embryonic remnants of the mesonephros and mesonephric duct are named according to this nomenclature. The asterisk is at the site of the developing gubernaculum, inguinal ring, and inguinal canal

b

Cloaca Cloacal membrane

Common excretory duct Ureter (ureteric bud)

Allantois Hind gut

Kidney Inguinal crest Mesonephric duct Mesonephric vesicles

Pregenitalis

Mesonephric duct

*

Fig. IV.3  View of the caudal primitive genitourinary tract in the middle of the fifth week approximately 2 days later than Fig. IV.2. (a) Left lateral view. (b) Dorsal view of the caudal curl. The ureteric bud and primitive kidney have appeared. The first curve of the mesonephric duct has separated from the ectodermal ring. The inguinal crest (asterisks) has

Cloaca

Ureter (ureteric bud)

Lower limb

Caudal mesonephric vesicles Mesonephric duct

Common excretory duct

*

*

Inguinal crest Kidney

appeared in the future site of the inguinal ring and canal to which the first curve of the mesonephric duct is now attached. The segment of the mesonephric duct between the inguinal crest and ureteric bud will become the vas deferens. The segment between the ureteric bud and the entrance to the cloaca is the common excretory duct

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In the male the first curve of the mesonephric duct becomes the tail of the epididymis, the gonadal ligament becomes the testicular ligament, and the gubernaculum remains as the gubernaculum. In the female the paramesonephric duct develops beside the mesonephric duct. The first curve of the paramesonephric duct develops within the gonadal ligament. As the mesonephric duct degenerates, the gonadal ligament remains attached to the paramesonephric duct and becomes the ovarian ligament. The gubernaculum becomes the round ligament, and the inguinal canal becomes the canal of Nuck. Thus, by the middle of the fifth week, the basic blue print of the primitive urinary tract is laid down [11, 12]. The mesonephric bodies are the largest visceral organs of the embryo. They extend from the midcervical to the midlumbar region as large raised, parallel, parasagittal ridges protruding from the posterior abdominal wall near the midline (see Fig. IV.5).

Head

They follow the line of the caudal curl of the embryo. The precursors of the caudal gonadal ligaments fix the caudal ends of the mesonephric bodies and the mesonephric ducts to the future inguinal ring and inguinal canal, a relationship that will never vary. The lower limb bud is present. The mesonephric bodies will begin to produce urine during the sixth week but will then degenerate in a rostral to caudal wave. The bodies will be replaced by the gonads and will disappear by the end of the eighth week [13]. At the middle of the fifth week, this primitive urinary system is ready for the beginning of the development of the bipotential genital tract and the definitive urinary tract.

Bipotential Genital Tract The bipotential genital tract is superimposed upon the primitive urogenital tract. It begins with development of the bipotential gonad on the ventromedial surface of the mesonephros during the fifth week and ends with completion of the paramesonephric duct at the end of the eighth week. The differentiation of the bipotential gonads into testes and ovaries begins before the development of the paramesonephric ducts is completed.

C4 Upper limb

Mesonephric duct Mesonephric body

Lower limb

*

*

L3 Cloaca Tail

Fig. IV.5  Mesonephric bodies in the early to mid-fifth week. The investing mesenchyme of the mesonephros and mesonephric duct is included in this figure and hides the underlying mesonephric nephrons. The mesonephric bodies are the largest organs in the embryo. They extend from the midcervical to the midlumbar area. The first curve of the mesonephric duct, the future tail of the epididymis, and indirectly the caudal end of the mesonephric bodies are attached to the inguinal crest, indicated by the asterisk, at the future site of the gubernaculum, internal inguinal ring, and inguinal canal

Bipotential Gonads The development of the gonads begins with the primordial germ cells. The primordial germ cells may be first identifiable in the inner cell mass of the blastocyst in the first postfertilization week [1, 14]. There is good evidence that they are in the amniotic ectoderm at the junction of the amnion with the epiblast at the caudal end of the bilaminar disc early in the second week [15]. During the dissolution of the primary yolk sac and the formation of the definitive yolk sac, they migrate by ameboid action through the extraembryonic mesenchyme of the body stalk and reach the endoderm of the caudal yolk sac near its junction with the allantois in the third week. During the fourth week, they continue to migrate through the wall and mesentery of the proximal hindgut and toward the adjacent ventromedial side of the mesonephric bodies [16]. In the middle of the fifth week, shortly after the appearance of the ureteric bud and crista inguinalis, the primordial germ cells enter the ventromedial border of the mesonephric bodies. At the same time, the mesonephric mesoderm proliferates, differenti-

110

ates into gonadal mesoderm, and induces the coelomic epithelium to proliferate and form a thickened layer. The thick coelomic epithelium and the enlarging underlying nest of differentiating mesoderm form the early gonadal ridge on the ventromedial surface of the mesonephric body [17]. The subsequent events are somewhat controversial. One view proposes that primordial germ cells and coelomic epithelium migrate into the differentiating mesoderm, and that these three components form the gonadal blastema [15]. The coelomic epithelium serves as a scaffolding around which the other components of the gonadal blastema are organized. The mesodermal tissue forms a network of poorly defined, irregular gonadal cords in the deep portion of the gonad near its border with the mesonephric body. The coelomic epithelium, primordial germ cells, and local mesenchyme lie between the cords. A recent study utilizing serial semithin sections of Epon-embedded gonads from human embryos and fetuses proposes that gonadal cords arise from mesonephric glomerular and tubular epithelium. These cords grow though the mesorchium into the hilum of the gonad and, hence, into the cortex. These primitive mesonephric cords never intermingle with coelomic epithelium and give rise to the rete and definitive sex cords [18]. The gonadal blastema, whatever its origin, rapidly enlarges and forms a long oval gonad with a broad attachment to the ventromedial surface of the mesonephric body [11]. It does not reach either the upper or lower pole of the mesonephric body (see Fig. IV.4). At the junction between the gonad and the mesonephric body, the gonadal blastema pushes deeply into the mesonephros. By the middle of the sixth week, the bipotential period of gonadal development ends. The mesonephros extends from the midcervical to the midlumbar regions, while the gonad extends from approximately T1 to T10 (see Fig. IV.4). A groove forms along the length of the junction between the gonad and mesonephros, and the two begin to separate. This separation continues during the following period of testicular and ovarian differentiation.

Bipotential Mesonephric and Paramesonephric Ducts The development of the bipotential mesonephric (Wolffian) ducts is completed early in the fifth week when they reach the cloaca. Subsequent

Part IV  Genitourinary Tract

remodeling occurs [2]. In the seventh week, the pelvis begins to form medial to the developing inguinal ring and gubernaculum. As the developing urogenital sinus sinks into the deepening pelvis, the mesonephric duct curves caudally over the lip of the pelvic inlet and into the pelvis. This is the second mesonephric curve [6]. The two curves divide the mesonephric duct into three segments. First, there is a rostral vertical segment attached to the dorsolateral border of the mesonephros extending to the first curve at the caudal end of the mesonephros. This will become the epididymis. The first curve will become the tail of the epididymis. The second is a horizontal segment extending from the first to the second curve, and the third is a vertical segment from the second curve to the entrance of the duct into the urogenital sinus, which will become the vas deferens. The bipotential paramesonephric (Müllerian) ducts begin in the mid to late sixth postfertilization week as invaginations of thickened areas of the coelomic epithelium at the rostral ends of the mesonephric bodies and are probably under the inductive influence of the mesonephric duct [6]. They grow caudally along the lateral margins of the mesonephric ducts with which they are in contact. The paramesonephric ducts may arise in part from the mesonephric ducts. By the end of the seventh week, the paramesonephric ducts reach the caudal ends of the mesonephric bodies and as they follow the first curve of the mesonephric ducts medially under the caudal end of the mesonephric bodies they cross over the ventral surfaces of the mesonephric ducts and come to lie medial to the mesonephric ducts [12]. In the female the paramesonephric duct replaces the mesonephric duct and assumes its caudal attachments. At the start of the eighth week, the paramesonephric ducts meet in the midline and follow the second curve of the mesonephric ducts caudally into the developing pelvis. Because of the crossing over, the paramesonephric ducts are medial to the mesonephric ducts. They fuse to form a single solid duct. By the end of the eighth week, the solid blind end of the fused paramesonephric ducts meets the mesenchyme of the posterior wall of urethral part of the vesicle urethral canal at the site of the sinual (Müllerian) tubercle between the openings of the mesonephric ducts (see Fig. IV.6). This ends the period of the bipotential genital ducts. Sexual differentiation of the ducts will begin shortly with the regression of the paramesonephric duct in the male and the mesonephric duct in female.

Part IV  Genitourinary Tract

111

Epigenitalis

G

G

Paragenitalis

*

B

*

Ureteral orifice Bladder outlet Urethra Müllerian tubercle

Urogenital sinus, pelvic part

Fig. IV.6  Bipotential genitourinary tract in the eighth week. The gonads (G) have replaced the mesonephric bodies. The mesonephric (red) and paramesonephric ducts (blue) are parallel, curve medially under the gonad (the first curve is at the asterisks), curve caudally (the second curve is at the arrows) over the pelvic inlet into the pelvis and fuse to the dorsal wall of the urogenital sinus at the Müllerian tubercle. The paramesonephric ducts cross in front of the mesonephric ducts at the first curve and fuse in the pelvic midline medial to the mesonephric ducts. The caudal gonadal ligaments and inguinal canal develop at the asterisks. The point of fusion of the mesonephric and paramesonephric ducts with the urogenital sinus at the Müllerian tubercle marks the division of the urogenital sinus into the vesicle urethral canal rostrally and the definitive urogenital sinus caudally. The vesicle urethral canal becomes the bladder and urethra. In the male the urethra from the bladder outlet to the rostral

edge of the Müllerian tubercle becomes the proximal prostatic urethra, and the Müllerian tubercle becomes the verumontanum. In the female the urethra from the bladder outlet to the rostral edge of the Müllerian tubercle greatly elongates and forms the entire length of the urethra except for its distal end, and in the process the Müllerian tubercle migrates to the distal end of the urethra and becomes the hymen. The mesonephric nephrons have all degenerated except for the remnants labeled epigenitalis and paragenitalis in this figure. In the male, the epigenitalis becomes the efferent ductules and the paragenitalis persist as embryonic remnants in the distal spermatic cord. In the female, the epigenitalis remnants become the epoophoron along the hilum of the ovary, and the paragenitalis remnants become the paroophoron between the medial end of the ovary and the uterus. B—bladder

 ate of the Paramesonephric Duct F in the Male

the testis as the appendix testis. The caudal portion often persists as the prostatic utricle at the verumontanum.

In the 10th week, the Sertoli cells secrete Müllerianinhibiting substance, which diffuses along the mesonephric duct and causes the regression of the ipsilateral paramesonephric duct beginning at the site at which the paramesonephric duct crosses in front of the mesonephric duct. The regression proceeds cranially and caudally from this point. The action of Müllerian inhibitory substance is local, not systemic, and does not effect the opposite para mesonephric duct. The regression of the duct cranial to the crossover point is complete except that a remnant may remain attached to the upper pole of

References 1. O’Rahilly R, Muller F. Human embryology & teratology. 3rd ed. New York: Wiley-Liss; 2001. 2. Felix W.  The development of the urinogenital organs. In: Keibel F, Mall FP, editors. Human embryology. Philadelphia: JB Lippincott; 1912. p. 752–979. 3. Torrey TW. The early development of the human nephros. Contrib Embryol. 1954;25:199–240. 4. Shikinami J.  Detailed form of the Wolffian body in human embryos of the first eight weeks. Contrib Embryol. 1926;28:49–61.

112 5. O’Rahilly R, Muller F. Developmental stages in human embryos. Washington, DC: Carnegie Institution of Washington; 1987. 6. O’Rahilly R, Muecke EC.  The timing and sequence of events in the development of the human urinary system during the embryonic period proper. Z Anat Entwickl Gesch. 1972;138:99–109. 7. O’Rahilly R, Muller F. The origin of the ectodermal ring in staged human embryos of the first 5 weeks. Acta Anat. 1985;122:145–57. 8. Hughes ESR. The development of the mammary gland. Ann R Coll Surg. 1950;122:3–23. 9. Jirásek JE. Development of the genital system and male pseudohermaphroditism. Baltimore: The Johns Hopkins Press; 1971. 10. Barteczko KJ, Jacob MI. The testicular descent in human. Adv Anat Embryol Cell Biol. 1999;156:1–98.

Part IV  Genitourinary Tract 11. Steding G.  The anatomy of the human embryo. Basel: Karger; 2009. 12. Jirásek JE.  An atlas of human prenatal morphogenesis. Boston: Martinus Nijhoff; 1983. 13. Hamilton WJ, Mossmann HW. Human embryology. 4th ed. Baltimore: William & Wilkins; 1972. 14. Hertig AT.  Human trophoblast. Springfield: Charles C. Thomas; 1968. 15. Jirásek JE.  An atlas of human prenatal developmental mechanics. London/New York: Taylor & Francis; 2004. 16. Witschi E.  Migration of the germ cells from the yolk sac to the primitive gonadal fields. Contrib Embryol. 1948;32:67–80. 17. van Wagenen G, Simpson ME. Embryology of the ovary and testis. New Haven: Yale University Press; 1965. 18. Satoh M. Histogenesis and organogenesis of the gonad in human embryos. J Anat. 1991;177:85–107.

8

Kidney Eduardo D. Ruchelli and Dale S. Huff

Introduction The histological appearance of the fetal kidney is significantly different from that of the adult kidney, mostly because it continues to develop almost until the end of gestation, when the formation of nephrons (nephrogenesis) comes to an end. Furthermore, the neonatal kidney is still immature, and significant changes occur as the kidney continues to grow and mature during infancy and childhood. A lack of familiarity with these differences may lead to mistakenly considering normal findings as pathologic ones, or to overlooking abnormalities of renal development, such as impaired nephrogenesis with reduced number of nephrons (as seen in intrauterine growth restriction) or abnormal tubular differentiation (as seen in renal tubular dysgenesis). The aim of this chapter is to review the dramatic changes in renal development in the fetal and neonatal period and highlight the important differences at various gestational ages.

Embryology Early in the fifth postfertilization week, the sacral intermediate mesoderm differentiates into the sacral nephrogenic cord, which separates completely from the mesonephros and differentiates into the metanephric blastema. The ureteric bud arises from the dorsomedial aspect of that segment of the mesonephric duct between its attachment to the inguinal crest and its attachment to the cloaca [1]. The blind end of the bud grows dorsally toward the ventral surface of the metanephric blastema, and by the end of the fifth week, it becomes dilated and pushes into the ventral surface of the metanephric blastema. The metanephric blastema surrounds the dilated blind end of the ureteric duct, forming the E. D. Ruchelli (*) · D. S. Huff Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]; [email protected]

metanephric cap. The straight duct-like segment of the ureteric bud becomes the ureter, and its dilated blind end becomes the rudimentary pelvis. The metanephric cap becomes the renal parenchyma. The segment of the mesonephric duct between the origin of the ureter and the cloaca is now referred to as the common excretory duct. Thus, at the end of the fifth week, the primitive kidney has formed. At the beginning of the sixth week, the blind end of the ureteric bud, which is buried within the ventrally facing hilum of the metanephric cap, dilates to form the cone-­ shaped ampulla [1, 2]. The ampulla is the actively proliferating and elongating end of the ureteric bud. The ampulla continues to grow into the metanephric cap by dichotomous branching morphogenesis. The first branching produces an upper pole branch and a lower pole branch of the primitive pelvis. This causes an elongation of the primitive kidney in the rostralcaudal direction, and the kidney becomes reniform [1]. The lower pole is closer to the midline than the upper pole, which explains why a horseshoe kidney is formed by the fusion of the lower poles. During the second branching, each polar branch of the pelvis divides into a secondary polar branch and an interpolar branch. This results in four primitive major calyces, each surrounded by its lobe of metanephric blastema. The interpolar branches are perpendicular to the polar branches. The polar branches subsequently branch more rapidly than the interpolar branches, causing the polar lobes to be complex, with two or more papillae emptying into one minor calyx, whereas the interpolar lobes remain simple, with only one papilla emptying into a minor calyx. The next few branches form additional primitive major and minor calyces, the cribriform plates of the future papillae, and the first collecting ducts. The primitive pelvis and calyces are narrow tubes that will later be remodeled and become dilated. Therefore, by the end of the sixth week, primitive lobes are recognizable. Although some of the early ureteric bud branches that will be subsequently remodeled into minor calyces induce rudimentary nephrons, many of these early nephrons degenerate and disappear. The rest become detached

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from the ampulla that induced their formation and may persist as rudimentary glomeruloid structures in the renal sinus. During the beginning of the seventh postfertilization week, the metanephric blastema surrounding the tips of the branching collecting ducts will be induced to form nephrons [1–3]. Each branch has a straight ductal stem and a dilated tip, the ampulla. The blastema around the ampulla will differentiate into nephrons, whereas the blastema around the stems will differentiate into specialized renal interstitium. Each ampulla of the first generation of collecting ducts induces its surrounding metanephric blastema to form a nephrogenic vesicle that becomes an “S”-shaped tubule within a few days. The fate map of the “S”-shaped tubule has been well established (Fig. 8.1). In the eighth week, the first generation, consisting of 18 nephrons, is completed [2]. By the end of the week, the

Fig. 8.1  Diagrammatic fate map of the “S”-shaped nephron: seventh week. The lower limb (green, 1) will become the glomerulus. The dark green (2) cells will become the visceral epithelium of the glomerular tuft. The light green (3) cells will become the parietal epithelium of Bowman’s capsule. The middle limb (pink, 6) will become the proximal convoluted tubule and is attached to the margin of the future glomerulus (5). The junction between the middle loop (blue) and the upper loop (purple) will become the descending loop of Henle (light blue, 7), the ascending loop of Henle (dark blue, 8), and the macula densa (purple, 9). One view holds that these precursors of the loop of Henle and the macula densa are the proximal part of the upper limb. The upper limb will become the distal convoluted tubule (orange, 10) and the arched connecting duct (yellow, 11). The arched connecting duct joins the collecting duct (black, 12) at the previous site of the ampulla, which induced the differentiation of the nephron. The brown tissue (4) between the macula densa and the glomerulus is the anlage of the vascular mesenchyme of the future vascular pole of the glomerulus. It will push into the glomerular visceral epithelium and will become the glomerular capillary loops, glomerular mesangium, extraglomerular mesangium (Goormaghtigh cells) of the juxtaglomerular apparatus, and afferent and efferent arterioles. The segment that will become the macula densa is anchored to this position at the vascular pole and will never move

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characteristic histology of each part of the nephron is well developed (Fig. 8.2). The proximal convoluted tubular epithelium has a well-developed brush border. The descending, thin, and ascending limbs of the loops of Henle are identifiable. The epithelial cells of the macula densa, the granular cells of the afferent and efferent arterioles, and the Goormaghtigh cells (also called “lacis cells” or “extraglomerular mesangial cells”) of the juxtaglomerular apparatus are differentiated. Renin activity is identifiable in the afferent arterioles. The distal convoluted tubular epithelium is well differentiated. Principal and intercalated epithelial cells of the collecting ducts are identifiable. Metanephric urine production begins. The pelvis and major calyces are remodeled, begin to dilate, and assume their definitive form, but the minor calyces remain as narrow tubes. The nephro-

Fig. 8.2  Nephron: eighth week. The color scheme is the same as in Fig. 8.1. The junction of the proximal convoluted tubule (pink) and the glomerulus (green) has shifted from the edge of the glomerulus to a site opposite the vascular pole. The proximal convoluted tubule passes behind, to the right, and above the glomerulus. The descending loop of Henle (light blue) lies adjacent to its collecting tubule of origin (black). The macula densa (purple) remains in its original position at the vascular pole. The distal convoluted tubule (orange) passes to the left and in front of the glomerulus. The arched connecting duct (yellow) passes in front of the proximal convoluted tubule and joins its collecting duct of origin (black). The afferent and efferent arterioles have appeared, and the juxtaglomerular apparatus is well differentiated. The glomerular mesangium and the Goormaghtigh cells (both brown) arise from the same primitive tissue (compare with Fig. 8.19)

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Fig. 8.3  Renal lobe: kidney at 19 weeks gestation. This low-power view demonstrates two well-demarcated lobes. Although the cortex is thin at this age, it is already well defined, as is the underlying medullary pyramid. The nephrogenic zone (NZ) can be seen at the periphery of the cortex as a basophilic band of developing nephrons under the capsule and in the column of Bertin (CB), where the two adjacent lobes meet. The renal sinus (RS) consists of loose mesenchymal tissue and is devoid of fat at this age (hematoxylin and eosin (H&E), 2×)

genic zone, definitive cortex, and medulla begin to be recognizable. By the 12th week, the renal pelvis, major and minor calyces, and p­ apillary tips have developed their definitive forms. By the 13th week, the nephrogenic zone, definitive cortex, and medullary pyramids have attained their final appearance (Fig. 8.3).

Histology Anatomically, the human kidney is organized in lobes (see Fig.  8.3) [4]. Each lobe consists of a medullary pyramid and its surrounding cortex. These lobes account for the characteristic gross appearance of the external surface of the fetal kidney, which shows prominent fissures between adjacent lobes, which divide the kidney into polygons. During the third trimester, partial lobar and calyceal fusion occurs, and the number of fissures, papillae, and calyces decreases. The fissures usually disappear during the first few years of life, secondary to renal growth. The peripheral portion of the lobe consists of the cortex, which surrounds the central portion, the medullary pyramid. A portion of the cortex lies under the renal capsule, but where two adjacent lobes meet laterally, the two layers of the cortex from each lobe (septal cortex) become confluent and form the so-called column of Bertin (see Fig. 8.3). The tips of the medulla drain into the papillae and calyceal system,

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and the columns of Bertin are in direct continuity with the mesenchymal tissues of the renal sinus, between the calyces. Both cortex and medullary elements are primarily formed in the nephrogenic zone. The nephrogenic zone is easily recognized at low power in renal sections of all preterm fetuses as a band situated in the outermost portion of the cortex (Fig.  8.4). During the midtrimester, abundant undifferentiated blastema cells can be seen in the background of the developing nephrons, which impart a deeply basophilic appearance to the nephrogenic zone (Fig. 8.5). The number of blastema cells decreases after 30 weeks, and as nephrogenesis comes to an end between weeks 32 and 36, the nephrogenic zone is no longer present in the normal term fetus. At term, mature tubules should be seen between the last generation of glomeruli and the capsule. By the end of gestation, the full complement of nephrons is attained, and subsequent renal growth reflects enlargement and maturation of the existing nephrons. During fetal life, subsequent generations of nephrons formed in the nephrogenic zone are added to the underlying developing cortex in ill-defined “layers.” These “layers” are histologically represented by rows of glomeruli (Fig. 8.6). As the result of this pattern of growth, the oldest (first-­ formed) glomeruli are those in the inner cortex, whereas the youngest (last-formed) glomeruli are those in the outer cortex, immediately under the nephrogenic zone (see Figs.  8.5 and 8.6). The timing of a developmental disturbance can sometimes be presumed based on whether abnormalities involve the entire cortex (an early event) or only the outer cortex (a late event). A few studies [5, 6] have proposed a histologic method to evaluate nephrogenesis by counting the number of “layers” of glomeruli as representative of nephrons, from inner to outer cortex. These estimates need to be performed on well-oriented sections in which the medullary rays are seen extending longitudinally from a well-defined corticomedullary junction to the outer cortex (see Fig.  8.6). Unfortunately, such well-oriented areas are not necessarily common on routine autopsy sections, and some interobserver variability may also exist in the process of identifying “layers.” Nevertheless, based on these studies, an estimate of the average number of layers is shown in Table 8.1. The stages of individual nephron development as described in the Embryology section above can be observed histologically in the nephrogenic zone until 36  weeks gestation (Figs.  8.7, 8.8, 8.9, 8.10, and 8.11). Once formed, fetal glomeruli resemble adult glomeruli but are smaller, and the podocytes have a typical cuboidal appearance. This cuboidal appearance is continuous around the glomerulus at birth, but podocytes gradually

116 Fig. 8.4  Renal cortex during gestation, at 16 weeks (a), 21 weeks (b), 27 weeks (c), and 35 weeks (d) gestation. The cortex becomes thicker as the number of nephrons formed in the nephrogenic zone increases until shortly before term. Notice that the nephrogenic zone is present at all ages except for the kidney at 35 weeks, which shows only a few foci of discontinuous nephrogenesis (see Fig. 8.5 for details) (H&E, a and b, 10×; c and d, 4×)

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flatten during the first year of life; remnants of the cuboidal layer are usually not seen in normal glomeruli after 12 months of age [7]. Glomeruli decrease in size between 12 and 20 weeks gestation (see Fig. 8.5) [8], remain the same size until birth, and then increase in diameter postnatally. Regional differences in glomerular size also exist within the cortex of the fetus and newborn. Juxtamedullary

glomeruli are larger than more immature glomeruli under the nephrogenic zone or capsule (Fig. 8.12). A small percentage of glomeruli (usually 1–2%) undergo involutional sclerosis in otherwise normal kidneys of fetuses and children (Fig. 8.13) [9]. As the glomeruli differentiate on one end of the developing nephron, tubular differentiation can be recognized

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Fig. 8.5  Nephrogenic zone during gestation at 16 weeks (a), 21 weeks (b), 27 weeks (c), and 35 weeks (d) gestation. The nephrogenic zone is continuous and readily observed under the capsule at 16, 21, and 27  weeks gestation. At 35  weeks gestation, it is almost completely absent; only rare foci of immature nephrons are present. Also notice the high density of undifferentiated blastema cells in the background of the

nephrogenic zone at gestational ages 16 and 21 weeks. At gestational age 27 weeks, the number of undifferentiated cells has diminished in the background, but developing nephrons are still present. The number of glomeruli has substantially increased from 16 to 35 weeks. Proximal convoluted tubules with distinct cytoplasmic eosinophilia are seen at all gestational ages (H&E, a, b, and c 20×; d 10×)

histologically relatively early (Fig. 8.14). Proximal tubule differentiation is evident immediately beneath the S-shaped bodies. The characteristic deep eosinophilia of the cytoplasm of proximal tubular epithelial cells allows for easy identification of the tubules, except for those cases with severe autolysis. Immunohistochemistry for CD10 and epithelial membrane antigen (EMA) is also helpful to identify different segments of the nephron (Fig. 8.15). This identification is of particular importance in cases of abnormal tubular differentiation (such as renal tubular dysgenesis), in which the absence or paucity of proximal convoluted

tubules could be overlooked. As the tubules differentiate, they also elongate and increase in diameter, separating the glomeruli from one another. This separation is more pronounced in the deeper cortex (Fig. 8.16). Postnatally, tubular growth is also responsible for the separation of the outermost glomeruli from the capsule and for the formation of a subcapsular zone largely devoid of glomeruli (Fig. 8.17) [7]. Development of the juxtaglomerular apparatus starts very early in gestation (Fig.  8.18). The renin-angiotensin system plays a critical role in kidney development, and the

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Fig. 8.6  “Rows” of glomeruli at 16 weeks (a), 24 weeks (b), and 32 weeks (c) gestation. Generations of nephrons formed in the nephrogenic zone are added to the underlying developing cortex in ill-defined “rows” (H&E, a, b 10×; c 4×)

Table 8.1  Development of glomeruli during gestation Gestational age, wk 23 or less 24 28 32 36

Rows of glomeruli 3 5–7 8–9 9–10 10–14

Data from Clapp and Croker [7] and Dorovini-Zis and Dolman [5]

activity of the intrarenal renin-angiotensin system is high during fetal and neonatal life, declining during postnatal maturation [10]. The juxtaglomerular apparatus is not easily identified on routine sections of fetal kidneys, but renin expression can be demonstrated by immunohistochemistry (Fig. 8.19). In the medulla, the loops of Henle are relatively short in the fetus; significant lengthening occurs after birth. The

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Fig. 8.7  Nephrogenesis. The branches of the ureteric bud form the collecting ducts (CD), seen here as a T-shaped structure. These ureteric bud branches induce the metanephric blastema (Bla) to condense around their tips (ampulla) and then to differentiate into a vesicle (Ves), which develops a lumen and fuses to the end of the T-shaped structure (one on each side). This vesicle will form the entire nephron (H&E, 40×)

Fig. 8.8  Nephrogenesis. Vesicles (Ves) have fused to the T-shaped ends of the ureteric bud branches and start to elongate to form the S-shaped body (H&E, 40×)

Fig. 8.9  Nephrogenesis. The S-shaped body (S) has formed, and a vascular cleft (large arrow) develops at the site where the glomerular capillaries will emerge. Below the cleft, the lower limb of the S-shaped body, most distant from the ureteric bud, differentiates into the parietal (Pa) and visceral (podocytes; Po) epithelium of the glomerulus. The future Bowman’s space (small arrow) can be seen between the two. The middle limb generates the proximal convoluted tubule, and the upper limb forms the loop of Henle, macula densa, and distal convoluted tubule (H&E, 40×)

Fig. 8.10  Nephrogenesis. The developing glomerulus is now better defined, as the capillary loops covered by the podocytes (Po) start to fill out into an expanding Bowman’s space (arrow). Above the glomerulus, cross sections of the developing tubules can be seen connected to the collecting duct (CD) (H&E, 40×)

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Fig. 8.11  Nephrogenesis. Nephrons in various stages of development can be seen within the nephrogenic zone. Developing glomeruli are indicated by arrows (H&E, 20×)

Fig. 8.12  Glomeruli: kidney at 19 weeks gestation. The most recently formed glomeruli increase in size as they become incorporated into the developing cortex under the nephrogenic zone (right, small arrow). Thus, the older juxtamedullary glomeruli (left, large arrow) are larger than more superficial ones in fetuses and infants. All glomeruli retain the cuboidal appearance of podocytes through the end of gestation (H&E, 20×)

Fig. 8.13  Sclerotic glomeruli: kidney at 37 weeks gestation. Scattered sclerotic glomeruli are commonly observed in fetuses and neonates. When present in small numbers (usually 1–2%, occasionally up to 10%) in the absence of other abnormalities, they should not be considered pathologic changes (H&E, 20×)

Fig. 8.14  Proximal convoluted tubules: kidney at 19 weeks gestation. Proximal convoluted tubules (PCT) differentiate very early in nephrogenesis and can readily be identified under the nephrogenic zone (NZ) owing to their distinct cytoplasmic eosinophilia. A proximal convoluted tubule can be seen draining Bowman’s space of a young glomerulus (arrow) (H&E, 20×)

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Fig. 8.15  Immunohistochemical characteristics of different nephron segments: kidney at 22 weeks gestation. (a) Normal tubular differentiation is seen in this optimally oriented field, in which collecting ducts (CD) can be seen extending longitudinally into the cortex. Similar to Fig.  8.14, proximal convoluted tubules display distinct cytoplasmic eosinophilia. (b) CD10 immunostain demonstrates nor-

mal expression on the apical aspect of proximal convoluted tubules (arrows) as well as podocytes and Bowman capsule in the glomeruli. (c) In contrast to CD10, epithelial membrane antigen (EMA) immunostain demonstrates normal expression in distal convoluted tubules and collecting ducts (a H&E, 10×; b CD10 immunostain, 10×; c EMA immunostain, 10 ×)

nephrons formed last (outer cortex) are the shortest, whereas the nephrons formed earliest (juxtamedullary) are the ­longest. In the fetus, thin portions of loops of Henle are present only in the nephrons formed earliest and only in the descending limb, not in the ascending limb [7]. Early in the midtrimester, the medulla contains primarily immature collecting ducts surrounded by abundant loose stroma

(Fig. 8.20). As the number of nephrons increases and the cortex thickens, the medulla becomes denser and is occupied by collecting ducts and elongating loops of Henle from the juxtamedullary nephrons (Fig.  8.21). Like the mature kidney, the fetal medulla displays more abundant interstitium toward the papillae between the ducts of Bellini (distal segment of the collecting duct) (Figs. 8.22 and 8.23).

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Fig. 8.17  Cortex at term. The nephrogenic zone is no longer present, and the full complement of nephrons has been attained. Glomeruli in the neonate are typically very close to the capsule before further tubular growth separates them and creates a subcapsular zone devoid of glomeruli (“cortex corticis”) during infancy (H&E, 4×)

Fig. 8.16  Tubular growth and maturation: kidney at 35 weeks gestation. The young glomeruli are more crowded in the outer cortex, whereas the older glomeruli are more widely separated near the medulla as the result of tubular elongation and increased diameter (H&E, 4×)

Fig. 8.18  Juxtaglomerular apparatus. The different segments of the S-shaped body are committed to become specific portions of the nephron very early in the differentiation process. At the same time, the cells that form the juxtaglomerular apparatus can be identified next to the glomerulus. DCT distal convoluted tubule, JGC juxtaglomerular cells, LH loop of Henle, MD macula densa, PCT proximal convoluted tubule (H&E, 40×)

Fig. 8.19  Renin-expressing cells (granular cells): kidney at 27 weeks gestation. Renin expression is high in the fetus and can be demonstrated by immunohistochemistry, primarily in the terminal afferent arteriole (arrows). The juxtaglomerular apparatus is otherwise not readily evident on routine stains of fetal kidneys (renin immunohistochemistry, 20×)

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Fig. 8.20  Medulla: kidney at 17 weeks gestation. Loose stroma separates immature collecting ducts (CD) and loops of Henle (LH). Thin segments of the loop of Henle are not evident at this age (H&E, 10×)

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Fig. 8.22  Medulla: kidney at 35 weeks gestation. The interstitium is normally more abundant near the papilla; this should not be mistakenly interpreted as fibrosis (H&E, 2×)

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Fig. 8.21  Medulla: kidney at 21 weeks gestation. (a) As the tubules elongate and increase in diameter, the medulla becomes denser. (b) Close-up view of the papilla (arrow) showing maturing collecting ducts (ducts of Bellini) and loops of Henle (H&E, a, 2×; b, 20×)

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Fig. 8.23  Collecting ducts: kidney at 35 weeks gestation. The collecting ducts in the outer medulla (a) are lined by cuboidal epithelium, whereas the collecting ducts in the inner medulla (ducts of Bellini), near the papillae (b), are lined by columnar epithelium (H&E, a, b 20×)

References 1. O’Rahilly R, Muecke EC. The timing and sequence of events in the development of the human urinary system during the embryonic period proper. Z Anat Entwicklungsgesch. 1972;138:99–109. 2. Potter EL.  Normal and abnormal development of the kidney. Chicago: Year Book Medical; 1972. 3. O’Rahilly R, Müller F. Developmental stages in human embryos. Washington, DC: Carnegie Institution Of Washington; 1987. 4. Sykes D. The morphology of renal lobulations and calices, and their relationship to partial nephrectomy. Br J Surg. 1964;51:294–304. 5. Dorovini-Zis K, Dolman CL.  Gestational development of brain. Arch Pathol Lab Med. 1977;101:192–5.

6. Hinchliffe SA, Sargent PH, Chan YF, van Velzen D, Howard CV, Hutton JL, Rushton DI. “Medullary ray glomerular counting” as a method of assessment of human nephrogenesis. Pathol Res Pract. 1992;188:775–82. 7. Clapp WL, Croker BP. Kidney. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 891–970. 8. Souster LP, Emery JL. The sizes of renal glomeruli in fetuses and infants. J Anat. 1980;130:595–602. 9. Emery JL, Macdonald MS. Involuting and scarred glomeruli in the kidneys of infants. Am J Pathol. 1960;36:713–23. 10. Yosypiv IV.  A new role for the renin-angiotensin system in the development of the ureteric bud and renal collecting system. Keio J Med. 2008;57:184–9.

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Urinary Bladder Eduardo D. Ruchelli and Dale S. Huff

Introduction The fetal urinary bladder extends high into the abdomen even when empty, whereas the adult bladder is found entirely within the pelvis minor. The fetal bladder is flanked by the two umbilical arteries, one on each side, and it is anchored to the umbilicus by the cord-like remnant of the urachus. In fact, the dome of the fetal bladder tapers off into the urachal remnant, giving the organ an almond-like shape. Bladder development is dependent on urine production, so any condition that impairs urine production (such as renal agenesis, severe renal cystic disease, or renal tubular dysplasia) will result in hypoplasia of the bladder. Conversely, the fetal bladder becomes enlarged and hypertrophic in the presence of prolonged obstruction of the bladder outlet or urethra. These pathologic changes should be distinguished from the normal variability of physiologic dilatation, which will also affect the thickness of the wall and its histological appearance.

Embryology The urinary bladder and urethra develop from the urogenital portion of the cloaca. The simple columnar epithelium of the entire cloaca is derived from endoderm, whereas the mesenchyme is derived from the splanchnopleuric mesoderm [1]. In the early part of the fifth postfertilization week, when the ureteric bud first appears, the site of the urorectal septum becomes visible on the external surface of the cloaca. By the early part of the sixth week, the urorectal septum delineates the primary urogenital sinus from the anorectum.

E. D. Ruchelli (*) · D. S. Huff Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]; [email protected]

The mesonephric duct distal to the takeoff of the ureteric bud is the common excretory duct, which enters the ventral or urogenital sinus portion of the cloaca. The entrance of the common excretory ducts divides the primary urogenital sinus into the vesicourethral canal rostrally and the definitive urogenital sinus caudally. The vesicourethral canal will form the bladder and proximal prostatic urethra in the male and the bladder and most of the urethra in the female [2]. The definitive urogenital sinus is divided into pelvic and phallic parts. The pelvic part will form the distal prostatic and membranous urethra in the male and the distal urethra in the female. By the end of the sixth week, the urorectal septum completely separates the urogenital sinus from the anorectum. During the seventh week, the common excretory duct gradually becomes incorporated into the posterior wall of the vesicourethral canal, and the ureters and mesonephric ducts enter the vesicourethral canal separately [3]. The ureteral orifices come to lie cranially and laterally to the openings of the mesonephric ducts, which migrate caudally and remain near the midline. The bladder outlet forms just cranially to the openings of the mesonephric ducts. The bladder enlarges and becomes distinct from the urethra. The trigone becomes defined. The trigone and the dorsal midline of the proximal prostatic urethra down to the orifices of the mesonephric ducts are thought to be of mesonephric origin, but a contribution from the urogenital sinus has been suggested. The simple columnar epithelium of the urethra, except at the sinual tubercle, becomes stratified during the seventh week and is composed of several layers. The epithelium of the bladder remains simple and later differentiates into stratified epithelium two or three layers thick [2, 4]. During the eighth week, beginning at the dome of the bladder, the undifferentiated mesenchyme begins to differentiate into lamina propria and muscularis propria [2]. Full maturation of the epithelium and muscularis propria is completed in fetal life.

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Histology The four layers of the bladder wall from the lumen outward consist of the urothelium, lamina propria, muscularis propria, and adventitia (Figs. 9.1 and 9.2) [5, 6]. Some authors describe a muscularis mucosae of the urinary bladder, which may be indistinct and composed of a thin, discontinuous layer of smooth muscle [7]. The urothelium becomes well differentiated early in the midtrimester (Fig. 9.3) and has a similar appearance to the adult urothelium, consisting of a variable number of cellular layers depending on the degree of bladder dilatation. The urothelium is usually up to six to seven layers thick in the contracted bladder, but in autopsy specimens, the urothelium often is poorly preserved, and its layers may be incomplete or completely absent (see Fig. 9.1). The lamina propria is composed of dense connective tissue in which there are blood vessels, lymphatics, and nerves (see Figs. 9.1 and 9.3). The major changes in fetal life in the histological appearance of the urinary bladder occur in the muscularis propria, which is relatively poorly developed in the early midtrimester (see Fig. 9.1) and becomes progressively thicker during the third trimester (Figs.  9.4, 9.5, and 9.6). Similar to the adult bladder, the layers of smooth muscle that constitute the muscularis propria frequently have no definite orientation, and three distinct layers cannot be identified, with the exception of the area of the bladder neck (see Figs. 9.4, 9.5, and 9.6) [6]. The adventitia is a relatively thin, fibrous tissue layer containing large vessels and nerves (see Figs. 9.2 and 9.4).

Fig. 9.2  Urinary bladder and uterus (top) at 19 weeks. This cross section of the bladder demonstrates the typical mucosal folds in the empty, contracted state. The muscularis propria is better developed than in Fig. 9.1, but the smooth muscle fascicles still remain separated by relatively abundant stroma (H&E, 2×)

Fig. 9.1  Urinary bladder at 16 weeks. A full-thickness section of the bladder wall is shown. The urothelium is frequently poorly preserved in autopsy specimens. The underlying lamina propria is relatively thick, depending on the degree of dilatation. The dense connective tissue of the lamina propria blends with the relatively abundant stroma that separates the smooth muscle fascicles of the immature muscularis propria. The adventitia consists of a thin layer of connective tissue, which may be covered by peritoneum in sections of the superior bladder (hematoxylin and eosin (H&E), 10×)

Fig. 9.3  Urinary bladder at 19 weeks. The epithelium is relatively well preserved, as shown here. This figure clearly exhibits urothelial differentiation, and the epithelium will remain unchanged throughout gestation (H&E, 10×)

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Fig. 9.4  Urinary bladder at 23 weeks. The muscularis propria continues to thicken, as shown here. The layers of the muscular coat (inner, central, and outer) have no definite orientation and are imperfectly separated (H&E, 2×)

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Fig. 9.5  Urinary bladder at 32 weeks. The muscularis propria occupies most of the bladder wall, but the smooth muscle fascicles normally remain separated by some connective tissue (compare Fig.  9.1). The other layers remain unchanged (H&E, 4×)

References 1. O’Rahilly R, Muecke EC.  The timing and sequence of events in the development of the human urinary system during the embryonic period proper. Z Anat Entwicklungsgesch. 1972;138:99–109. 2. Felix W. The development of the urinogenital organs. In: Keibel F, Mall FP, editors. Human embryology. Philadelphia: JB Lippincott; 1912. p. 752–79. 3. O’Rahilly R, Muller F. Human embryology and teratology. 3rd ed. New York: Wiley-Liss; 2001. p. 264. 4. Yasuda Y. Genital system. In: Nishimura H, editor. Atlas of human prenatal histology. Tokyo: Igaku-Shoin; 1983. p. 147–69. 5. Valdes-Dapena MA.  Histology of the fetus and newborn. Philadelphia: Saunders; 1979. p. 469–77. 6. Reuter VE, Al-Ahmadie A, Tickoo SK. Urinary bladder, ureter and renal pelvis. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 971–85. 7. Weaver MG, Abdul-Karim FW. The prevalence and character of the muscularis mucosae of the human urinary bladder. Histopathology. 1990;17:563–6.

Fig. 9.6  Urinary bladder at 35 weeks. This full-thickness view of the bladder wall near term illustrates how little the microscopic features of the bladder wall change during the second and third trimesters of gestation, except for an increase in smooth muscle in the muscularis propria (seen in the previous figures). The urothelium (inset) is fully differentiated prenatally (H&E, 4×; inset, ×20)

Testis

10

Dale S. Huff and Chrystalle Katte Carreon

Introduction

versial. One view is that the cords are formed by coelomic epithelium invading the mesoderm [2]; another view is that Testicular development is not a smooth, continuous process. the cords arise from mesonephric glomerular and tubular Instead, it is a series of five abrupt distinct steps of rapid mesodermal epithelium [1]. The cords lack lumens, but as transformation separated by periods of relative inactivity. soon as they become sharply outlined structures, some refer Each step is triggered by a rapid rise in production of gonad- to them as seminiferous tubules. otropins and androgens, which triggers a new step in maturaDuring the seventh week, the testis is attached to the tion of germ cells, Leydig cells, and Sertoli cells. The first remnants of the mesonephros by a thick mesorchium [3]. step begins at the end of the 6th week postfertilization; the The capsule first appears at this stage; it is composed of a second, during the 14th postmenstrual week; the third, dur- simple cuboidal epithelium that covers the surface and is ing the 2nd postnatal month; the fourth, during the 5th post- derived from the visceral layer of the tunica vaginalis and, natal year; and the fifth, during the 9th postnatal year. Some underlying it, the primitive tunica albuginea [4]. Each semof these steps produce dramatic changes in testicular histol- iniferous tubule forms a loop similar to an inverted “U” ogy; others only produce subtle changes in the cytology of with the two ends of the “U” at the hilum, the two straight germ cells, Leydig cells, and Sertoli cells. This chapter parallel limbs radiating toward the periphery, and the arch describes the details of these five steps in testicular differen- of the “U” under the capsule. The two hilar ends of the tiation (Table 10.1). looped seminiferous tubules are narrow, delicate, and solid and are composed of small, dark cells. These are the precursors of the straight tubules and rete testis [5]. The oppoEmbryology: The First Step in Testicular site arched ends of the tubules may blend into the tunica albuginea [4], which explains the occasional inclusion of Differentiation remnants of primitive testicular blastema or individual The first step in testicular differentiation, the transformation germ cells in the tunica albuginea in later fetal or postnatal of the bipotential gonad into the testis, begins toward the end life [6]. These inclusions should not be mistaken for tesof the sixth postfertilization week. In the presence of sex-­ ticular dysgenesis [7]. The cells in the tubules differentiate determining region Y (SRY) protein, also known as testis-­ into fetal Sertoli cells, which are small, basophilic, and determining factor, the primitive gonadal cords of the spindle-shaped. The long axis of the Sertoli cells is perpenbipotential gonad transform into solid testicular seminifer- dicular to the long axis of the seminiferous tubules [4]. ous cords [1]. This process is the first histological evidence Primordial germ cells migrate by amoeboid action from the of testicular differentiation. The origin of the cords is contro- coelomic epithelium and interstitium into the center of the tubules and transform into gonocytes [8, 9]. The transformation of primordial germ cells into gonocytes is the first step in the maturation of male germ cells [10, 11] (see Table 10.1). Gonocytes are the fetal stem cells of the male germ cell line. At 7 weeks postfertilization, gonocytes are D. S. Huff ∙ C. K. Carreon (*) difficult to recognize in routine sections stained with hemaDepartment of Pathology and Laboratory Medicine, The Children’s toxylin and eosin (H&E), but they are demonstrated by Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA immunohistochemical stains for several markers of early e-mail: [email protected]; [email protected]

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_10

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130 Table 10.1  Maturation process of germ cells PFA postfertilization age, PMA postmenstrual age, PNA postnatal age

D. S. Huff and C. K. Carreon Age Embryo (7 weeks PFA): bipotential gonad becomes testis; pool of fetal stem cells established

Least mature Primordial germ cells

Fetus (14 weeks PMA): fetal Leydig cells dominate histologic picture; spermatogonia appear

Gonocytes (fetal stem cells)

Neonate (2 months PNA): pool of adult stem cells established; apoptosis of gonocytes causes reduced number of germ cells

Gonocytes (fetal stem cells)

→

→

Intermediate

More mature Gonocytes (fetal stem cells)

Intermediate → cell

Fetal spermatogonia

Fetal → Spermatogonia

Apoptosis (gonocytes disappear) Adult pale spermatogonia

Adult dark spermatogonia (adult stem cells) Prepubertal (5 years PNA): meiosis established

Adult dark spermatogonia (adult stem cells)

Adult pale spermatogonia; B spermatogonia

Puberty (9 years PNA): complete spermiogenesis, including mature sperm expected by 14 years; pool of adult Leydig cells established

Adult dark spermatogonia (adult stem cells)

Adult pale spermatogonia, B spermatogonia, primary spermatocytes, secondary spermatocytes, early spermatids, late spermatids

germ cells such as Oct-4, placental alkaline phosphatase (PLAP), and CD117. Rising maternal plasma levels of human chorionic gonadotropin stimulate precursors of Leydig cells in the interstitium to differentiate into fetal Leydig cells. Testosterone production begins, and Sertoli cells begin to produce Müllerian-inhibiting substance. The two major results of the first step in testicular differentiation are the transformation of the bipotential gonad into the testis and the formation of the fetal stem cell pool. By the eighth postfertilization week, the shape of the testis has changed from a long and flat structure typical of a bipotential gonad to a short, ovoid, and smooth-surfaced structure typical of a testis. The simple cuboidal surface epithelium may be focally multilayered, and the tunica albuginea increases in thickness. The mesonephric body has almost completely degenerated. Five to twelve of the epigenital nephrons adjacent to the hilum of the testis do not completely degenerate (see Fig. IV.4 in the Part IV introduction). The collecting ducts of these nephrons remain. The blind ends of these degenerating collecting ducts lie adjacent to the tubules of the rete testis but do not connect with them.

Adult dark spermatogonia (adult stem cells)

Primary spermatocytes (prophase first meiotic division) Mature sperm

The opposite ends are attached to the mesonephric duct. The remnants of these mesonephric nephrons will become the efferent ductules of the head of the epididymis [3]. The seminiferous tubules become more sharply outlined by a distinct basement membrane [4], and their peripheral looped ends are distinctly separated from the tunica albuginea. Gonocytes become recognizable in some well-preserved and well-­ prepared routine sections, and they occupy the center of the tubule. The levels of human chorionic gonadotropin reach their peak. The gradual differentiation of fetal Leydig cells continues, but their numbers remain relatively low, and the cells are only moderately enlarged with increasingly eosinophilic cytoplasm [4, 12]. They are not a prominent histological feature of the embryonic testis. The interstitium from local mesonephric mesenchyme extends between the seminiferous tubules in the hilum. This loose interstitium contains fibroblasts, vessels, nerves, mast cells, and pre-Leydig cells. By the end of the eighth postfertilization week, the germ, Sertoli, and Leydig cell lines are present, and the production of Müllerian-inhibiting substance and testosterone is established.

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Histology General Overview At term, the testis is covered by a thick, smooth capsule, except at the hilum. From the outside in, the capsule comprises three layers: (1) a layer of simple epithelium on the surface (tunica vaginalis); (2) a layer of dense connective tissue (tunica albuginea); and (3) the deepest layer, composed of loose vascular connective tissue (tunica vasculosa). The tunica vaginalis and tunica albuginea are present when the testis first differentiates from the bipotential gonad. The tunica vasculosa gradually develops between 20 and 30 weeks gestation. The parenchyma is composed of seminiferous tubules and interstitium. The seminiferous tubules are tightly coiled. The tubules contain Sertoli cells and germ cells and are surrounded by a basement membrane, a layer of peritubular myoid cells, and a layer of peritubular fibroblastic spindle cells. Interlobular septa divide the seminiferous tubules into approximately 250 lobules, each containing one to four looped seminiferous tubules. The interstitium contains Leydig cells, spindled stromal cells, vessels, nerves, mast cells, and macrophages. The hilar ends of the seminiferous tubules become straight tubules that connect to the mediastinal component of the rete testis, which in turn connects to the head of the epididymis through the efferent ductules.

 0–20 Weeks Postmenstrual Age: The Second 1 Step in Testicular Differentiation The second step in testicular differentiation begins at 14 weeks postmenstrual age. It causes a dramatic change in the histology of the fetal testis. By the 11th week, the hormonal control of testicular differentiation switches from placental human chorionic gonadotropin to fetal pituitary gonadotropins. Fetal pituitary luteinizing hormone triggers this second step in testicular differentiation. Most of the change occurs by 16  weeks. Thereafter, fetal Leydig cells gradually regress in size and number. Two important events occur during this step. The first is the rapid proliferation and maturation of fetal Leydig cells, which produce increasing amounts of testosterone. The second is the transformation of gonocytes to fetal spermatogonia, which is triggered by the increasing levels of testosterone (see Table 10.1).

 aturation of Fetal Leydig Cells M The most dramatic histological manifestation of the second step in testicular differentiation is the rapid increase in the number and size of fetal Leydig cells [12]. Fetal luteinizing hormone stimulates pre-Leydig cells in the interstitium to begin to mature into fetal Leydig cells. Pre-Leydig cells are spindle cells, which are difficult to distinguish from other spindle cells in the stroma and are derived either from coelomic epithelium or from mesonephric mesodermal epithelium. In the process of transforming into fetal Leydig cells, they

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acquire a large amount of bright pink cytoplasm and become round to polygonal cells with sharp cytoplasmic borders. Their nuclei become large and contain a large, round central nucleolus. Mature fetal Leydig cells are identical to adult Leydig cells except that they lack the Reinke crystalloids of the adult form. Fetal Leydig cells resemble fetal adrenal cortical cells, to which they are closely related. By 14 weeks gestation, fetal Leydig cells constitute half the volume of the testis. Longitudinal sections through the hilum demonstrate straight seminiferous tubules separated by broad bands of fetal Leydig cells (Fig. 10.1). Many of the bands of Leydig cells are broader than the seminiferous tubules [13]. Sections perpendicular to the long axis of the tubules demonstrate widely scattered tubular cross sections in a sea of Leydig cells (Fig. 10.2). The relative number and size of fetal Leydig cells (Figs.  10.1, 10.2, 10.3, 10.4, 10.5, 10.6, 10.7, 10.8, and 10.9) and the levels of testosterone are greatest between 14 and 19 weeks gestation.

 aturation of Germ Cells M By 14–15 weeks gestation, rapidly increasing levels of testosterone in the fetal testis and plasma trigger the second step in the maturation of germ cells: the transformation of gonocytes into fetal spermatogonia (see Table  10.1) [14]. During this transformation, the gonocytes in the center of the tubule extend a pseudopod, which attaches to the basement membrane. The gonocytes migrate toward the basement m ­ embrane and transform into intermediate cells. On arrival at the basement membrane, the intermediate cells transform into fetal spermatogonia. Because gonocytes are the stem cells of the fetal male germ cell line, they not only proliferate to produce increasing numbers of fetal spermatogonia, but they also maintain their own numbers. The classification of the subtypes of germ cells used in this chapter is from Fukuda et al. [9] and Wartenberg et al. [15]. Several confusing nomenclatures have been used in the classification of the subsets of fetal germ cells (Table 10.2) [16].

Fig. 10.1  Fetal testis at 15  weeks gestation. The straight loops of the uncoiled seminiferous tubules in the left half are cut longitudinally; those in the right half are cross-sectioned. The hilar ends of the seminiferous tubules abruptly become the narrow, straight tubules. The narrow zone that contains the short, straight tubules is the septal compartment of the rete testis. The straight tubules connect the seminiferous tubules with the mediastinal portion of the rete testis. The broad bands of pink tissue between the tubules are fetal Leydig cells. The tunica albuginea of the capsule is apparent. LP lower pole of testis, RT mediastinal compartment of the rete testis, TEp tail of the epididymis, UP upper pole of testis (H&E, 2×)

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Fig. 10.2  Fetal testis at 15 weeks gestation. This is a higher magnification of Fig. 10.1. Fetal Leydig cells constitute approximately half of the volume of the testis (H&E, 4×)

Fig. 10.4  Rete testes at 15 weeks gestation. This is a higher magnification of Fig. 10.1. This image shows details of the septal compartment of the rete. At the top of the figure, the hilar ends of two seminiferous tubules abruptly become narrow and form two straight tubules (arrows). Collectively, the straight tubules constitute the septal compartment of the rete testis. In the middle of the figure, the two straight tubules join the mediastinal compartment of the rete testis (asterisks), which occupies the lower half of the figure. In adults, at the junctions of the seminiferous tubules and straight tubules, the Sertoli cells are oriented parallel to the long axis of the tubule with the apices of the cells pointing downstream toward the straight tubules. The tips of the cells extend into the rete testis to form a valve that prevents retrograde flow of fluid and cells from the rete testis back into the seminiferous tubules. This structure is either not yet developed or very difficult to identify in the fetus. The rete testis is not yet canalized. In the septal compartment of the rete testis, fetal Leydig cells fill the area between the straight tubules (H&E, 20×)

D. S. Huff and C. K. Carreon

Fig. 10.3  Capsule of the fetal testis at 15 weeks gestation. This is a higher magnification of Fig. 10.1. The capsule consists of two layers. The visceral epithelium of the processus vaginalis forms the flat to low cuboidal epithelium that covers the surface and is called the tunica vaginalis. The second layer, the tunica albuginea, is a uniformly thick, homogeneous band composed of several horizontal, parallel layers of spindled fibroblasts and collagen. Its deep surface is sharply demarcated from the underlying Leydig cells and is parallel to the surface of the testis. The fetal Leydig cells form broad sheets between the tubules and between the tips of the tubules and the tunica albuginea. In contrast to the situation during the embryonic period, the tips of the tubules no longer blend into the tunica albuginea. The ends of the loops of seminiferous tubules are not coiled (H&E, 20×)

Fig. 10.5  Seminiferous tubule of the fetal testis at 15 weeks gestation. This image is a high-power view of a cross section of one seminiferous tubule from the same testis shown in Fig. 10.1. The tubule is surrounded by one or two layers of peritubular myoid and fibroblastic cells. The tubule contains six or eight large germ cells with large, round nuclei, some of which contain one or two large nucleoli (asterisks). It is difficult to distinguish between gonocytes, intermediate cells, and fetal spermatogonia. The germ cells have a large amount of clear cytoplasm, and several are located on the basement membrane. The smaller cells with oval or elongated nuclei and little cytoplasm are fetal Sertoli cells. The large fetal Leydig cells surrounding the tubule have large amounts of eosinophilic cytoplasm similar to adult Leydig cells, but Reinke crystalloids are not seen (H&E, 60×)

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Fig. 10.6  Fetal testis at 17 weeks gestation. This low-power view is of a testis that is 2  weeks older than the testis in Figs.  10.1, 10.2, 10.3, 10.4, and 10.5. The upper pole, with an attached solid appendix testis, is to the left. The epididymis is at the bottom of the testis. The efferent ductules (ED) extend from the rete testis to the epididymis. The tail of the epididymis (TEp) is embedded in the gubernaculum (Gu). The bright red Leydig cells constitute much of the volume of the testicular tissue. BEp body of epididymis, HEp head of epididymis (H&E, 1×)

Fig. 10.8  Fetal testis at 17 weeks gestation. This is a higher magnification of Fig. 10.6. This image shows cross sections of seminiferous tubules and early regression of fetal Leydig cells. The fetal Leydig cells constitute a large portion of the volume of the testis, but some are dedifferentiating into smaller, spindle-shaped juvenile Leydig cells (H&E, 10×)

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Fig. 10.7  Fetal testis at 17  weeks gestation. This is a higher-power view of Fig. 10.6. The capsule is composed of the visceral epithelium of the tunica vaginalis on the surface and the tunica albuginea. A third layer of loose, vascular connective tissue is beginning to form deep in the tunica albuginea and will become the tunica vasculosa. The most distal portions of the seminiferous tubules are slightly coiled. The fetal Leydig cells are beginning to dedifferentiate and regress in size and number. The columns of Leydig cells between the tubules become progressively narrower. There are very few Leydig cells between the tips of the tubules and the tunica albuginea. Some of the fetal Leydig cells are becoming smaller and spindle-shaped, especially in the column closest to the right edge of the figure. This change indicates that fetal Leydig cells are regressing into juvenile Leydig cells (H&E, 10×)

Fig. 10.9  Seminiferous tubule of the fetal testis at 17 weeks gestation. This image shows a seminiferous tubule of Fig. 10.6 at high power. The peritubular myoid and fibroblastic cells are one or two cells thick. At least nine germ cells are present and are marked with asterisks above their nuclei. Even in this well-preserved specimen, the germ cells are difficult to differentiate from Sertoli cells. Identification of subtypes of germ cells is impossible in most autopsy specimens. In the right lower quadrant of the tubule, three large germ cells on or near the basement membrane are easily distinguished from the smaller, elongated, or oval Sertoli cells between them. Some of the Leydig cells closest to the tubule, especially at approximately 3 o’clock, are small and spindle-­shaped. These cells are dedifferentiating juvenile Leydig cells and will form a pool from which fetal and adult Leydig cells will differentiate in the future (H&E, 60×)

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Table 10.2  Alternative nomenclature for subsets of germ cells in the embryo and fetus Names used in this chapter Primordial germ cells Gonocytes Intermediate cells Fetal spermatogonia

Alternative namesa Gonocytes Primitive germ cells Primitive germ cells Primordial germ cells Prespermatogonia M, T1, or T2 prespermatogoniab Prospermatogonia M, T1, or T2 prospermatogoniab Gonocytes

Terminology may overlap and is sometimes used interchangeably. Each alternative name may refer to any of the four names used in this chapter or to any combination of the four b Prespermatogonia and prospermatogonia have been subdivided into three subtypes: M (multiplying), TI (transitional nonproliferating), and T2 (transitional proliferating) a

loose inner layer of connective tissue, the tunica vasculosa, forms on the deep surface of the tunica albuginea (see Figs.  10.3 and 10.7). Septa begin to form from the tunica vasculosa, and the fibrous septa in the hilum begin to extend peripherally. The seminiferous tubules begin to elongate, and their peripheral ends begin to coil (see Figs. 10.6, 10.7, 10.8, and 10.9). The median tubular diameter increases to 59 μm [13] (see Figs. 10.5 and 10.9). Peritubular myoid cells appear before 15  weeks gestation and become more prominent thereafter (see Figs. 10.5 and 10.9). Some of the collecting tubules of the epigenetic mesonephric nephrons begin to form a solid junction with the rete, to form efferent ductules. This union is called the urogenital union [3] (see Fig. 10.6). The solid junctions become canalized late in fetal life or early in postnatal life.

20–40 Weeks Postmenstrual Age In contrast to the dramatic change in testicular histology caused by the maturation and proliferation of Leydig cells, this step in the maturation of germ cells causes very little change in the cytologic appearance of individual seminiferous tubules [13]. Distinguishing between gonocytes, intermediate cells, and fetal spermatogonia is very difficult in routine sections from autopsies (see Fig. 10.5). Gonocytes occupy the center of the tubules. They have a large, round central nucleus; one or two large, round nucleoli; and a scant amount of clear glycogen-rich, PAS-positive cytoplasm. Intermediate cells are larger than gonocytes, round to oval, and are close to or on the basement membrane. They have a large amount of clear glycogenrich cytoplasm. Fetal spermatogonia are larger than intermediate cells. They are oval, lie flat on the basement membrane, and have a large, round nucleus and an inconspicuous or absent nucleolus. The distinction between the cell types is more easily made in very fresh tissue obtained within an hour of death, in semi-thin (0.5–1.0 μm thick) sections of epon-embedded samples, and in electron micrographs. The immunophenotypes of the morphological subsets of germ cells in the fetal and neonatal periods have been described [17, 18]. Each cross section of a seminiferous tubule usually contains six or more germ cells of various types, which can be sometimes challenging to distinguish (see Fig. 10.5) [13]. The second type of cell in the tubule is the fetal Sertoli cell. Fetal Sertoli cells are the most numerous cells in the tubule. They are small, oval, dark cells with very little cytoplasm. They are oriented perpendicularly to the basement membrane and fill up the tubule around the germ cells. Their microscopic appearance does not change during fetal life. Although they look unimportant, they play vital roles in the maintenance and regulation of the maturation of germ cells.

Other Changes Between 10 and 20  weeks gestation, the capsule begins to change. The tunica albuginea continuously increases in thickness and becomes a layer of dense, fibrous tissue beneath the surface epithelium of the tunica vaginalis. A

 egression and Dedifferentiation of Fetal Leydig R Cells The most dramatic histologic change in this period is the continued regression, dedifferentiation, and apparent disappearance of fetal Leydig cells. They become smaller, vacuolated, and finally revert to spindle-shaped cells similar to pre-Leydig cells, which are difficult to distinguish from other connective tissue cells of the interstitium [4, 12, 13]. These are juvenile Leydig cells, which, along with pre-Leydig cells, form a pool from which more fetal and adult Leydig cells can be recruited. Some may undergo apoptosis. Because they are difficult to identify in routine histological sections from autopsies, Leydig cells have been said to be virtually absent in some near-term fetuses. However, they are recog-

Fig. 10.10  Fetal testis at 21 weeks gestation. This testis is dramatically different from the 17-week testis shown in Figs. 10.6, 10.7, 10.8, and 10.9. This image shows the capsule and peripheral ends of the seminiferous tubules. The fetal Leydig cells have almost completely disappeared, and the tubules are extensively coiled and packed back-to-­back. The tunica vasculosa (TV) is more distinct; its deep surface is scalloped because fibrovascular septa arise from it and extend down into the parenchyma, dividing it into lobules. The surface epithelium, the tunica vaginalis, is cuboidal and contains nests of germ cells. The germ cells and Sertoli cells are unusually well preserved and distinct (H&E, 10×)

10 Testis

Fig. 10.11  Seminiferous tubule of the fetal testis at 21 weeks gestation. This high-power view is of a cross section of one seminiferous tubule from the same testes as seen in Fig. 10.10. The peritubular myoid and fibroblastic cells are more numerous than in the testes at earlier gestational ages. The germ cells and Sertoli cells have not changed and are unusually distinct in this specimen. A total of 14 germ cells are marked with asterisks above their nuclei. The Leydig cells are mostly juvenile forms and blend into the peritubular spindle cells (H&E, 60×)

Fig. 10.13  Fetal testis at 23 weeks gestation. This is a higher-power view of Fig. 10.12. The fetal Leydig cells are partially regressed into smaller, elongated juvenile Leydig cells. TA tunica albuginea, TV tunica vasculosa (H&E, 4×)

nizable in semi-thin sections of epon-embedded tissue and in electron micrographs. As fetal Leydig cells regress, the seminiferous tubules become more and more crowded. Compare the appearance of the testis at 21 weeks gestation (Figs. 10.10 and 10.11), 23  weeks gestation (Figs.  10.12, 10.13, and 10.14), 26 weeks gestation (Figs. 10.15, 10.16, 10.17, 10.18, 10.19, and 10.20), 39  weeks gestation (Figs.  10.21, 10.22, 10.23, 10.24, 10.25, and 10.26), and 41  weeks gestation (Figs. 10.27, 10.28, 10.29, and 10.30).

 aturation of Germ Cells M The appearance of germ cells does not change in routine histological sections (Figs.  10.11, 10.18, 10.23, and 10.29).

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Fig. 10.12  Fetal testis at 23 weeks gestation. The tunica vasculosa is very vascular, and the fibrovascular interlobular septa extend more deeply into the parenchyma toward the hilum than in earlier specimens. The seminiferous tubules are extensively coiled. Fetal Leydig cells are less numerous than in the testis at 21 weeks gestational age. The timing of the maturational changes varies from fetus to fetus by several weeks. BEp body of epididymis, HEp head of epididymis (H&E, 2×)

Fig. 10.14  Fetal testis at 23 weeks gestation. This is a higher-power view of Figs. 10.12 and 10.13. Most of the Leydig cells are regressing, small, spindle-shaped juvenile forms. The germ cells are almost impossible to identify, even in this relatively well-preserved section (H&E, 20×)

However, the immunohistochemical phenotypes of germ cells may change, and additional subsets of germ cells seen between 20 and 40  weeks gestation may be identified in the future. Apoptosis of germ cells is an important feature in their maturation [19].

Other Changes The next most obvious histological change is the increased coiling of the seminiferous tubules owing to their increasing length [4, 13]. Coiling becomes maximal by approximately 30  weeks gestation. The increased coiling adds to the effect of the regression of Leydig cells, to result in the tightly packed appearance of the tubules. Compare the

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Fig. 10.15  Fetal testis at 26 weeks gestation. In this low-power view of the entire testis, the extratesticular portion of the epididymis, seen at the lower border of the upper pole (UP), joins with the efferent ductules (ED) that stream into the head of the epididymis (HEp). The vas deferens (VD) is joining the spermatic cord (SC). There are no visible Leydig cells, and the interlobular septa, which are accentuated owing to edema, extend from the tunica vaginalis to the hilum. Gu gubernaculum, LP lower pole of testis, RT mediastinal compartment of the rete testis, TEp tail of the epididymis (H&E, 1×)

Fig. 10.17  Seminiferous tubules of the fetal testis at 26 weeks gestation. This is a higher magnification of Fig.  10.16. Leydig cells are extremely difficult to identify. A few possible single juvenile Leydig cells and clusters of Leydig cells are indicated by arrows. The layers of peritubular spindle cells are very thick; the outer layers of these spindle cells are thought to be composed of completely regressed Leydig cells, which can be referred to as pre-Leydig cells. Germ cells are difficult to identify in this section (H&E, 20×)

D. S. Huff and C. K. Carreon

Fig. 10.16  Fetal testis at 26 weeks gestation. This image is a higher magnification of the capsule and peripheral loops of the seminiferous tubules. The capsule is composed of the simple, cuboidal epithelium of the visceral layer of the tunica vaginalis (TVag) on the surface; the outer dense, fibrous layer called the tunica albuginea; and the inner loose, highly vascular layer that gives rise to the interlobular septa, now called the tunica vasculosa (TVas). The peripheral loops of the seminiferous tubules are tightly coiled and back-to-back. The fetal Leydig cells have all regressed into small, spindled juvenile forms that cannot be distinguished from the stromal fibroblasts (H&E, 10×)

Fig. 10.18  Seminiferous tubule at 26 weeks gestation. This image is a higher magnification of a single tubule from Fig.  10.16. Regressed, spindle-­shaped juvenile Leydig cells are marked by arrows. Germ cells within the tubule in the center are marked by asterisks above their nuclei (H&E, 60×)

10 Testis

Fig. 10.19  Rete testis at the hilar aspect of the upper pole of the testis at 26  weeks gestation. This is from the same testis as shown in Fig. 10.15. The seminiferous tubules surround the top and two sides of the rete testis, and the bottom of the rete testis is at the hilar surface. The septal compartment of the rete testis is composed of the straight tubules, which are the short, narrow tubules connecting the hilar ends of the seminiferous tubules to the mediastinal compartment. The mediastinal compartment is the largest compartment of the rete testis. The extratesticular compartment, which connects the mediastinal compartment to the efferent ductules, is in the right lower border of the rete testis. Two cross sections and one oblique section of efferent ductules are in the right lower corner of this figure. The efferent ductules are heading toward the head of the epididymis, which is to the right (not shown). Lumens are not present in the rete testis (H&E, 20×)

Fig. 10.21  Fetal testis at 39  weeks gestation. In this low-power view of the testis, the three layers of the capsule—the tunica vaginalis, the tunica albuginea, and the tunica vasculosa—are sharply defined. The interlobular septa extend from the tunica vasculosa to the hilum. The seminiferous tubules are tightly coiled and back-toback. Leydig cells are not recognizable at this magnification (H&E, 1×)

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Fig. 10.20  Septal compartment of the rete testis at 26 weeks gestation. The junctions between two seminiferous tubules and two straight tubules are marked by arrows. The straight tubules are narrow and short, and they join the mediastinal compartment (not shown) (H&E, 20×)

Fig. 10.22  Seminiferous tubules and Leydig cells at 39 weeks gestation. This is a higher magnification of Fig. 10.21. Although Leydig cells are not recognizable at low magnification, numerous partially regressed and dedifferentiated juvenile Leydig cells are present between the tubules. Many of the germ cells are apparently undergoing apoptosis. Note the difference between the testis shown here at 39 weeks gestation, in which Leydig cells are easily identified, and the previous images at 26 weeks, in which Leydig cells are difficult to identify even at high power. The timing of maturational changes in the testis varies widely from fetus to fetus (H&E, 20×)

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Fig. 10.23  Single seminiferous tubule at 39 weeks gestation. This is a high-power view of Fig. 10.21. Four viable germ cells are marked with asterisks above their nuclei, and two apoptotic germ cells are marked with arrows. Partially regressed juvenile Leydig cells (LC) are seen in the interstitium (H&E, 60×)

Fig. 10.24  Rete testis at 39 weeks gestation. This is a higher magnification of Fig.  10.21. In this medium-power image of the rete testis, seminiferous tubules are present along the upper edge. Efferent ductules are at the lower right and are streaming toward the head of the epididymis, which is out of the picture to the right. The septal, mediastinal, and extratesticular compartments are visible, but only a few lumens are present (H&E, 4×)

Fig. 10.25  Septal compartment of the rete testis at 39 weeks gestation. This is a higher magnification of Fig.  10.24. The junctions between two seminiferous tubules and two straight tubules are marked with arrows, and lumens are present in some straight tubules. The irregular cuboidal to columnar epithelium of the rete testis is becoming recognizable. Irregular bundles of dense collagen appear between the channels of the rete testis (H&E, 20×)

Fig. 10.26  Extratesticular compartment of the rete testis at 39 weeks gestation. This is a higher magnification of Fig. 10.24. The large, round efferent ductules occupy the right lower half of the picture. The efferent ductules are lined by pseudostratified columnar and cuboidal epithelium with apical blebs. The arrows point to three sites at which the blind ends of efferent ductules abut against the blind ends of extratesticular tubules of the rete testis. These junctions have not yet canalized. Nests of erythroid extramedullary hematopoiesis are present (H&E, 20×)

appearance of the tubules at the various gestational ages. The tunica albuginea continues to increase in thickness, and the third layer of loose connective tissue, the tunica vasculosa, increases in thickness and vascularity (see Figs. 10.10, 10.12, 10.13, 10.15, 10.16, 10.21, and 10.27). The fibrous septa extend from the hilum to the tunica vasculosa (see Figs.  10.15, 10.21, and 10.27). The septal, mediastinal, and extratesticular portions of the rete become more recognizable. The septal portion is composed of the

straight tubules (also known as tubuli recti), and the mediastinal portion is the labyrinth of anastomosing clefts. In the extratesticular portion, the labyrinth coalesces into 5–12 tubules that join with the efferent ductules. Lumens begin to form in the rete at approximately 24 weeks gestation [20] and are prominent by 40  weeks gestation. Canalization of the junction of the rete and the efferent

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Fig. 10.27  Fetal testis at 41  weeks gestation. This low-power view shows a cross section of the testis. The cause of the interstitial edema localized to the right half of the testis is not known, but it accentuates the interlobular septa. BEp body of the epididymis, Mo mesorchium, RT mediastinal compartment of the rete testis (H&E, 1×)

Fig. 10.28  Fetal testis at 41 weeks gestation. This image is a medium-­ power view of seminiferous tubules and interstitium from Fig. 10.27. The small, roundish cells with frayed pink cytoplasm in the interstitium are juvenile Leydig cells (asterisks) (H&E, 20×)

Fig. 10.29  One seminiferous tubule at 41 weeks gestation. Four germ cells are marked by asterisks above their nuclei. The cell in the upper left of the tubule is undergoing apoptosis. The arrows point to five juvenile Leydig cells. The spindle-shaped cell on the right side of the tubule blends in with the peritubular myoid and fibroblastic cells (H&E, 60×)

Fig. 10.30  Rete testis at 41 weeks gestation. The rete testis fills the center of the image. The seminiferous tubules are around the right, upper, and left sides of the rete testis. The hilum is at the bottom of the image. The septal compartment is at the right upper edge of the mediastinal compartment. The rete testis is completely canalized, and the irregular squamoid and cuboidal lining epithelium of the rete testis can be appreciated (H&E, 4×)

ductules may not be complete until postnatal life (Figs.  10.19, 10.20, 10.24, 10.25, 10.26, and 10.30). Extramedullary hematopoiesis and lipochrome pigment increase with increasing gestational age [13].

 Months Postnatal Age: The Third Step 2 in Testicular Differentiation—Mini-puberty The third step of testicular differentiation occurs at 2–3 months postnatal age. It is a fundamental transforma-

tion with two major results: the appearance of the pool of adult male stem cells (adult dark spermatogonia) and the pruning of the number of germ cells through extensive apoptosis of gonocytes. This step is often referred to as the “mini-­puberty.” At the onset of this event, the seminiferous tubules contain numerous germ cells that are gonocytes, intermediate cells, and fetal spermatogonia (Figs.  10.31 and 10.32). The mini-puberty is initiated by a transient surge in luteinizing hormone secretion, which stimulates

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Fig. 10.31  Postnatal testis, 2  months. This high-power view is of a seminiferous tubule from an infant prior to the onset of the mini-puberty at 2 months postnatal age. The cytology of the germ cells and Leydig cells is distinct. Approximately 18 germ cells are present. The two germ cells in the center of the tubule labeled “1” are gonocytes. The one labeled “2” is a gonocyte that has attached itself to the basement membrane by a pseudopod. The two on the basement membrane at the right end of the tubule (labeled “3”) are intermediate cells. The one on the basement membrane at the right end of the tubule and the one in the lower right corner are fetal spermatogonia (both labeled “4”). The tubule in the lower left corner contains a gonocyte migrating to the basement membrane (“2”) and two fetal spermatogonia (“4”) on the basement membrane. The other germ cells are not further classified. The arrows point to juvenile Leydig cells (Toluidine blue, 60×)

Fig. 10.33  Postnatal testis, 6 months. Shown here is the seminiferous tubule from an infant after completion of the mini-puberty at 6 months of age. The number of germ cells is greatly reduced, and gonocytes are no longer present. Adult dark spermatogonia (labeled “5”) are present. A few fetal spermatogonia (labeled “4”) remain. Leydig cells (asterisks) are difficult to identify even in this preparation; they are hidden among the peritubular and interstitial spindle cells (Toluidine blue, 60×)

D. S. Huff and C. K. Carreon

Fig. 10.32  Postnatal testis, 2 months. Shown here is the same testis as in Fig. 10.31. The interstitium is filled with juvenile Leydig cells. The arrows point to clusters of Leydig cells that, in routine autopsy sections, would look like those in Figs. 10.15, 10.16, 10.27, and 10.28. In routine autopsy sections, the elongated Leydig cell marked with an asterisk would look like the long cell next to the tubular basement membrane in Fig. 10.29 (Toluidine blue, 60×)

the recruitment and maturation of fetal Leydig cells from pre-Leydig cells and from the regressed juvenile Leydig cells. Secretion of testosterone triggers the third step in the maturation of germ cells: the transformation of gonocytes, the fetal stem cells, into adult dark spermatogonia, the adult stem cells (see Table 10.1) [21]. It is not clear whether gonocytes transform directly into adult dark spermatogonia, indirectly through fetal spermatogonia, or both. Future immunohistochemical studies may more accurately identify the stem cells of the germ cell line. A minority of gonocytes transform into adult dark spermatogonia; the majority undergo apoptosis. Fetal spermatogonia remain but may have a different immunophenotype than those present during the fetal period. As stem cells, the adult dark spermatogonia mature into adult pale spermatogonia and also maintain their own numbers. The mini-puberty and the transformation of gonocytes into adult dark spermatogonia last for several weeks, beginning to recede by the third or fourth postnatal month. Secretion of luteinizing hormone falls, fetal Leydig cells again regress to juvenile Leydig cells, and secretion of testosterone falls. When this surge in activity is over, gonocytes have disappeared and are replaced by adult dark and adult pale spermatogonia. The number of germ cells per tubular cross section falls from six or more to two because of the apoptosis of gonocytes. Juvenile Leydig cells are difficult to identify in

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Fig. 10.34  Prepubertal testis at early childhood. Shown here is seminiferous tubule from a 5-year-old boy. Few gonocytes (“1”) are present toward and along the basement membrane. Fetal spermatogonia are no longer identified (Toluidine blue, 60×)

routine sections but are identifiable in epon-embedded 0.5-­ μm sections (Fig.  10.33). In normal testes, gonocytes should be absent by 6 months of age [14]. The presence of some cells positive for PLAP, Oct-4, CD117, and other markers of immature germ cells during the first 6 months of life is normal and not necessarily diagnostic of carcinoma in situ. The successful completion in this step in germ cell maturation is essential for the development of normal fertility. Figures 10.31, 10.32, 10.33, 10.34, 10.35, 10.36, and 10.37 are biopsies that were immediately fixed in glutaraldehyde, embedded in epon, sectioned at 0.5-μm thickness, and stained with toluidine blue.

 Years Postnatal Age: The Fourth Step 5 in Testicular Differentiation—Onset of Meiosis The fourth step in testicular differentiation usually begins at 5 years postnatal age, but its onset may range from 3 to 6 years. The histologic hallmark of this step is the appearance of primary spermatocytes, which are cells in prophase of the first meiotic division. The ability of the male germ cell line to enter meiosis is a major hurdle in male germ cell maturation. Although primary spermatocytes are thought to be ­present in all testes at this age, they may be present in only one or two tubules of a given biopsy or sample of a testis. Primary spermatocytes are large cells with large nuclei, with the chromatin patterns characteristic of cells in meiosis. They usually occur in small clusters

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Fig. 10.35  Prepubertal testis at early childhood. Shown here is the same testis as in Fig. 10.34. Rare scattered primary spermatocytes may be seen at about this age, but they can be very difficult to find. Note the presence of a single primary spermatocyte (“1”) toward the center of the seminiferous tubule in this biopsy, which was incidentally found on deeper sections (Toluidine blue, 60×)

of two to four cells located in the center of the tubule (Figs. 10.34 and 10.35). This is a transient step; from 6 to 9 years postnatal age, primary spermatocytes are no longer seen. It is as if the testis is practicing how to enter meiosis.

 Years Postnatal Age: The Fifth Step 9 in Testicular Differentiation—Puberty and Complete Spermiogenesis The pubertal surge in gonadotropins stimulates the gradual maturation of juvenile Leydig cells into adult Leydig cells and the increased production of testosterone, which in turn triggers the reappearance of primary spermatocytes and meiosis. Complete spermiogenesis gradually unfolds with the sequential appearance of secondary spermatocytes, early spermatids, late spermatids, and finally mature sperm. Concomitantly, Leydig cells continue to mature, increase in size and number, and acquire Reinke crystals. Immature Sertoli cells also mature into adult Sertoli cells. The exact timing of these events varies from individual to individual, but mature sperm are almost always present by 14 years of age, and some of the features may begin to manifest at around 9 years of age. This results in a dramatic change in the testicular histology, including a dramatic enlargement of tubular diameter, a dramatic increase in the number of germ

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Fig. 10.36  Early pubertal testis. Shown here is the seminiferous tubule from a 12-year-old boy. Rare spermatogonia are present along the basement membrane (“1”). In comparison with the prepubertal testis, this testis has a higher number of primary spermatocytes (“2”), which almost form a cluster toward the center of the tubule and are a prominent feature in this testis, indicating active spermiogenesis. Observe the less spindly appearance of the Leydig cells in the intertubular region, indicating responsiveness to hormonal stimulation (Toluidine blue, 60×)

D. S. Huff and C. K. Carreon

Fig. 10.37  Pubertal testis. Shown here is the seminiferous tubule from a 15-year-old boy. Rare scattered intermediate cells are present. In comparison with the early pubertal testis, this testis, in addition to demonstrating numerous primary spermatocytes (“1”), also shows early and late spermatids (“2”) indicating achievement of the last stages of spermiogenesis, which is a prominent feature of the pubertal testis (Toluidine blue, 60×)

cells, enlargement of Sertoli cells, and an increased number and size of adult Leydig cells (Figs. 10.36 and 10.37).

Special Considerations Paramesonephric and mesonephric embryonic remnants are often described as being distinctly different. Paramesonephric remnants (appendix testis) are said to be solid and covered by prominent cuboidal and columnar epithelium. Mesonephric remnants (appendix epididymis) are said to be cystic and covered by flat, inconspicuous mesothelium. Although these definitions are sometimes true, the descriptions below emphasize that the two may be impossible to differentiate by their histologic appearances.

Paramesonephric Remnants

Fig. 10.38  Solid filiform appendix testis. The fingerlike appendix is attached to the testicular capsule and points to the left. It is covered with simple, low cuboidal epithelium. Ap appendix testis, Ep head of the epididymis, Cp capsule of the testis, Te upper pole of the testis (H&E, 4×)

The appendix testis, sometimes called the hydatid of Morgagni, is thought to be a remnant of the paramesonephric duct and is described by some as the homologue of the fimbriated end of the Fallopian tube (Figs. 10.38, 10.39, and 10.40). It is attached

to the upper pole of the testis and may be polypoid or filiform, sessile, or pedunculated. It is usually solid but may be cystic or may contain a tubule. The cyst or tubule may communicate with the surface, similar to a fimbria. The appendix is often covered

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Fig. 10.39  Cystic appendix testis. The appendix contains a central cyst and is attached by a thin pedicle to the capsule of the testis. The epithelium covering the surface and lining the cyst is similar (shown in Fig. 10.40). The capsule of the testis includes the tunica vaginalis, the tunica albuginea, and the tunica vasculosa. Ap appendix testis, Cp capsule of testis, Ep head of epididymis, Pd remnant of paramesonephric duct, Te upper pole of testis (H&E, 4×)

Fig. 10.40  Epithelium of the appendix testis. This is a higher-power image of the epithelium that covers the surface of the appendix testis shown in Fig. 10.39. The epithelium is pseudostratified columnar and cuboidal, with patches of cilia (H&E, 60×)

Fig. 10.41  Cystic, sessile appendix epididymis. The appendix (Ap) is attached to the head of the epididymis (Ep). The epithelium lining the cyst is similar to that in Figs. 10.40 and 10.43 (H&E, 4×)

Fig. 10.42  Solid, pedunculated appendix epididymis. The appendix (Ap) is attached to the head of the epididymis (Ep) by a short pedicle. The surface epithelium is shown in Fig. 10.43. Te upper pole of testis (H&E, 2×)

by cuboidal to columnar, focally ciliated, and basophilic epithelium. If it is cystic, the cyst is usually lined by similar epithelium. The surface epithelium is sometimes flat mesothelium. The stroma may contain scattered smooth muscle fibers and, if tubular, may demonstrate a muscularis [3, 22, 23].

of the mesonephric duct (Figs. 10.41, 10.42, and 10.43). It is polypoid or filiform, pedunculated, or sessile. It is usually cystic or contains a rudimentary tubule that usually does not communicate with the surface. It is usually covered by simple, flat epithelium similar to the adjacent covering of the tunica albuginea, but it may be covered by an epithelium identical to that typical of an appendix testis. If it is cystic, the cyst is lined by simple cuboidal or columnar epithelium that may be focally ciliated. This description is almost identical to that of the appendix testis. Differentiation between an appendix testis and an appendix epididymis may be impossible by histological criteria alone and may depend on location. Some

Mesonephric Remnants Several mesonephric remnants are often present [3, 22, 23]. The appendix epididymis is attached to the head of the epididymis and is thought to be a remnant of the blind rostral end

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Fig. 10.43  Surface epithelium of the appendix epididymis shown in Fig. 10.42. This epithelium is identical to that of the cystic appendix testis seen in Fig. 10.40 (H&E, 60×)

Fig. 10.44  Embryonic rests in the distal spermatic cord. These mesonephric embryonic rests (arrows) are found in the distal spermatic cord adjacent to or rostral to the head of the epididymis. In this fetus, they appear to arise from the epigenital mesonephric nephrons related to those that form the efferent ductules. These rests are intermingled with the vessels of the pampiniform plexus entering the distal spermatic cord. BEp body of epididymis, Gu gubernaculum, HEp head of epididymis, SC cut end of spermatic cord (H&E, 1×)

Fig. 10.45  Embryonic rests with glomeruloid structures in the distal spermatic cord. Ep, edge of the epididymis; Me, a nest of tubular and glomeruloid mesonephric embryonic remnants at the edge of the pampiniform plexus in the distal end of the spermatic cord; Te, hilum of the testis; VD, vas deferens, as it nears the distal end of the spermatic cord (H&E, 2×)

Fig. 10.46  Higher magnification of the embryonic rests in Fig. 10.45. The rest is composed of mesonephric tubular and glomerular structures (H&E, 10×)

appendages are attached at the junction between the epididymis and the testis; these may be difficult to classify. Blind-ended remnants of mesonephric collecting ducts, which remain attached to the mesonephric duct near the upper and lower poles of the testis, are called superior and inferior aberrant ducts, respectively, or organs of Haller. They are analogous to the epoophoron in the female and are inconspicuous. Ductular remnants seen between the epididymis and testis may be either mesonephric or paramesonephric in origin. Ductular remnants associated with the vas deferens in the distal end of the spermatic cord are the remnants of the para-

genital mesonephric nephrons and have been called the paradidymis or organ of Giraldés (Fig. 10.44). They are analogous to the paroophoron, are lined by simple cuboidal or columnar epithelium, and are accompanied by sparse, smooth muscle fibers. In our material, some of the embryonic remnants in the pampiniform plexus near the hilum, adjacent to the head of the epididymis, or in the distal spermatic cord appear to arise from partially degenerated epigenitalis mesonephric nephrons, which stream from the testicular hilum into the pampiniform plexus rostrally toward the spermatic cord. Any of these mesonephric ductular r­ emnants may be associated with mesonephric glomeruloid structures (Figs. 10.45 and 10.46).

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Fig. 10.47 Shown here is a testis from a normally developed 38-weeks-­gestation neonate. A strong positive immunohistochemical staining for CD10 is demonstrated by the epididymis (left upper corner), and a much weaker and patchy positive staining is observed in the rete testis (upper right). Both of these structures, along with the efferent ducts, ductus deferens, and seminal vesicles, are mesonephric duct/Wolffian duct derivatives. In contrast, the seminiferous tubules show diffusely absent staining for CD10 (CD10 immunohistochemistry, 4×)

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Fig. 10.48  Adrenal cortical rest of Marchand at the junction of the rete testis and the efferent ducts. The adrenal rest is outlined by the arrows; it is composed of definitive and fetal adrenal cortex. At the asterisk, the channels of the extratesticular compartment of the rete testis intermingle with the adrenal rest. ED efferent ductules, RT extratesticular compartment of the rete testis (H&E, 4×)

Similar embryonic remnants are found in the spermatic cord and in the wall of hernial sacs, where they may be confused with vas deferens or epididymis. Recent evidence that epithelium of the mesonephric duct (Wolffian duct) derivatives may be positive for CD10, whereas paramesonephric duct (Müllerian duct) epithelium is not, may provide an immunohistochemical method for determining the origin of the embryonic remnants discussed above (Fig. 10.47) [24].

Adrenal Cortical Rests Adrenal cortical rests or ectopic adrenal nodules of Marchand may be found in the testis, the rete testis (Fig. 10.48), the testicular hilum, the epididymis, the paratesticular soft tissue, or the spermatic cord. They are encapsulated and composed of definitive and fetal cortex without the medulla. They may arise from a rostral area of mesonephric mesodermal epithelium, which gives rise to both adrenal and testicular epithelium. They may be affected by the same lesions as fetal adrenal cortex, such as adrenal cortical cytomegaly, cytomegalic inclusion disease, and other infections.

Testicular Embryonic Remnants Several embryonic remnants may be found in and around the testis. Nests of testicular blastema or clusters of seminiferous

Fig. 10.49  Ectopic germ cells in the cuboidal epithelium of the tunica vaginalis and in the tunica albuginea. The ectopic germ cells sometimes form clusters, or they may be single. They disappear with advancing gestational age (H&E, 60×)

tubules are occasionally present in the tunica albuginea [6, 7]. On occasion, these may extend onto the surface as grossly visible reddish-brown spots, giving the testis an appearance similar to a speckled bird’s egg. Whether these remnants should be considered congenital anomalies or variations of normal is unclear. They should be distinguished from testicular dysgenesis. Solitary germ cells may be found in the tunica albuginea (Fig. 10.49), testicular interstitium, or rete testis. They are seen infrequently and more likely during midpregnancy than at other periods of fetal life. Ectopic Leydig cells may be present in the tunica albuginea or in paratesticular connective tissue and nerves (Fig.  10.50); these are often

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Fig. 10.50  Ectopic Leydig cells in nerves of the paratesticular soft tissue near the hilum. These may be confused with ganglion cells (H&E, 60×)

numerous in conditions associated with hyperplasia of Leydig cells. They may mimic ganglion cells. Acknowledgments  The authors thank Dr. Catherine T.-S. Chung from The Hospital for Sick Children, Toronto, Canada, for invaluable assistance in selecting microscopic slides for photography, taking photomicrographs, and editing the original manuscript.

References 1. Satoh M. Histogenesis and organogenesis of the gonad in human embryos. J Anat. 1991;177:85–107. 2. Jirásek JJ. An atlas of human prenatal developmental mechanics: anatomy and staging. Boca Raton: Parthenon; 2004. 3. Felix W. The development of the urinogenital organs. In: Keibel F, Mall FP, editors. Human embryology. Philadelphia: JB Lippincott; 1912. p. 752–979. 4. van Wagenen G, Simpson M. Embryology of the ovary and testis, Homo sapiens and Macaca mulatta. New Haven: Yale University Press; 1965. 5. O’Rahilly R. The timing and sequence of events in the development of the human reproductive system during the embryonic period proper. Anat Embryol (Berl). 1983;166:247–61. 6. Cajaiba MM, Garcia-Fernandez E, Reyes-Mugica M, Nistal M. The spectrum of persistence of testicular blastema and ectopic testicular parenchyma: a possible result of focal delay in gonadal development. Virchows Arch. 2007;451:89–94.

D. S. Huff and C. K. Carreon 7. Nistal M, Panigua R.  Non-neoplastic diseases of the testes. In: Bostwick D, Cheng L, editors. Urologic surgical pathology. 2nd ed. Philadelphia: Elsevier; 2008. p. 616–755. 8. Falin LI.  The development of genital glands and the origin of germ cells in human embryogenesis. Acta Anat (Basel). 1969;72: 195–232. 9. Fukuda T, Hedinger C, Groscurth P.  Ultrastructure of developing germ cells in the fetal human testis. Cell Tissue Res. 1975;161:55–70. 10. Culty M. Gonocytes, the forgotten cells of the germ cell lineage. Birth Defects Res C Embryo Today. 2009;87:1–26. 11. Hermo L, Pelletier RM, Cyr DG, Smith CE.  Surfing the wave. Cycle, life history, and genes/proteins expressed by testicular germ cells. Part 1: background to spermatogenesis, spermatogonia, and spermatocytes. Microsc Res Tech. 2010;73:241–78. 12. Svechnikov K, Landreh L, Weisser J, Izzo G, Colón E, Svechnikova I, Söder O. Origin, development and regulation of human Leydig cells. Horm Res Paediatr. 2010;73:93–101. 13. Waters BL, Trainer TD.  Development of the human fetal testis. Pediatr Pathol Lab Med. 1996;16:9–23. 14. Hadziselimovic F.  Testicular development. In: Gillenwater JY, Grayhack JT, Howards SS, Duckett JW, editors. Adult and pediatric urology. 2nd ed. St Louis: Mosby; 1987. p. 2173–88. 15. Wartenberg H, Holstein AF, Vossmeyer J. Cytology of the prenatal development of human gonads. II. Electronmicroscopic studies on the cytogenesis of gonocytes and foetal spermatogonia in the testis. Z Anat Entwicklungsgesch. 1971;134:165–85. 16. Wartenberg H. Human testicular development and the role of the mesonephros in the origin of a dual Sertoli cell system. Andrologia. 1978;10:1–21. 17. Gaskell TL, Esnal A, Robinson LL, Anderson RA, Saunders PT. Immunohistochemical profiling of germ cells within the human fetal testis: identification of three subpopulations. Biol Reprod. 2004;71:2012–21. 18. Franke FE, Pauls K, Rey R, Marks A, Bergmann M, Steger K.  Differentiation markers of Sertoli cells and germ cells in fetal and early postnatal human testis. Anat Embryol (Berl). 2004;209:169–77. 19. Helal MA, Mehmet H, Thomas NS, Cox PM, Ralph DJ, Bajoria R, Chatterjee R. Ontogeny of human fetal testicular apoptosis during first, second, and third trimesters of pregnancy. J Clin Endocrinol Metab. 2002;87:1189–93. 20. Yasuda Y. Testis. In: Nishimura H, editor. Atlas of prenatal histology. Tokyo: Izaku-Shoin; 1983. p. 148–51. 21. Clermont Y.  Renewal of spermatogonia in man. Am J Anat. 1966;118:509–24. 22. Willis R.  The borderland of embryology and pathology. London: Butterworth; 1958. 23. Hamilton W, Mossman HW.  Hamilton, Boyd, and Mossman’s human embryology. 4th ed. Baltimore: Williams and Wilkins; 1972. 24. Cerilli LA, Sotelo-Avila C, Mills SE.  Glandular inclusions in inguinal hernia sacs: morphologic and immunohistochemical distinction from epididymis and vas deferens. Am J Surg Pathol. 2003;27:469–76.

Epididymis

11

Dale S. Huff and Eduardo D. Ruchelli

Introduction

The efferent ductules and head of the epididymis are derived from the collecting ductules of 5–12 of the most rosThe primary function of the epididymis is to provide a con- tral degenerating epigenital mesonephric nephrons. The duit for the transport of sperm and seminiferous fluid from body and tail of the epididymis are derived from the mesothe rete testis to the vas deferens, but the epididymis per- nephric duct. When the embryonic period closes at the end of forms much more elaborate functions, including the resorp- the tenth postmenstrual week, the rudimentary epididymis is tion of some of the fluid by the head of the epididymis, final composed of 5–12 straight efferent ductules, which will maturation and development of sperm motility in the body, form the head. Each efferent ductule is attached to the rostral and storage of sperm in the tail. Performance of these func- part of the mesonephric duct, which descends along the postions requires the development of an elaborate rostral-to-­ terolateral hilar border of the testis to the lower pole, and caudal gradient of epithelial specialization from the efferent then curves medially under the lower pole. The segment of ductules to the beginning of the vas deferens, which is not the mesonephric duct from the junction with the efferent completed until puberty. Knowledge of the changing fetal ducts to the lower pole of the testis will become the body of histology described in this chapter aids in the understanding the epididymis. The segment that curves under the lower of the congenital anomalies of the epididymis that are seen in pole of the testis will become the tail of the epididymis. association with cryptorchidism [1], renal and ureteral Histologically, the mesonephric duct is lined by simple, anomalies, cystic fibrosis, and other underlying conditions. nonciliated, columnar epithelium [2], and its wall is composed of concentric layers of undifferentiated mesenchyme, which will become the circular muscular coat. In fetuses at Embryology 11 weeks gestation, the undifferentiated mesenchyme of the caudal end of the mesonephric duct becomes organized into The early embryology of the genitourinary system and the concentric layers, and this transformation extends rostrally mesonephric duct is described in the Part IV introduction, into the epididymis. Cilia begin to appear in fetuses at and the development of the rete testis and efferent ductules is approximately 11–12 weeks gestation [2]. described in Chap. 10. The development of the epididymis is During the fetal period, the simple, straight efferent ductone part of the rostral-to-caudal segmental differentiation of ules and straight ductus epididymis become elaborately the mesonephric duct into its various components: the head, coiled, with a rostral-to-caudal gradient in the degree of coilbody, and tail of the epididymis; the vas deferens; the ampulla ing, epithelial specialization, and thickness of the muscular of the vas deferens; the seminal vesicles; and the ejaculatory coat. At 11 weeks gestation, the solid tubules of the epigeniducts. This segmental differentiation is similar to that seen in tal mesonephric collecting ductules connect with the solid the gastrointestinal tract and is under elaborate genetic con- cords of the rete. This is called the urogenital union. The trol. The viability and maturation of all the structures is epigenital ductules are now called the efferent ductules, and androgen-dependent. the mesonephric duct is called the ductus epididymis. Each efferent ductule connects to the upper end of the ductus epididymis. The efferent ductules and their connections to the D. S. Huff · E. D. Ruchelli (*) rete testis and to the ductus epididymis begin to develop Department of Pathology and Laboratory Medicine, The Children’s lumens in the third trimester, a process that is not complete Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA until after birth. e-mail: [email protected]; [email protected]

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_11

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Histology: General Overview The epididymis is the first segment of the excretory system of the testis. It is “C”-shaped and fits around the upper pole, the posterolateral or hilar margin, and the lower pole of the testis. The bulbous head (caput) of the epididymis is tightly attached to and pressed into the upper pole of the testis. The body (corpus) is joined to the hilar margin of the testis by a short mesentery, the mesorchium. The tail (cauda) curves under and is tightly fused to the lower pole of the testis by the fibrous remains of the testicular ligament and gubernaculum. The head is joined to the rete testis by the efferent ductules, and the tail is continuous with the vas deferens. The head and all but the most caudal end of the body are intraperitoneal and are covered with a fibrous capsule similar to, but not as well developed as, the tunica albuginea of the testis. The capsule is covered by simple, flat mesothelial epithelium derived from the visceral epithelium of the processus vaginalis. The caudal end of the body and the tail are retroperitoneal and are buried in the connective tissue remnants of the testicular ligament and gubernaculum [3, 4]. The head is composed of 5–12 lobules called coni vasculosi and a second, ill-defined region that connects the coni vasculosi with the body. Each of the coni vasculosi is composed of one extensively coiled efferent ductule. One end of each efferent ductule is connected to the extratesticular portion of the rete testis, and the other end is connected to the ductus epididymis (formerly the mesonephric duct). The coni vasculosi are separated by fibrous septa, and the coils of the efferent ductules within the coni vasculosi are embedded in an interstitium of connective tissue. The body and tail of the epididymis are composed of the single, coiled ductus epididymis, which is continuous caudally with the vas deferens. The coils of the efferent ductules and ductus epididymis in the body and tail are all set in an interstitium of connective tissue. The histology of the adult human epididymis has been extensively described [5–7]. The rostral-to-caudal gradient in specialization of epithelial histology and function in adults has been shown to be much more elaborate than previously thought. It has been divided into seven segments: anterior and posterior efferent ductules; anterior, middle, and posterior body; and anterior and posterior tail [5]. The histologic organization is similar throughout all the multiple segments of the epididymis. The epithelium lining the ductules lies upon a basement membrane, which is surrounded from the inside out by spindle-shaped periductular myoid cells, an inconspicuous lamina propria, and a circular layer of smooth muscle. The epithelium is pseudostratified cuboidal to columnar and has stereocilia, but the epithelium varies in a rostral-to-caudal gradient. In the head, the pseudostratified epithelium is composed of short, cuboidal, nonciliated cells and tall, columnar cells with stereocilia. Five ultrastructurally distinct zones have been described within the ciliated epithelium lining the efferent ductules [6]. In the body, the

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pseudostratified columnar epithelium consists of four types of cells: tall, columnar stereociliated principal cells; tall, slender, apical mitochondria-rich cells; clear cells; and basal cells. The length of the stereocilia varies gradually in a rostral-­to-caudal gradient: They are longest at the rostral end of the ductus epididymis, where they nearly fill the lumen; they are shortest at the caudal end of the tail. The thickness of the circular muscle coat varies in a rostral-to-caudal gradient: It is thinnest in the head and thickest in the tail. Longitudinal fibers appear in the tail [8].

Fetal Histology The changing histology of the fetal epididymis is due to increased coiling, maturation of the epithelium, and development of the muscular coat, each of which varies in a rostral-­to-­ caudal gradient. The histological plan of the fetal epididymis is similar to that of the adult. The primitive, pseudostratified columnar epithelium lies on a basement membrane that is surrounded by one or two layers of spindle-shaped myoid or myofibroblastic cells, which is in turn surrounded by a cuff of swirled, primitive mesenchyme. The primitive mesenchyme differentiates into the lamina propria, the circular smooth muscle coat, and the adventitia. The lamina propria and adventitia are inconspicuous, but the smooth muscle becomes prominent and important. Immunohistochemical studies of inhibin [9], androgen and estrogen receptors [10], epithelial membrane antigen, cytokeratins, and estradiol-­related protein receptors [11] in the fetal epididymis have been reported, but the histology of the fetal epididymis has not been reported in a large series of cases from multiple gestational ages. At 13 weeks, through the developmental process known as tubulogenesis [12], the distal ends of the efferent ducts close to their attachment to the ductus epididymis begin to coil and form the coni vasculosi. The proximal ends at the rete remain straight and parallel. Each coiled duct forms a pyramidal lobule, with its apex at the testicular hilum and its base toward the mesonephric duct. These are the coni vasculosi; collectively, they form the head of the epididymis. The body and tail begin as the uncoiled mesonephric duct, which becomes the uncoiled ductus epididymis. When the efferent ductules begin to coil, the ductus epididymis of the body and tail also elongates and coils through coiling tubulogenesis. The coiling follows a rostral-to-caudal gradient and is most extensive in the efferent ductules, less extensive in the body, and least extensive in the tail. This coiling results in a rostral-to-caudal gradient in the diameter of the epididymis; the diameter of the head is the largest, and that of the tail is the smallest. The coiling of the epididymis continues throughout fetal life and postnatal life and causes a dramatic increase in the length of the tube. Histologically, increased coiling results in increased numbers of ductular cross sections, which become more closely packed.

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Fig. 11.1  Epididymis at 17 weeks gestation. This image is a longitudinal section of the epididymis and testis. The epididymis is very primitive, and the thin connective tissue capsule is visible. The head (H) of the epididymis is bent away from the testis (TE). The lobules formed by the individual efferent ductules are faintly visible in the head. The efferent ductules are slightly coiled. The body (B) and tail (T) of the epididymis are even less coiled than the efferent ductules, and the path of the gently curved mesonephric duct, which is differentiating into the epididymis, can be seen in the body and tail. The vas deferens (V) is straight. The peritubular swirls of dark blue, primitive mesenchyme, which will become the stroma and smooth muscle of the epididymis, are seen around the tubules. PP pampiniform plexus (Hematoxylin and eosin (H&E), 4×)

Before about the middle of the second trimester, coiling is not extensive. The diameter of the head is only slightly larger than that of the body, and it is bent away from the upper pole of the testis and is not attached to it. The ductus epididymis of the body and tail shows even less coiling and is gently undulating. The simple, ciliated, columnar epithelium of the embryonic efferent ductules and ductus epididymis has differentiated into primitive, pseudostratified stereociliated cuboidal and columnar epithelium, which is similar from the head to the tail. The primitive periductular mesenchyme is thinnest in the head and thickest in the tail, but it shows no evidence of smooth muscle differentiation (Figs. 11.1, 11.2, 11.3, and 11.4). After the middle of the second trimester, the size of the head of the epididymis grows rapidly due to the extensive coiling of the efferent ductules. Within a few weeks, the head enlarges into a cap-shaped structure that is bent over and pressed into the upper pole of the testis, to which it becomes tightly attached. The gestational age at which the head is attached to the upper pole of the testis has not been established with certainty; the attachment probably varies by several weeks from fetus to fetus, but in some cases it occurs shortly after 20 weeks gestation. The attachment of the head of the epididymis to the upper pole of the testis is best visualized in complete longitudinal sagittal sections of the testis and epididymis. The increased coiling results in more numerous and more tightly packed ductular cross sections. The epithelium continues to mature but remains primitive and similar

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Fig. 11.2  Efferent ductules forming the head of the epididymis at 17 weeks gestation. The visceral mesothelial epithelium of the processus vaginalis covers the thin, fibrous capsule, similar to the visceral mesothelial epithelium of the tunica vaginalis covering the tunica albuginea of the testis. The ductules are lined by crowded, simple columnar epithelium and are surrounded by concentric swirls of primitive mesenchyme. The luminal surface is slightly irregular, and apical blebs and occasional stereocilia are recognizable in this imperfectly preserved section (H&E, 40×)

Fig. 11.3  Body of the epididymis at 17 weeks gestation. This image is a higher magnification of the body of the epididymis shown in Fig. 11.1. The histology of the body is nearly identical to that of the head. The cross sections of the tubule are slightly smaller and more round than those of the efferent ductules. The simple cuboidal epithelial cells are very crowded. The luminal surface is sharply outlined, and terminal bars appear (H&E, 40×)

throughout the length of the epididymis. Smooth muscle has been reported to appear in the primitive periductular mesenchyme at approximately 19–20 weeks gestation [2], but the time of the appearance of smooth muscle differentiation may vary; in some cases, it is not seen until later in gestation (Figs. 11.5, 11.6, 11.7, 11.8, 11.9, 11.10, and 11.11). During the second trimester, coiling continues to increase throughout the epididymis. By the end of the second trimester or the beginning of the third, the primitive epithelium dif-

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Fig. 11.4  Tail of the epididymis at 17 weeks gestation. This image is a higher magnification of the tail seen in Fig.  11.1. The histological appearance of the tail is identical to that of the body (H&E, 20×)

Fig. 11.6  Coni vasculosi of the head of the epididymis at 22  weeks gestation. This image is a higher magnification of the coni vasculosi shown in Fig.  11.5. The cross sections of the tightly coiled efferent ductules are closely packed and back-to-back. The periductular primitive mesenchyme is thin (H&E, 10×)

ferentiates into epithelium that is characteristic of epididymis. The epithelium throughout the epididymis increases in thickness. Principal cells and apical mitochondria-rich cells are present in the head, body, and tail of the epididymis by 24–27 weeks gestation [11]. Basal cells and clear cells may

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Fig. 11.5  Epididymis at 22 weeks gestation. This image shows a longitudinal section of the entire epididymis and testis. The enlarged head (H) is bent around and pressed against the upper pole of the testis (TE). Coiling of the efferent ductules in the head has increased dramatically. The pyramidal coni vasculosi of five or six efferent ductules are sharply outlined by septa. The testicular end of each is only slightly coiled and therefore is narrow, whereas the opposite end is very coiled and therefore is wide. The periductular mesenchymal cuffs are indistinct. The demarcation between the head and body (B) (arrow) is vague. The coiling of the body has increased, but not as dramatically as in the head. The peritubular mesenchymal cuffs are thick and condensed. Because of the extensive coiling, the diameters of the head and body have increased. The tail (T) is less coiled than the body, but the peritubular mesenchymal cuffs of the tail and the vas deferens (V) are thicker and more condensed than in the body (H&E, 1×)

Fig. 11.7  Coni vasculosi of the head of the epididymis at 22 weeks gestation. This is a higher magnification of the efferent ductules shown in Fig. 11.6. The nuclei of the cuboidal epithelial cells are irregular in size and shape and are becoming pseudostratified. A layer of spindled myoid cells one or two cells thick surrounds the epithelium, and a thin irregular layer of primitive mesenchyme surrounds the layer of myoid cells (H&E, 40×)

be difficult to recognize in routine histological sections. In the head, the outline of the lumen becomes irregular, and the epithelium is pseudostratified cuboidal and columnar. The tall columnar cells have stereocilia, whereas the cuboidal

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Fig. 11.8  Body of the epididymis at 22 weeks gestation. This image is a higher magnification of the body shown in Fig. 11.5. The cross sections of the ductus epididymis are more widely spaced than those of the efferent ductules, because the ductus epididymis is less coiled than the efferent ductules. The primitive periductular mesenchyme is much thicker than that of the efferent ductules (H&E, 10×)

Fig. 11.9  Body of the epididymis at 22 weeks gestation. This image is a higher magnification of the body shown in Fig. 11.8. Occasional tall, thin columnar cells are interspersed between the tightly packed cuboidal cells, especially in the tubule in the right lower corner. These cells may be primitive apical mitochondria-rich cells. The peritubular mesenchyme is composed of spindle cells, but smooth muscle differentiation is not seen (H&E, 40×)

Fig. 11.10  Tail of the epididymis at 22 weeks gestation. This image is a higher magnification of the tail shown in Fig.  11.5. The tail is less coiled than the body, and the peritubular mesenchyme is thicker than that of the body (H&E, 10×)

Fig. 11.11  Tail of the epididymis at 22 weeks gestation. This image is a higher magnification of the tail shown in Fig. 11.10. The epithelium is similar to that of the body, but it lacks the tall, thin columnar cells seen in the body (H&E, 40×)

surface cells are not ciliated. Smooth muscle differentiation within the primitive periductular mesenchyme is not typically recognizable in routine sections of the head. In the body, the outline of the lumen is smooth and round. The mucosa is thicker than in the head, and the epithelium is composed of tall, wide stereociliated principal cells and tall, slender, dark apical mitochondria-rich nonciliated cells. As is the case in the head, smooth muscle differentiation within the primitive periductular mesenchyme is usually not recognizable in the body. In the tail, the mucosa is similar to but thinner than that in the body. The periductular cuff of primi-

tive mesenchyme is thicker than in the body and head. Circular smooth muscle cells that are thin and delicate differentiate within the periphery of the primitive periductular mesenchyme. Longitudinal bundles of large smooth muscle cells appear around the tail of the epididymis and the proximal vas deferens. For the most part, these longitudinal bundles are separate from the smooth muscle differentiating within the periductular mesenchyme, but in some areas they swirl around and blend into the periductal and vascular smooth muscle. These longitudinal bundles are not much discussed or illustrated in the literature but may be the longi-

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Fig. 11.12  Epididymis at 34 weeks gestation. This image shows a longitudinal section of the entire epididymis and testis. The head (H) of the epididymis is bent away from the upper pole of the testis. The head of this epididymis is less well developed than the epididymis at 22 weeks gestation, shown in Fig. 11.5. This figure illustrates the wide range of the timing of developmental events. Otherwise, little has changed in the low-magnification appearance of the epididymis. The arrow indicates the indistinct junction between the head and the body of the epididymis. B body of epididymis, T tail of epididymis, TE testis, V vas deferens (H&E, 1×)

Fig. 11.13  Head of the epididymis at 34 weeks gestation. This image is a longitudinal section of the head shown in Fig. 11.12. The coni vasculosi are separated by fine interlobular septa. The efferent ductules are more extensively coiled than previously, and their profiles are curved (H&E, 4×)

Fig. 11.14  Efferent ductules at 34  weeks gestation. This image is a higher magnification of the efferent ductules shown in Fig. 11.13. The epithelium is becoming more columnar and pseudostratified. The broad columnar cells are principal cells, and occasional tall, thin columnar cells reach from the basement membrane to the lumen. These cells are apical mitochondrial-rich cells. Some stereocilia are preserved, and the outline of the lumen is slightly irregular. There is a peritubular layer of spindle-shaped myoid cells and a substantial layer of peritubular mesenchyme (H&E, 40×)

Fig. 11.15  Body of the epididymis at 34 weeks gestation. This image is a higher magnification of the body seen in Fig. 11.12. The ductule is lined by pseudostratified columnar epithelium. The mesenchyme forms a dense layer of spindle cells, but histologically identifiable smooth muscle is not present (H&E, 10×)

tudinal muscle referred to above (Figs. 11.12, 11.13, 11.14, 11.15, 11.16, 11.17, and 11.18) [8]. By the end of the third trimester, the epithelial maturation has increased. The dramatic rostral-to-caudal gradient in the length of cilia that is seen in adults, with very long cilia ros-

trally and short cilia caudally, is not prominent in fetuses. Differentiation of smooth muscle within the primitive periductal mesenchyme of the tail and the caudal portion of the body continues. Smooth muscle differentiation may not be present in the mid and rostral body and the head of the epididymis at the end of the third trimester (Figs. 11.19, 11.20, 11.21, 11.22, 11.23, and 11.24).

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Fig. 11.16  Body of the epididymis at 34 weeks gestation. This image is a higher magnification of the body seen in Fig. 11.15. Compared with efferent ductules of the head, the ductus of the body has a more regular round lumen, a taller pseudostratified columnar epithelium with the nuclei confined mostly to the lower half of the thickness of the epithelium, and more apical mitochondria-rich cells. The pseudostratified tall columnar epithelium rests on a thin basement membrane. Of the four types of epithelial cells described in the adult, principal cells and apical mitochondria-rich cells are present. Two types of apical mitochondria-­ rich cells are seen: the tall, slender, dark cells that extend from the basement membrane to the lumen and the small cells at the lumen that contain dark, oval nuclei in the upper half of the epithelium. The remainder are principal cells. Clear cells and basal cells are difficult to identify in this preparation. Smooth muscle is not present (H&E, 40×)

Fig. 11.17  Tail of the epididymis at 34 weeks gestation. This image is a higher magnification of the tail seen in Fig. 11.12. This photomicrograph is from the distal end of the tail, close to its junction with the vas deferens; therefore, it is only slightly coiled. Smooth muscle is present. The most well-differentiated smooth muscle is organized into broad bundles (asterisks), which partially surround the outside of the epididymis like a capsule and extend around the outside of the proximal vas deferens. The muscle cells forming the bundles are unusually large. They sometimes swirl around the ductule and adjacent vessels and blend in with the outer layers of the periductular mesenchyme and the vascular smooth muscle. The significance of this smooth muscle is not clear. The peritubular cuffs of spindled mesenchymal cells are very condensed and much thicker than those in the body, but only the outer layers are recognizable smooth muscle (H&E, 10×)

Fig. 11.18  Tail of the epididymis at 34 weeks gestation. This image is a higher magnification of the tail seen in Fig. 11.17. The multilayered, pseudostratified epithelium is cuboidal rather than columnar, as in the body. Numerous stereocilia are preserved in this section. The peritubular mesenchyme is vaguely divided into two or three layers, of which the outer layer is recognizable as smooth muscle (H&E, 40×)

Fig. 11.19  Head of the epididymis at 39 weeks gestation. The epididymis is surrounded by a fibrous capsule, which is covered by a layer of simple, flat mesothelial epithelium. The coni vasculosi are separated by connective tissue septa, and the efferent ductules are tightly coiled. The periductular mesenchymal cuffs are thicker and more condensed than in younger fetuses (H&E, 4×)

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Fig. 11.20  Efferent ductules of the head of the epididymis at 39 weeks gestation. This image is a higher magnification of the efferent ductules seen in Fig. 11.19. The lumens are large and slightly irregular. The epithelium is thin and single-layered in some foci. A few apical mitochondria-­rich cells of both types are present. Smooth muscle differentiation is recognizable in the peritubular mesenchyme (H&E, 40×)

Fig. 11.21  Body of the epididymis seen at 39 weeks gestation. This image of the body is from the same specimen shown in Figs. 11.19 and 11.20. The fibrous capsule is thick and covered by mesothelium. The duct of the epididymis is more tightly coiled than previously, and the lobules are separated by septa (H&E, 4×)

Fig. 11.22  Body of the epididymis at 39 weeks gestation. This image is a higher magnification of the body seen in Fig. 11.21. The lumens are smaller and more round than those in the head. Principal and apical mitochondria-rich cells and basal cells are present. The basement membrane and peritubular myoid cells are prominent, and smooth muscle cells are recognized in the peritubular mesenchyme (H&E, 40×)

Fig. 11.23  Tail of the epididymis at 39 weeks gestation. This image of the tail is from the same case seen in Fig. 11.19. This section is from the gradual transition from the body (B) in the upper left to the tail (T) in the lower half of the picture. Smooth muscle is seen in the periphery of the periductular mesenchyme of some cross sections of the duct (H&E, 4×)

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Fig. 11.24  Tail of the epididymis at 39 weeks gestation. This image is a higher magnification of the tail seen in Fig. 11.23. The lumen is lined by pseudostratified cuboidal and columnar epithelium with stereocilia. The distinction between principal and apical mitochondria-rich cells is not as clear as in the body (H&E, 40×)

References 1. De Miguel MP, Mariño JM, Gonzalez-Peramato P, Nistal M, Regadera J.  Epididymal growth and differentiation are altered in human cryptorchidism. J Androl. 2001;22:212–25. 2. Felix W. The development of the urinogenital organs. In: Keibel F, Mall FP, editors. Human embryology. Philadelphia: JB Lippincott; 1912. p. 752–979.

155 3. Hadziselimovic F.  Cryptorchidism. Berlin: Springer-Verlag; 1983. 4. Hadziselimovic F, Kruslin E.  The role of the epididymis in descensus testis and the topographical relationship between the testis and epididymis from the sixth month of pregnancy until immediately after birth. Anat Embryol (Berl). 1979;155: 191–6. 5. Dacheux JL, Belghazi M, Lanson Y, Dacheux F. Human epididymal secretome and proteome. Mol Cell Endocrinol. 2006;250: 36–42. 6. Yeung CH, Cooper TG, Bergmann M, Schulze H. Organization of tubules in the human caput epididymidis and the ultrastructure of their epithelia. Am J Anat. 1991;191:261–79. 7. Trainer T. The testis and excretory duct system. In: Mills SE, editor. Histology for pathologists. 3rd ed. Philadelphia: Lippincott Williams & Wilkins; 2007. p. 943–63. 8. Ladman A.  The male reproductive system. In: Greep RO, Weiss L, editors. Histology. 3rd ed. New  York: McGraw-Hill; 1973. p. 847–90. 9. Nistal M, Gonzalez-Peramato P, De Miguel MP. Immunodetection of inhibin in the human testis and epididymis during normal development and in non-tumoural testicular lesions. Reprod Fertil Dev. 2010;22:558–63. 10. Shapiro E, Huang H, Masch RJ, McFadden DE, Wu XR, Ostrer H.  Immunolocalization of androgen receptor and estrogen receptors alpha and beta in human fetal testis and epididymis. J Urol. 2005;174:1695–8; discussion 1698 11. Regadera J, Cobo P, Paniagua R, Martínez-García F, Palacios J, Nistal M. Immunohistochemical and semiquantitative study of the apical mitochondria-rich cells of the human prepubertal and adult epididymis. J Anat. 1993;183:507–14. 12. Joseph A, Yao H, Hinton BT.  Development and morphogenesis of the Wolffian/epididymal duct, more twists and turns. Dev Biol. 2009;325:6–14.

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Vas Deferens Linda M. Ernst, Eduardo D. Ruchelli, and Dale S. Huff

Introduction

Histology

The vas deferens, also known as the ductus deferens or spermatic duct, is a tubular conduit that begins at the tail of the epididymis and ends by joining the excretory duct of the seminal vesicle to form the ejaculatory duct. During the in situ inspection of a perinatal autopsy prior to testicular descent, the vas deferens is grossly recognized as a thin structure traveling from the tail of the epididymis over the ipsilateral umbilical artery toward the midline and terminating posterior to the urinary bladder at the base of the prostate. After testicular descent, the vas deferens can be seen emanating from the internal inguinal ring and terminating again at the prostatic base. The histology of the vas deferens in the fetus and neonate is not well studied, but some significant changes, especially in the structure of the muscular wall, are known to occur over the fetal period. This chapter reviews the development and histological changes in the vas deferens during gestation.

The fetal vas deferens is a tubular structure with an epithelium-­ lined lumen and a mesenchymal or muscular coat. In the adult vas deferens, a thin layer of connective tissue containing elastic fibers is seen beneath the epithelium, but these fibers are lacking in infants and children [1]. On hematoxylin and eosin (H&E)-stained sections of the fetal vas deferens, the epithelium appears to be in close contact with the muscular coat, with little to no intervening connective tissue. In the early midtrimester, the vas deferens is thin, with a round to ovoid lumen (Figs. 12.1 and 12.2). The epithelium is well developed and appears as a columnar to pseudostratified columnar epithelium. The apical border of the lining cells displays a prominent eosinophilic terminal bar, a marker of the stereocilia present on the surface of these cells. Unfortunately, the stereocilia may appear sloughed or are not

Embryology The vas deferens is derived from the mesonephric duct. As reviewed in the Part IV introduction, at approximately the seventh gestational week, two curves divide the mesonephric duct into three segments. The two most caudal segments, distal to the tail of the epididymis, become the vas deferens and terminate in the urogenital sinus.

L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected] E. D. Ruchelli · D. S. Huff Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]; [email protected]

Fig. 12.1  Vas deferens at 17 weeks gestation. Early in the midtrimester, the vas deferens is characterized by a central lumen surrounded by mesenchyme, with very little hint of smooth muscle differentiation. The lining of the vas deferens consists of a columnar epithelium with a well-­ defined terminal bar and cilia on the apical surface (H&E, 20×)

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Fig. 12.2  Vas deferens at 20 weeks gestation. The epithelial lining has not changed significantly, but the surrounding mesenchyme is slightly more organized, with increased cytoplasm noted in the spindle-shaped cells (H&E, 20×)

Fig. 12.3  Vas deferens at 24 weeks gestation. By the late midtrimester, the wall of the vas deferens appears more organized and shows clear smooth muscle differentiation. The epithelium appears as pseudostratified columnar epithelium with cilia (H&E, 20×)

Fig. 12.4  Vas deferens at 29 weeks gestation. This section is from the ampullary region of the vas deferens; the lumen is slightly larger and irregular. The epithelium is pseudostratified columnar, and cilia are not well preserved. The muscular coat is thick, and smooth muscle differentiation can be seen (H&E, 20×)

Fig. 12.5  Vas deferens at 34  weeks gestation. At this gestational age, the wall of the vas deferens exhibits well-developed smooth muscle, and the beginning of layers can be identified. The epithelium is pseudostratified columnar, with cilia noted on the apical surface. Basal cells can be seen along the basement membrane (H&E, 20×)

easily seen in sections from autopsy material, as preservation is typically suboptimal. The muscular coat of the vas deferens in this early stage of gestation is present; it appears as concentric layers of closely packed, spindle-shaped mesenchymal cells without significant smooth muscle differentiation (see Figs. 12.1 and 12.2). As gestation progresses into the late midtrimester to the early third trimester, the muscular coat develops further and thickens (Figs.  12.3 and 12.4). The vas deferens widens in diameter. The epithelium remains unchanged, but the mesenchyme of the muscle wall begins to take on a more eosinophilic appearance as smooth muscle differentiation progresses.

Near term, the muscular wall of the vas deferens becomes quite thick, and spindle-shaped cells show definitive smooth muscle differentiation. The formation of distinct layers of muscle also becomes apparent (Figs.  12.5 and 12.6), although some research has stated that evidence of separate layers of muscle is not apparent until after birth [2]. In the mature, adult vas deferens, there are inner and outer longitudinal muscle layers and a middle oblique or circular layer [1]. The appearance of the lining epithelium of the vas deferens remains constant throughout gestation as a columnar to pseudostratified columnar epithelium with

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Fig. 12.6  Vas deferens at 38 weeks gestation. This low-power image shows that the vas deferens is larger at term. Note the well-defined smooth musculature of the wall, now divided into distinct layers (H&E, 10×)

Fig. 12.7  Vas deferens at 38  weeks gestation. This higher-power image shows the luminal lining characterized by a pseudostratified columnar epithelium with poorly preserved cilia (H&E, 40×)

Fig. 12.8  Ampulla of the vas deferens at 34 weeks gestation. This low-­ power image shows the larger, wider lumen of the vas deferens in the ampullary region. Note the outpouchings of the lumen characteristic of the ampulla (H&E, 10×)

Fig. 12.9  Ampulla of the vas deferens at 34  weeks gestation. This higher-power image shows the widened and more complex ampullary lumen. Note that the epithelium is not distinctly different from the more proximal vas deferens. Basal cells and stereocilia can be appreciated (H&E, 20×)

stereocilia (Fig. 12.7). A layer of basal cells is described to be present in the mature vas deferens [1], but they are difficult to appreciate in routine fetal sections. Ultrastructural studies have identified at least four different cell types within the epithelium of the mature vas deferens: principal cells, pencil or peg cells, mitochondria-enriched cells, and basal cells [1]. The proximal portion of the vas deferens has a round to ovoid lumen (see Figs. 12.1, 12.2, and 12.3), but distally, in the area anatomically known as the ampulla, the lumen of the vas deferens widens. In sections of the distal vas deferens, this wider lumen can be appreciated with small

outpouchings of the lumen, imparting a somewhat stellate appearance to its shape (see Figs. 12.4, 12.8, and 12.9).

References 1. Trainer T.  Testis and excretory duct system. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 1003–26. 2. Dixon JS, Jen PY, Gosling JA.  Structure and autonomic innervation of the human vas deferens: a review. Microsc Res Tech. 1998;42:423–32.

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Seminal Vesicle Linda M. Ernst, Eduardo D. Ruchelli, and Dale S. Huff

Introduction

Embryology

There are three accessory glands of the male genital system: the seminal vesicles, the prostate, and the bulbourethral glands. The seminal vesicles are paired structures on the posterolateral aspect of the base of the bladder. Their main function is to secrete fluid that will form part of the semen; secretions of the seminal vesicles contain nutrients for the spermatozoa (such as fructose), as well as other proteins, enzymes, and mucus [1]. Because spermatozoa are not yet developed in fetal life, the physiological functions of the seminal vesicles are not necessary for the maintenance of life, but proper development of all the components of the male genital tract is important for future fertility. Aberrations of the proper development of the seminal vesicles, such as agenesis, fusion, duplication, hypoplasia, diverticulum, and cysts [2], can be seen during fetal life and can occur in association with other genitourinary tract anomalies, owing to the close embryological origin of these complex tissues. Therefore, knowledge of the normal development and histologic appearance of the seminal vesicle during fetal life can be useful. This chapter reviews the embryology of the seminal vesicles and highlights the histologic features of the fetal seminal vesicles at various points in gestation.

The three accessory glands of the male genital system—the seminal vesicles, the prostate, and the bulbourethral glands— originate at the most caudal end of the mesonephric duct, where it joins the endodermally derived pelvic urethra. The seminal vesicles begin to develop in the 10th–12th weeks as bilateral, dorsolateral outpouchings from the mesonephric duct proximal to the prostatic outpouchings, at the angle between the vertical and horizontal portions of the duct [3, 4]. By the 13th week, the outpouchings enlarge, and distal constrictions are visible. These represent the beginning of the developing ejaculatory ducts [4], which will receive both the ampulla of the distal vas deferens and the ipsilateral seminal vesicle. Further development of the seminal vesicles occurs in fetal life, as described below.

L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected] E. D. Ruchelli · D. S. Huff Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]; [email protected]

Histology General Overview The overall appearance of the seminal vesicle is that of a coiled, tubular structure. The wall is composed of an epithelial layer, lamina propria, and muscularis. The mucosa of the adult seminal vesicle is extensively folded and alveolus-like [5]. These mucosal convolutions do begin to develop in fetal life and appear fairly significant by term, but they do not reach the complexity seen in adulthood. The mature seminal vesicle epithelium consists of a layer of luminal columnar, secretory cells with microvilli and an underlying layer of basal cells. Adult columnar epithelial cells contain lipofuscin pigment [5], which is not a feature seen in fetal seminal vesicles. The lamina propria is the connective-tissue layer beneath the basement membrane of the epithelium; it contains connective-tissue elements and blood vessels. The outer coat of the mature seminal vesicle comprises a thin, external longitudinal smooth-muscle layer and a thicker, internal circular smooth-muscle layer [5].

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Second Trimester By 15 weeks gestation, the seminal vesicle anlage becomes “kinked” and consists of a medial, horizontal portion and a lateral, vertical portion [4]. In the early midtrimester, the lumen of the seminal vesicle shows only occasional diverticula, folds, or outpouchings; significant folding of the epithelium has not yet occurred (Figs.  13.1 and 13.2). The surrounding mesenchyme appears to be undifferentiated and

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consists of haphazardly arranged spindle-shaped cells (see Fig. 13.2). Later in the midtrimester, the seminal vesicle is more coiled, and the lumen develops more epithelial convolutions, which are especially noticeable in the distal half (Fig. 13.3) [4]. In addition, the mesenchyme surrounding the lumen matures, and early formation of the lamina propria and muscularis is seen (Fig. 13.4). The epithelium appears as a simple columnar type or a pseudostratified columnar type (Fig. 13.5).

Fig. 13.1  Seminal vesicle at 17 weeks gestation. The architecture of the seminal vesicles (SV) appears to be similar to the ampulla of the vas deferens (V). The lumen shows minimal outpouchings. The surrounding mesenchyme is immature, with little definitive smooth-muscle differentiation (hematoxylin and eosin (H&E), 4×)

Fig. 13.2  Seminal vesicle at 17 weeks gestation. This image shows a higher-power view of the seminal vesicle. The lumen shows minimal outpouchings, and the surrounding mesenchyme is immature, with little definitive smooth-muscle differentiation (H&E, 10×)

Fig. 13.3  Seminal vesicle at 24  weeks gestation. This low-power image shows the overall architecture of the seminal vesicles (SV) with increased coiling, especially distally (H&E, 1×). U urethra

Fig. 13.4  Seminal vesicle at 24  weeks gestation. This low-power image shows the overall architecture of the seminal vesicle, with increased coiling. The epithelium also shows increased outpouchings. The surrounding mesenchyme appears to show two indistinct layers: an inner, less-differentiated mesenchyme layer (lamina propria) and the outer layer, showing more smooth-muscle differentiation (muscularis) (H&E, 4×)

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Fig. 13.5  Seminal vesicle at 24  weeks gestation. This high-power image highlights the epithelium of the seminal vesicle, which appears columnar and pseudostratified (H&E, 40×)

Fig. 13.6  Seminal vesicle at 29  weeks gestation. This low-power image shows the overall coiled architecture of the seminal vesicles (SV), the increasing definition of the wall, and the relationship with the vas deferens (V) and urethra (U) (H&E, 1×)

Fig. 13.7  Seminal vesicle at 29 weeks gestation. This image highlights the development of the wall of the seminal vesicle. The lamina propria and muscularis are more easily appreciated, and the epithelium continues to show folds (H&E, 10×)

Fig. 13.8  Seminal vesicle at 29  weeks gestation. This higher-power view highlights the epithelium, which still appears partially pseudostratified. A hint of two populations of epithelial cells is seen, with a basal layer and a more superficial layer (H&E, 20×)

Third Trimester

still appear pseudostratified, but the formation of two distinct layers can be seen as some cells along the basement membrane take on the appearance of basal cells (Fig. 13.8). As the third trimester progresses toward term, the epithelial folding increases to involve the majority of the organ, with the most prominent folding seen at term (Figs. 13.9, 13.10, 13.11, and 13.12). The epithelial differentiation into basal and columnar cells also becomes more apparent late in gestation.

In the early third trimester, the coiling of the seminal vesicle and folding of the epithelium increases (Fig. 13.6). The surrounding muscularis thickens and takes on a more eosinophilic appearance as smooth-muscle differentiation also increases. The lamina propria is distinct beneath the epithelium, containing tightly packed spindle cells without eosinophilia in their cytoplasm (Fig. 13.7). The epithelium may

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Fig. 13.9  Seminal vesicle at 34  weeks gestation. This low-power image shows the coiled seminal vesicle with slightly more complex epithelial folding (H&E, 10×)

Fig. 13.10  Seminal vesicle at 34 weeks gestation. Epithelial folding is more complex, and the distinction between basal epithelial cells and apical cells is more obvious (H&E, 20×)

Fig. 13.11  Seminal vesicle at 38 weeks gestation. This low-power image shows the coiled nature of the seminal vesicle; by term, the most complex folding of the epithelium is appreciated. Note the eosinophilic appearance of the muscularis and the paler-appearing lamina propria (H&E, 4×)

Fig. 13.12  Seminal vesicle at 38 weeks gestation. In this higher-power image of the epithelium, note the paler basal cells and columnar luminal cells with secretions present in the lumen (H&E, 20×)

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References 1. Huggins C, Scott W, Heinen JJ.  Chemical composition of human semen and of the secretions of the prostate and seminal vesicles. Am J Phys. 1942;136:467–73. 2. Arora SS, Breiman RS, Webb EM, Westphalen AC, Yeh BM, Coakley FV. CT and MRI of congenital anomalies of the seminal vesicles. AJR Am J Roentgenol. 2007;189:130–5.

165 3. Schoenwolf GC, Larsen WJ. Larsen’s human embryology. 4th ed. Philadelphia: Churchill Livingstone/Elsevier; 2009. 4. Brewster SF. The development and differentiation of human seminal vesicles. J Anat. 1985;143:45–55. 5. Trainer T.  Testis and excretory duct system. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 1003–26.

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Prostate Gland Linda M. Ernst, Eduardo D. Ruchelli, and Dale S. Huff

Introduction There are three accessory glands of the male genital system: the seminal vesicles, the prostate gland, and the bulbourethral glands. The prostate gland is the largest of these three glands. Its main purpose is to store prostatic secretions and, when needed, to expulse those secretions, which form part of the ejaculate. In fetal life, the physiological functions of the prostate gland are not necessary for the maintenance of life, but proper development of all the components of the male genital tract is important for future fertility. The prostate gland has both epithelial and stromal components that can be recognized histologically, and the early development of these components is seen during fetal life. Aberrations of the proper development of the prostate gland can be seen in some syndromes, such as androgen insensitivity syndrome, prune belly syndrome, and posterior urethral valves [1]. Therefore, knowledge of the proper development and histological appearance of the prostate gland during fetal life can be useful. This chapter highlights the histological features of the fetal prostate gland at various points in gestation.

Embryology The three accessory glands of the male genital system—the seminal vesicles, the prostate gland, and the bulbourethral glands—originate at the most caudal end of the mesoneph-

L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected] E. D. Ruchelli · D. S. Huff Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]; [email protected]

ric duct, where it joins the endodermally derived pelvic urethra. Under the inductive influence of the surrounding mesenchyme, multiple evaginations of the urethral endoderm begin the formation of the prostate gland in the tenth week [2]. These epithelial evaginations elongate and bifurcate in a tightly regulated process known as branching morphogenesis [3], which produces the ducts and acini of the prostate gland. The factor or factors secreted by the mesenchyme that induce prostatic development are still the subject of scientific investigation, but the process of prostatic development and growth is known to be androgendependent [4]. Initially, there are at least five endodermal outgrowths consisting of solid cords of epithelial tissue [2]. The cords develop lumens in the 11th week and, by branching morphogenesis, form the ducts and acini. By the 15th week, secretory activity of the prostate gland begins. The prostatic fibromuscular stroma is derived from the mesenchyme surrounding the epithelial components, which differentiates into the smooth muscle and connective tissue of the prostate [2]. The paramesonephric ducts largely regress in the male, except for the cranial and caudal ends. The caudal end of the fused paramesonephric ducts frequently remains and is incorporated into the developing prostate, where it is known as the prostatic utricle. The definitive embryological origin of the prostatic utricle is a matter of scientific debate. Origin from the urogenital sinus alone or from the urogenital sinus in combination with the paramesonephric ducts has been considered [5, 6]. The prostatic utricle can appear as a solid cord of cells, but it frequently acquires a lumen and can be cystically dilated [7, 8]. The lining cells typically appear as a stratified squamous epithelium. The bilateral bulbourethral glands, also known as Cowper’s glands, further develop caudal to the prostatic origin as additional endodermal buds from the pelvic urethra. The surrounding mesenchyme gives rise to the connective tissue and smooth muscle elements of the glands.

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Histology

The ducts are canalized and have a double-­layered epithelium consisting of a cuboidal basal cell layer and a cuboidal to General Overview columnar luminal cell lining (Fig. 14.2). The prostatic urethra is the largest epithelium-lined lumen The prostate gland is situated anatomically at the distal end that can be seen in cross sections including the central portion of the bladder, surrounding the prostatic urethra. In fetuses it of the gland. Within the prostatic urethra, there is an elonis typically recognized as a small, firm, ovoid structure at the gated median elevation or ridge that protrudes into the lumen, base of the bladder. On cross section, the fetal prostate gland known as the verumontanum or seminal colliculus. The verhas a discoid to round shape and is surrounded by an ill-­defined umontanum produces the inverted-U shape of the prostatic capsule. Skeletal muscle can be seen at the anterior aspect of urethra seen on cross sections (Fig. 14.3). The verumontanum the gland and likely contributes to the capsule (Fig. 14.1) [1]. is an important area of the urethra, which receives the ejaculaThe prostate is composed of epithelial and mesenchymal/stro- tory ducts and can be seen in most fetal specimens (Figs. 14.3 mal elements. The stroma generally is more prominent in the and 14.4). Also seen at the verumontanum is the prostatic anteromedial portions of the gland, producing the convexity utricle, which typically appears as an ovoid, tubular structure of the gland anteriorly. Epithelial elements consist of ducts in with a stratified squamous epithelium, believed to be squathe central portion of the gland and acini more peripherally. As mous metaplasia related to estrogen-related hormone effect gestation progresses, the ducts and acini continue to develop (Fig. 14.5) [1]. Squamous metaplasia is also seen in the ureby increasing in number and volume within the prostate gland. thra, especially along the posterior wall (Fig. 14.6) [9]; it is also seen in ducts (Fig. 14.7). As the prostate gland matures, foci of squamous metaplasia become less prominent. Epithelial Elements As gestation progresses, the epithelial component of the prostate gland occupies a larger proportion of the gland, increasThe most centrally located, endodermally derived, epithelium-­ ing the gland-stroma ratio. In one histologic study of 107 fetal lined structures in the prostate gland are the ductal elements prostates at various gestational ages, 3 stages for the developof the prostate gland, which drain to the prostatic urethra. The ment of the ductal and acinar structures are described [9]: ducts consist of the branching epithelial structures that predominate in the central portion of the prostate gland around • The bud stage (at approximately 20–30 weeks gestation) the prostatic urethra. In fact, in the early midtrimester, ducts is characterized by simple, solid cellular buds at the ends compose most of the epithelial elements and are most promiof the ducts (Figs. 14.8, 14.9, 14.10, 14.11, 14.12, 14.13, nent in the posterior aspect of the prostate (see Fig. 14.1) [1]. and 14.14).

Fig. 14.1  Prostate gland at 17 weeks gestation. This low-power image of the fetal prostate gland shows a paucity of epithelial elements in the anterior portion (upper) and a few simple ductal structures posteriorly (lower). The prostatic urethra is present in the center of the gland. The stroma appears to be basophilic and shows minimal smooth muscle differentiation. An ill-defined connective tissue capsule is present, with a band of skeletal muscle noted anteriorly. The bilateral ejaculatory ducts can be seen in the lower aspect of the section (hematoxylin and eosin (H&E), 4×)

Fig. 14.2  Prostate gland at 17  weeks gestation. This higher-power image of the prostate gland highlights the ductal elements. They are mostly round to focally stellate in shape and are lined by a double-­ layered epithelium consisting of cuboidal basal cells and slightly more columnar apical cells. The surrounding mesenchymal stroma is composed of ovoid to spindle-shaped cells in a sheetlike arrangement with minimal smooth muscle differentiation (H&E, 20×)

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Fig. 14.3 Verumontanum at 17  weeks gestation. This low-power image of the prostate gland shows the verumontanum, which bulges into the prostatic urethra. The bilateral ejaculatory ducts (arrows) are visible within the verumontanum (H&E, 4×)

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Fig. 14.4  Ejaculatory ducts at 17 weeks gestation. This higher-power image shows the ejaculatory ducts with a pseudostratified columnar epithelium (H&E, 20×)

Fig. 14.6  Prostatic urethra with squamous metaplasia. This low-power image shows the prostatic urethra with its inverted-U shape. Note that it is lined by urothelium along the anterior (upper) wall and that there is squamous metaplasia of the epithelium along the posterior (lower) wall (H&E, 4×)

Fig. 14.5  Prostatic utricle at the verumontanum. This low-power image shows the inverted-U-shaped urethra and the protruding prostatic utricle (PU) at the verumontanum. The prostatic utricle is somewhat dilated and cystic and is frequently lined by squamous epithelium. Note the adjacent prostatic ducts coursing toward the urethra, and the ejaculatory ducts (arrows) located more posteriorly (H&E, 2×)

• In the bud-tubule stage (at 31–36 weeks gestation), cellular buds and acinar structures in small collections are identified (Figs. 14.15, 14.16, and 14.17). • The acinotubular stage (near term) is recognized by acinotubular clusters arranged in lobules. Acinar development is seen best at the periphery of the gland, whereas ductal elements are still seen in the central regions (Figs. 14.18, 14.19, and 14.20). The glandular elements at term maintain their double-­ layered epithelium characterized by basal cells along the base-

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Fig. 14.7  Squamous metaplasia of prostatic ducts. Squamous metaplasia of ducts is shown in this fetal prostate at 29 weeks gestation. This change becomes less frequent as gestation progresses (H&E, 20×)

Fig. 14.8  Prostate gland at 20 weeks gestation. At this gestational age, the prostate gland appears larger than at 17 weeks (see Fig. 14.1), and the central prostatic urethra is larger. The epithelial elements are still somewhat sparse and consist predominantly of ducts. The prostatic utricle is present posteriorly, between the ejaculatory ducts, and is lined by stratified squamous epithelium. The cross-sectional level is more proximal to the verumontanum, which is not shown here (H&E, 4×)

Fig. 14.9  Prostate gland at 20 weeks gestation. In this higher-power view, the prostatic ducts, which are slightly more complex than at 17 weeks, have similar double-layered epithelium (H&E, 20×)

Fig. 14.10  Prostate gland at 24  weeks gestation. At this gestational age, the gland is larger still, and this low-power image shows an increase in epithelial elements. Note the seminal vesicles located posteriorly (H&E, 2×)

ment membrane and secretory cells located apically. Basal cells have a cuboidal to flattened appearance by term and can be highlighted with immunohistochemical stains for highmolecular-weight cytokeratin and/or p63 (Fig. 14.21). Basal cells of the prostate gland do not contain muscle filaments, so they are not discernible with smooth muscle actin (SMA) immunohistochemistry, as seen with myoepithelial cells [10].

Stromal Elements The stromal elements of the prostate include the fibromuscular tissue, which intervenes between the epithelial elements

and the capsule. In the early midtrimester, the stroma of the fetal prostate gland has an immature appearance composed of spindle-shaped cells without much smooth muscle differentiation, and the capsule is an ill-defined connective tissue layer. Skeletal muscle bands are often seen in the anterior prostate (see Figs. 14.1 and 14.15). As gestation progresses, the stroma develops a more eosinophilic appearance as some of the stromal cells acquire the cytoplasmic properties of smooth muscle, with numerous spindle-shaped cells with a fibroblastic phenotype intermixed (see Figs.  14.11, 14.13, 14.16, and 14.20). Near term, the capsule is also thicker, with

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Fig. 14.11  Prostate gland at 24  weeks gestation. The epithelial elements at the periphery appear in the bud stage with solid buds at the ends of the ducts. The fibromuscular stroma shows more differentiation of the smooth muscle component (H&E, 10×)

Fig. 14.12  Prostate gland at 24  weeks gestation. This higher-power image shows the basal cells and more apical cells of the ductal lining epithelium. Note the small cellular buds emanating from the ends of the ducts (arrows), which is characteristic of the bud stage (H&E, 20×)

Fig. 14.13  Prostate gland at 29  weeks gestation. At this gestational age, still more epithelial development is appreciated. Note the ductular structures centrally and the beginning of acinar collections peripherally (H&E, 4×)

Fig. 14.14  Prostate gland at 29  weeks gestation. This higher-power image shows the epithelial elements near the periphery of the gland. The ducts terminate in buds that are just beginning to form acinar structures. Note the obvious smooth muscle differentiation in the stroma surrounding the epithelial elements (H&E, 40×)

a more well-defined connective tissue layer (see Figs. 14.15 and 14.18). Autonomic innervation of the prostate gland, which can be demonstrated using immunohistochemistry for protein gene product 9.5, is established during the fetal period. One study has shown that autonomic nerve fibers are first seen within the prostatic capsule at 13 weeks gestation, are

seen in the outer part of the gland by 17 weeks gestation in association with acini, and are seen throughout the gland by 26  weeks gestation [11]. Interestingly, the development of these autonomic fibers appears to follow the development of smooth muscle bundles within the stroma [11].

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Fig. 14.15  Prostate gland at 34 weeks gestation. Overall, the gland is larger still, and the epithelial elements are quite prominent, especially in the posterior aspect of the gland. The verumontanum is seen protruding into the prostatic urethra. Prostatic ducts drain into the urethra at this level. The connective tissue capsule is present, with a band of skeletal muscle incorporated anteriorly (H&E, 1×)

Fig. 14.16  Prostate gland at 34  weeks gestation. This higher-power image shows the epithelial components of the gland. The ducts now terminate in more well-formed small acinar structures (bud-tubule stage). Note the obvious smooth muscle differentiation in the stroma surrounding the epithelial elements (H&E, 10×)

Fig. 14.17  Prostate gland at 34  weeks gestation. This image is a higher-power view of the acinar structures at the terminus of the duct (H&E, 40×)

Fig. 14.18  Prostate gland at 39  weeks gestation. In this low-power image, the prostate gland is larger still. Note the prominence of the epithelial elements, involving the majority of the gland. The verumontanum is seen protruding into the prostatic urethra with numerous ductal structures entering the urethra. A connective tissue capsule is seen around most of the gland (H&E, 1×)

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Fig. 14.19  Prostate gland at 39  weeks gestation. This image shows ducts at the left and the formation of small lobules of acini (acinotubular stage) in the center. The acinar-lining cells show a clearing of apical cytoplasm with secretions. The elongated ducts present on the left side of the picture drain toward the urethra. The stroma has obvious smooth muscle differentiation (H&E, 4×)

Fig. 14.21  Prostate gland at 39 weeks gestation. PIN-4 cocktail immunohistochemistry, which includes high-molecular-weight cytokeratin and p63, is used to highlight the basal cells along the basement membrane of the acini. Basal cells form a continuous cuboidal to flattened layer at term (PIN-4 immunohistochemistry, 40×)

References 1. Popek EJ, Tyson RW, Miller GJ, Caldwell SA.  Prostate development in prune belly syndrome (PBS) and posterior urethral valves (PUV): etiology of PBS—lower urinary tract obstruction or primary mesenchymal defect? Pediatr Pathol. 1991;11:1–29.

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Fig. 14.20  Prostate gland at 39  weeks gestation. This higher-power image shows the secretions by the apical cells. Basal cells remain present along the basement membrane of the acini (H&E, 20×)

2. Schoenwolf GC, Larsen WJ. Larsen’s human embryology. 4th ed. Philadelphia: Churchill Livingstone/Elsevier; 2009. 3. Risbridger GP, Almahbobi GA, Taylor RA. Early prostate development and its association with late-life prostate disease. Cell Tissue Res. 2005;322:173–81. 4. Thomson AA, Cunha GR, Marker PC.  Prostate development and pathogenesis. Differentiation. 2008;76:559–64. 5. Shapiro E, Huang H, McFadden DE, Masch RJ, Ng E, Lepor H, Wu XR. The prostatic utricle is not a Müllerian duct remnant: immunohistochemical evidence for a distinct urogenital sinus origin. J Urol. 2004;172:1753–6; discussion 1756 6. Wernert N, Kern L, Heitz P, Bonkhoff H, Goebbels R, Seitz G, et al. Morphological and immunohistochemical investigations of the utriculus prostaticus from the fetal period up to adulthood. Prostate. 1990;17:19–30. 7. Glenister TW.  The development of the utricle and of the so-­ called “middle” or “median” lobe of the human prostate. J Anat. 1962;96:443–55. 8. Valdés-Dapena MA.  Histology of the fetus and newborn. Philadelphia: Saunders; 1979. 9. Xia T, Blackburn WR, Gardner WA Jr. Fetal prostate growth and development. Pediatr Pathol. 1990;10:527–37. 10. Fine SW, McKenney JK. Prostate. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 987–1002. 11. Jen PY, Dixon JS. Development of peptide-containing nerves in the human fetal prostate gland. J Anat. 1995;187:169–79.

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Ovary Eduardo D. Ruchelli and Dale S. Huff

Introduction The ovary is much smaller during fetal life than in adulthood (30 times lighter at birth than at puberty). Yet the number of germ cells is several times higher before birth than during the reproductive years (6–7 million at 20 weeks gestation, 0.5–1 million at birth, 300,000 at puberty, and less than 1000 at the end of menopause [1]). Thus, it is not surprising that the gross and microscopic features of the ovary continue to change throughout life, reflecting the steady decline of germ cells and the differences in folliculogenesis during fetal development, childhood, and after puberty. This chapter focuses on the histological features of the fetal ovary and highlights its differences from the adult ovary.

Embryology Ovarian differentiation of the bipotential gonad occurs at the end of the sixth week postfertilization, a day or two after testicular differentiation [2]. In the female, rather than forming testicular cords, the gonadal cords in the hilar area become indistinct. This area is called the central zone and will become the medulla of the ovary. The gonadal blastema surrounding the hilar area becomes thicker and is called the peripheral zone, which will become the ovarian cortex. The peripheral zone is composed of a monotonous sheet of cells, which include small, primordial germ cells and slightly smaller, oval interstitial cells, the precursors of granulosa and theca cells. The two cell types may be nearly indistinguishable except in very well-preserved specimens. The cell of origin of the interstitial cells is controversial; it may be the coelomic epithelial cells [3] or the mesonephric epithelial cells [4]. The ovary is E. D. Ruchelli (*) · D. S. Huff Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]; [email protected]

long, oval, has an irregular surface, and is distinct from the mesonephros at this age, but it is attached to the mesonephric body along its ventromedial border by a broad mesovarium. The ovarian tissue pushes into the mesonephros and abuts the mesonephric glomeruli; this may explain the occasional embryonic mesonephric glomeruloid remnants found in the ovarian hilus. Mesonephric mesenchyme begins to invade the central zone through the mesovarium. During the seventh week postfertilization, the peripheral zone becomes thick and develops very broad and indistinct cords. The enlargement of the peripheral zone is caused by rapid proliferation by mitosis of all cells, especially the primordial germ cells. In the central zone, remnants of the fragmented gonadal cords form broad epithelial cords (sex cords) that contain rare germ cells. The central cords extend as narrow strands of dark cells into the mesovarium; these narrow strands will become the rete ovarii. A study based on serial semi-thin sections of Epon-embedded ovaries suggests that central and peripheral cords arise from mesonephric glomerular and tubular epithelium and that coelomic epithelium does not contribute to the cords [4]. During the eighth week postfertilization, the peripheral and central zones become definite cortex and medulla, respectively. The ingrowth of stroma from the mesovarium into the medulla breaks up the medullary cords except for those of the rete. The deep margin of the thickening cortex becomes lobulated. The primordial germ cells in the deep cortex begin to differentiate into oogonia, and the differentiation into oogonia gradually spreads up from the deep cortex into more superficial cortex.

Histology The fetal ovary is tan, elongated, and flattened, in contrast to the adult ovary, which is pink-white, ovoid, and becomes increasingly convoluted during reproductive life. In near-­ term fetuses, follicular cysts may cause a lobulated external

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appearance. Although the neonatal ovary weighs only 0.3 g (compared to 5–8  g in the adult), it is almost as long as the fallopian tube. The ovary is attached at the hilus to the posterior aspect of the broad ligament by the mesovarium (Fig. 15.1). The peritoneum of the mesovarium is contiguous with the surface epithelium of the ovary at the hilus. From the hilus, the blood supply of the ovary penetrates through an ill-defined medulla into the cortex. The main cellular components of the ovary include the surface epithelium, the stroma and its derivatives, the germ cells, and the sex cord-derived granulosa cells, which will surround the germ cells and form the follicles. The surface epithelium consists of a single, focally pseudostratified layer of modified cuboidal peritoneal cells, which focally lacks a distinct basement membrane (Fig. 15.2). The tunica albuginea, a thin fibrous layer located under the surface epithelium, is not as distinct as in the testis (Fig. 15.3). In contrast to the adult ovary, the stroma of the fetal ovary is very scanty, as 90% of the cortex consists of crowded germ cells (Fig. 15.4).

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at 20  weeks gestation, when the largest number of germs cells can be observed, and then declines through week 28 [1]. Simultaneously, starting at 12–13  weeks gestation, some oogonia in the inner cortex enter meiosis and become oocytes (see Figs.  15.2 and 15.6). Oocytes are arrested in the very last stage of prophase for as long as 40  years or more; only a small fraction (about 400) will complete the first meiotic division at the time of ovulation. Between 14 and 18  weeks gestation, individual oocytes lose the

Oogonia Primordial germ cells begin to differentiate into oogonia during the early fetal period [5, 6]. Oogonia are premeiotic germ cells that represent the mitotically active pool from which meiotic oocytes develop and differentiate. The series of mitosis preceding meiosis results in a syncytium of clusters of oogonia, which are intermingled with groups of pregranulosa cells (Fig. 15.5). Oogonial mitotic activity peaks

Fig. 15.1  Ovary at 19  weeks gestation. The cortex (Co) comprises most of the ovary and overlies the medulla (Me). Crowded germ cells occupy 90% of the cortex. At the hilus (Hi), the ovary is attached to the mesovarium (Mo). The entire ovary is covered by a distinct, single layer of cuboidal surface epithelium, and the surface is irregular. The surface epithelium is contiguous with the peritoneal mesothelium of the mesovarium at the hilus (arrow) (hematoxylin and eosin (H&E), 2×). FT fallopian tube

Fig. 15.2  Surface epithelium: ovary at 21 weeks gestation. This closer view of the surface epithelium (right) demonstrates a single layer of cuboidal cells that focally lack a distinct basement membrane. The tunica albuginea is not developed at this age. The outer cortex is primarily occupied by a syncytium of oogonia (small arrows) and occasional primordial follicles (large arrows) (H&E, 20×)

Fig. 15.3  Tunica albuginea (TA): ovary at 28  weeks gestation. The tunica albuginea is better developed at this age than it was in Fig. 15.2. Also notice the larger number of primordial follicles with a layer of flattened granulosa cells and slightly increased spindle-shaped stromal cells (H&E, 20×)

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Fig. 15.4  Stroma: ovary at 19 weeks (a) and 35 weeks (b) gestation. Although spindle-shaped stromal cells remain a minor component of the ovary throughout fetal life (in contrast to the adult ovary), they become better developed during the third trimester (b). Notice the

marked increase in primordial and primary follicles (arrows). The edge of a secondary follicle, probably undergoing atresia, with a stratified layer of granulosa cells (Gr) and a rim of theca cells (Th), can be seen to the right in b (H&E, 20×)

Fig. 15.5  Oogonia: ovary at 16  weeks gestation. Mitotically active oogonia comprise the majority of cells in the cortex as the ovary approaches the stage with the largest number of germ cells throughout prenatal and postnatal life. The groups of oogonia represent a syncytium kept together by intercellular bridges between the germ cells. Pregranulosa cells and germ cells may be nearly indistinguishable except in very well-preserved specimens (H&E, 40×)

Fig. 15.6  Oocytes and follicles: ovary at 24 weeks gestation. Oocytes (arrows) are larger than oogonia and display a characteristic nucleus arrested in the last stage of prophase of the first meiotic division. In this field, they can be seen within primordial follicles (PrF) with a layer of flattened granulosa cells and within primary follicles (PriF) with a layer of cuboidal granulosa cells (H&E, 40×)

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intercellular bridges that kept them in a syncytial arrangement and become surrounded by a layer of flattened pregranulosa cells to form primordial follicles.

Folliculogenesis Follicles are the functional unit of the ovary. Each follicle consists of an oocyte surrounded by at least one layer, or up to several layers, of somatic granulosa cells (see Figs. 15.6 and 15.7). The follicle is separated from the surrounding stroma by a basement membrane, and the granulosa cells are devoid of any vascular supply. Folliculogenesis refers to the continuous process whereby primordial follicles undergo maturation. During reproductive life, only one follicle achieves complete maturation each month, culminating in ovulation. The other follicles that have begun simultaneous maturation undergo atresia at earlier stages of their development. Atresia accounts for the normal, steady decline in germ cells starting at 20 weeks gestation, when the loss of oogonia outpaces the mitotic activity of the remaining oogonia. Folliculogenesis and atresia occur prenatally and throughout childhood following the same stages that can be seen in the adult ovary, but the majority of these follicles will not reach the preovulatory stage. Several maturational stages can be observed prenatally and during childhood: • • • • •

The primordial follicle The primary follicle The secondary or preantral follicle The tertiary or antral follicle The mature or Graafian follicle (less common)

Fig. 15.7  Follicles: ovary at 28 weeks gestation. Initially, primordial and primary follicles are more numerous in the inner cortex. Because of poor preservation or handling, the surface epithelium, as shown here, is frequently denuded in autopsy specimens (H&E, 10×)

E. D. Ruchelli and D. S. Huff

Primordial follicles are dormant, quiescent follicles that represent the ovarian reserve. Histologically, they are characterized by a single oocyte or, less commonly, multiple oocytes surrounded by a single layer of flattened, mitotically inactive, granulosa cells resting on a basement membrane. The oocyte measures 40–70  μm in diameter (Fig.  15.8). Primordial follicles should be observed in the normal ovary by 18  weeks gestation [7]. However, autolysis frequently impairs the evaluation of folliculogenesis and ovarian dysgenesis in autopsy specimens, and caution should be used when tissues are not well preserved. Primary follicles represent the first morphological evidence of follicular maturation (Fig. 15.9). In contrast to their

Fig. 15.8  Primordial follicles: ovary at 28 weeks gestation. Primordial follicles consist of an oocyte 40–70 μm in diameter, surrounded by a single layer of flattened granulosa cells. The nucleus of the oocyte displays chromosomes arrested in the dictyate stage of prophase and one or more nucleoli. Rare oocytes may have multiple nuclei (H&E, 40×)

Fig. 15.9  Primary follicles: ovary at 30 weeks gestation. As the granulosa cells begin to become mitotically active, they assume a cuboidal shape, which characterizes the primary follicle (arrow) (H&E, 40×)

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flattened appearance in the primordial follicle, the granulosa cells assume a cuboidal to columnar shape and the oocyte enlarges. Secondary or preantral follicles are characterized by stratification of the granulosa cells into three to five concentric layers around the oocyte (Fig. 15.10). In addition, the oocyte becomes surrounded by an eosinophilic, homogeneous, acellular layer known as the zona pellucida. Secondary follicles measure from 50 to 400 μm in diameter. Simultaneously, the surrounding stromal cells begin to become specialized into several layers of theca interna cells and an ill-defined layer of theca externa cells (Fig. 15.11). Tertiary or antral follicles are seen when the secretion of fluid by granulosa cells results in the formation of fluid-filled clefts that eventually coalesce to form a single cavity lined by several layers of granulosa cells (Fig. 15.12). The oocyte continues to enlarge and may adopt an eccentric position at one pole of the follicle. When granulosa cells proliferate around the oocyte and form the so-called cumulus oophorus, the follicle is known as a mature or Graafian follicle (Fig.  15.13). In the nearterm fetus, multiple grossly visible Graafian follicles may be present.

Follicular Atresia Follicular atresia may begin at any stage of follicular maturation. Atresia of primordial and primary follicles is characterized by pyknosis and nuclear fragmentation of the oocyte and by cytoplasmic vacuolation. The granulosa cells degenerate,

Fig. 15.10  Secondary follicle: ovary at 32  weeks gestation. Mitotic activity in the granulosa cells results in the formation of multiple layers around the central oocyte. A Call-Exner body (arrow) can be seen within the layers of granulosa cells. The oocyte enlarges, and the stromal cells surrounding the follicle begin to differentiate into theca cells (H&E, 20×)

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and the follicle disappears without a trace [8]. In contrast, atresia of antral follicles usually results in the formation of a scar, the corpus fibrosum. Some follicles do not become completely obliterated and remain as atretic cystic follicles for an indefinite period (see Fig. 15.13). Follicles that exceed 3 cm are designated as follicular cysts [8]. Structures resembling microscopic gonadoblastomas and sex cord tumors with annular tubules have been identified within atretic follicles in up to 35% of normal fetuses and infants [8].

Fig. 15.11  Secondary follicle: ovary at 38 weeks gestation. This secondary follicle (left) is already surrounded by several layers of differentiating theca cells. The theca interna has three to four layers of plump cells, whereas the theca externa is ill-defined and merges with the adjacent ovarian stroma. Elsewhere in the field, there are numerous primordial and primary follicles (H&E, 20×)

Fig. 15.12  Tertiary or antral follicle: ovary at 38  weeks gestation. Granulosa cells begin to secrete fluid and create spaces that coalesce to form a single cavity. The tertiary follicle (TeF) is surrounded by better-­ differentiated theca cells (Th). Two Call-Exner bodies (arrow) can be seen within the layers of granulosa cells. A primary follicle can be seen to the left, whereas the outer cortex under the tunica albuginea (TA) contains mostly primordial follicles (PrF) (H&E, 20×)

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Hilus Cells

Rete Ovarii and Mesonephric Duct Remnants

Ovarian hilus cells are morphologically identical to testicular Leydig cells. Similarly to Leydig cells, they can be recognized during fetal life but become inconspicuous after birth, only to reappear at the time of puberty. Clusters of hilus cells are typically found in the hilus and mesovarium but can be found within the ovary at some distance from the hilus (Fig. 15.14). The cells are 15–25  μm in diameter and are round to oval, with abundant eosinophilic cytoplasm. Hilus cells should be distinguished from adrenal cortical rests, which are usually found in the mesovarium (see Chap. 20, Fig. 20.21).

The rete ovarii, the female analogue of the rete testis, is found at the hilus of the ovary. It consists of a network of clefts, tubules, and cysts with focal intraluminal papillae, lined by cuboidal to columnar epithelium. A cuff of spindle-­ shaped stromal cells surrounds the rete structures. The rete may juxtapose and communicate with mesonephric remnants within the mesovarium. Mesonephric remnants are commonly found in the vicinity of the ovary in fetal autopsies (Figs.  15.15 and 15.16). The remnants usually consist of tubules lined by cuboidal to columnar epithelium, sur-

Fig. 15.13  Graafian follicle: ovary at 32 weeks gestation. The oocyte has adopted an eccentric position and is surrounded by a mantle of granulosa cells, the cumulus oophorus (arrow). This tertiary follicle is undergoing atresia, as the granulosa cell layers at the periphery of the cystic follicle have thinned out and exfoliated (H&E, 1×)

Fig. 15.14  Hilus cells: ovary at 35 weeks gestation. Hilus cells (arrow) are morphologically identical to Leydig cells. Although typically seen at the hilus, they can be observed in clusters within the inner cortex, as in this case. The eosinophilic cytoplasm is very distinct in the fetus, but it becomes inconspicuous after birth, and the cells cannot be readily identified until they reappear after puberty (H&E, 20×)

Fig. 15.15  Mesonephric remnants: ovary at 19  weeks gestation. Blind-ending tubules in the mesovarium, corresponding to the efferent ductules and epididymis in the male, are known as epoophoron. This is a common finding in autopsy sections of the fetal ovary or fallopian tube (H&E, 4×)

Fig. 15.16  Mesonephric remnants: ovary at 19 weeks gestation. This closer view of the epoophoron demonstrates tubules lined by cuboidal to columnar epithelium, surrounded by a concentric layer of loose, spindle-shaped cells. Because of their common mesonephric origin, these structures are reminiscent of the fetal vas deferens in the male (H&E, 10×)

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rounded by concentric layers of spindle cells. Those found in the mesovarium next to the ovary are known as epoophoron; this consists of a few blind tubules and a duct that correspond to the efferent ductules and epididymis in the male. Closer to the uterus, some rudimentary tubules may persist as the paroophoron, and remnants corresponding to the vas deferens and ejaculatory duct in the male may persist as Gartner duct cysts between the layers of the broad ligament along the lateral wall of the uterus and in the lateral wall of the entire length of the vagina to the introitus [9].

References 1. Oktem O, Oktay K. The ovary: anatomy and function throughout human life. Ann N Y Acad Sci. 2008;1127:1–9. 2. Van Wagenen G, Simpson ME. Embryology of the ovary and testis: Homo sapiens and Macaca mullata. New Haven: Yale University Press; 1965.

181 3. Jirásek JE.  Development of the genital system and male pseudohermaphroditism. Baltimore: Johns Hopkins University Press; 1971. 4. Satoh M.  Histogenesis and organogenesis of the gonad in human embryos. J Anat. 1991;177:85–107. 5. Motta PM, Nottola SA, Makabe S.  Natural history of the female germ cell from its origin to full maturation through prenatal ovarian development. Eur J Obstet Gynecol Reprod Biol. 1997;75: 5–10. 6. Motta PM, Makabe S, Nottola SA.  The ultrastructure of human reproduction. I.  The natural history of the female germ cell: origin, migration and differentiation inside the developing ovary. Hum Reprod Updat. 1997;3:281–95. 7. McFadden DE, Pantzar T.  Genital system. In: Dimmick JE, Kalousek DK, editors. Developmental pathology of the embryo and fetus. Philadelphia: Lippincott; 1992. 8. Gilks CB, Clement PB.  Ovary. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 1119–48. 9. Moore KL, Persaud TVN, Torchia MG.  The developing human: clinically oriented embryology. 10th ed. Philadelphia: Saunders; 2015.

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Fallopian Tubes Eduardo D. Ruchelli and Dale S. Huff

Introduction The fallopian tubes and uterus have a common embryologic origin, so it is not surprising that they have a similar anatomic organization. During fetal life, the layers of the wall of the fallopian tubes mature and differentiate in a similar fashion to the uterus. In the second and third trimesters, the fallopian tubes typically exhibit a markedly convoluted gross appearance. Similar to the adult fallopian tube, the fetal fallopian tube can be divided into four segments: the intramural segment, within the wall of the uterine cornu; the adjacent isthmus, with a thick, stout wall and narrow lumen; the ampulla, which is thin-walled and tortuous; and the infundibulum, which opens into the peritoneal cavity by way of the fimbriated end.

Embryology The fallopian tubes, the uterus, and much of the vagina are derived from the paramesonephric (Müllerian) ducts, the development of which is described in the Part IV introduction. Each duct has two curves, which divide it into three segments: • The first segment, the most rostral, is vertical and extends from the abdominal ostium (the future fimbriated end) caudally along the posterolateral border of the ovary and mesonephros to the first curve [1]. This segment becomes the fallopian tube. The first curve, which bends medially under the caudal end of the ovary, is attached to the caudal end of the ovary by the ovarian ligament and by the round ligament to the inguinal canal. E. D. Ruchelli (*) · D. S. Huff Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]; [email protected]

• The second segment is horizontal and extends medially from the first curve to the midline, where it meets and fuses with the opposite duct and curves caudally (the second curve). This segment becomes incorporated into the ipsilateral lateral wall of the uterus. • The third segment is vertical; after fusing with the opposite duct, it extends caudally to its attachment to the posterior wall of the urogenital sinus at the level of the Müllerian tubercle. This segment becomes the uterus and vagina. The fallopian tube immediately develops a lumen that is continuous with that of the abdominal ostium. The epithelium of the abdominal ostium derives from the coelomic epithelium, whereas that of the remainder of the tube is of paramesonephric origin. The undifferentiated mesenchyme of the wall is derived from paramesonephric mesoderm and possibly partially from mesonephric mesoderm. The mucosa and muscularis remain undifferentiated until later in fetal life [1, 2].

Histology Three layers can be identified in sections of the mature fallopian tube: the mucosa, the muscularis (myosalpinx), and the serosa [3]. The tubal mucosa is invaginated into the lumen, forming longitudinal branching folds or plicae, which increase in complexity from the isthmus to the infundibulum (Fig. 16.1). During the early midtrimester, the mucosal stroma blends into the muscularis, and there is no clear distinction between the two (Figs. 16.2, 16.3, 16.4, and 16.5). The lumen is lined by undifferentiated simple or slightly pseudostratified cuboidal to columnar epithelium [4]. Cilia can be observed in well-preserved specimens, but not all cells are ciliated. Two other cell types have been described: secretory cells and intercalated cells. However, secretory cells are difficult to discern in the immature epithelium of the fetus,

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Fig. 16.1  Fallopian tube at 23  weeks. This slide includes two cross sections of the fallopian tube and the ovary (Ov) in the middle. On the left is the isthmus (Ist); on the right is the infundibulum (Inf). Notice how the complexity of the plicae increases and the thickness of the muscular coat decreases from the isthmus to the infundibulum (see Fig. 16.9) (hematoxylin and eosin (H&E), 1×)

Fig. 16.2  Fallopian tube at 19 weeks. Plicae are already developed, but the muscular layer is indistinct and blends with the stroma of the mucosa. Note the relatively thick connective tissue of the serosal layer (H&E, 10×)

Fig. 16.3  Fallopian tube at 19 weeks. This closer view shows a single layer of columnar cells with focally crowded nuclei along the mucosal surface. Cilia are barely discernible in some cells. The mucosal stroma blends into the developing muscular layer (H&E, 40×)

Fig. 16.4  Fallopian tube at 21 weeks. The demarcation between mucosal stroma and smooth muscle remains indistinct, as shown. Note the relatively thick connective tissue of the serosal layer (H&E, 10×)

and intercalated cells are modified secretory cells seen during the menstrual cycle after puberty [5, 6]. Toward the end of the midtrimester, the muscularis gradually becomes distinct (Fig. 16.6). The muscular layer is composed of an external longitudinal layer and an internal circular layer arranged

in a basket-weave fashion (Figs. 16.6, 16.7, and 16.8). The muscular coat is always thicker toward the uterine end of the tube (Fig.  16.9). The serosa comprises the peritoneum and underlying loose connective tissue. This layer is particularly thick during the midtrimester (see Figs. 16.2 and 16.4), but it also remains relatively prominent at term. The mucosal epithelium changes little during gestation (Fig. 16.10).

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Fig. 16.5  Fallopian tube at 21 weeks. This closer view shows no significant smooth-muscle differentiation. The epithelium consists of a single layer of columnar cells (H&E, 20×)

Fig. 16.6  Fallopian tube at 23 weeks. The myosalpinx becomes more distinct as the smooth-muscle fibers show better differentiation. The epithelium between plicae now almost rests on top of the muscular coat (H&E, 20×)

Fig. 16.7  Fallopian tube at 32 weeks. The mucosal stroma has more collagen, and the smooth-muscle fibers begin to be oriented in different directions (H&E, 10×)

Fig. 16.8  Fallopian tube at 38 weeks. This section closer to the ovarian end of the tube demonstrates increased complexity of the plicae and thinning of the muscular coat (H&E, 10×)

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b

Fig. 16.10  Fallopian tube at 38  weeks. As shown, the epithelium changes little during gestation, remaining a single layer of columnar cells (H&E, 20×) (Compare with Figs. 16.3 and 16.5)

References

Fig. 16.9  Fallopian tube at 23 weeks. This closer view is of the same segments of fallopian tube as depicted in Fig. 16.1. The muscularis is thicker in the isthmus (a) than in the infundibulum (b) (H&E; a 10×; b 4×)

1. Felix W. The development of the urinogenital organs. In: Keibel F, Mall FP, editors. Human embryology. Philadelphia: J.B. Lippincott Company; 1912. p. 752–979. 2. Hamilton WJ, Mossman HW.  Human embryology. Baltimore: Williams & Wilkins; 1972. 3. Valdes-Dapena MA.  Histology of the foetus and newborn. Philadelphia: Saunders; 1979. p. 469–77. 4. Jirásek JE. An atlas of human prenatal developmental mechanics. New York: Taylor & Francis; 2004. 5. Atkins KA, Hendrickson MR, Kempson RL. Normal histology of the uterus and fallopian tubes. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 1071–117. 6. Anderson MC, Robboy SJ, Russell P. The fallopian tube. In: Robboy SJ, Anderson MC, Russell P, editors. Pathology of the female reproductive tract. Philadelphia: Elsevier; 2002. p. 415–44.

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Uterus Eduardo D. Ruchelli and Dale S. Huff

Introduction The uterus of the fetus is relatively small, particularly when compared to the ovaries, which have about the same length as the fallopian tube. The relative size of the corpus and the cervix is the reverse of the adult uterus, the corpus being about one fifth to one third of the length of the cervix during gestation. This cervicocorpus disproportion is maintained into childhood. Most developmental anomalies of the uterus are the result of insufficient development of one or both Müllerian (paramesonephric) ducts or incomplete fusion, which results in different types of anomalous uterovaginal septation. Whenever the dome of the uterus is noticed not to be round or flat, serial cross sections of the corpus and cervix should be submitted for microscopic examination in order to evaluate the extent of septation of the lumen. The lumen of the fetal uterus is a slit-like potential space with a smooth lining, whereas the lining of the much larger cervix displays numerous delicate mucosal folds toward the end of gestation. The histological characteristics of the fetal uterine corpus and cervix are the focus of this chapter.

Embryology In the middle of the eighth postfertilization week, the paramesonephric ducts meet in the midline at an acute, “V”-shaped angle where the rostral wall of the uterine fundus will develop [1]. The ducts fuse and grow caudally to end in the posterior wall of the urogenital sinus at the site of the sinual (Müllerian) tubercle at the end of the eighth week [2]. This fused segment of the paramesonephE. D. Ruchelli (*) · D. S. Huff Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]; [email protected]

ric ducts is the uterovaginal canal, which will form the uterus and the rostral four fifths of the vagina [1]. Each of the two fused ducts immediately develops a lumen that is lined by undifferentiated, simple to slightly pseudostratified, cuboidal to low columnar epithelium derived from the paramesonephric mesodermal epithelium. The epithelium becomes thick, columnar, and stratified. The undifferentiated mesenchyme of the uterovaginal canal forms a wall that is distinctly thicker than that of the developing fallopian tube [3]. The mesenchyme is derived from the paramesonephric mesoderm and, to some extent, from mesonephric mesoderm of the mesonephric ducts that are embedded in the outer layers of the mesenchymal wall of the uterovaginal canal. The “V”-shaped point of fusion of the paramesonephric ducts is remodeled partly by retrograde fusion of the ducts in a rostral direction, so that the top of the uterine fundus becomes flat and then finally acquires its definitive domed shape. During about the 11th and 12th postmenstrual weeks, the common wall between the two paramesonephric ducts disappears, and the uterovaginal canal has a single lumen. In about the 13th and 14th postmenstrual weeks, the uterine corpus becomes anatomically distinct from the uterine cervix, and the anterior and posterior vaginal fornices appear and form a distinct demarcation between the uterine cervix and the vagina. Not until the 16th postmenstrual week does the vaginal epithelium become stratified squamous epithelium that is distinct from the uterine stratified columnar epithelium [3].

Histology The corpus and cervix change little histologically until the beginning of the third trimester [4]. Both the endometrium and endocervix are devoid of glands until the end of the midtrimester, when cervical glands start to develop (Figs. 17.1, 17.2, and 17.3).

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Fig. 17.1  Uterine corpus at 19 weeks: low magnification. The slit-like lumen of the uterus is lined by columnar epithelium, which is devoid of glands. The endometrial stroma is very cellular and blends into the myometrium with little demarcation. The serosa is thin and extends laterally into the broad ligament on each side of the uterus (arrows) (hematoxylin and eosin (H&E), 4×)

Fig. 17.2  Uterine corpus at 19  weeks: higher magnification. This higher magnification shows the smooth endometrial lining without glands. The epithelium retains the appearance of the early uterovaginal canal, with a stratified or pseudostratified columnar architecture. The basement membrane is usually prominent in the fetal and neonatal endometrium, and the stroma is poorly demarcated from the outer myometrium (H&E, 10×)

Fig. 17.3  Uterus at 19  weeks. This longitudinal section shows the internal os (Int OS). The endometrial (right) and endocervical (left) epithelial lining are very similar at this age, and both are devoid of glands (H&E, 10×)

Fig. 17.4  Uterine corpus at 21 weeks: low magnification. A few simple glands can be seen at this age. These early glands are rather simple, straight, and wide-mouthed; compare them to branching cervical glands at a similar gestational age as shown in Fig. 17.14. The thickness of the endometrium equals that of the myometrium, which is becoming slightly more eosinophilic; compare to Fig. 17.1 (H&E, 4×)

Uterine Corpus During the midtrimester, the surface epithelium of the endometrium retains the appearance of the early uterovaginal canal, with a stratified or pseudostratified columnar architecture. The underlying stroma is very cellular and composed of ovoid cells with scanty cytoplasm. The endometrial stroma is poorly demarcated from the myometrium (see Fig. 17.2). Late in the midtrimester, the first few glands begin to appear in the endocervix, but the appear-

ance of endometrial glands is very variable [5]; when present, they are short, straight, simple, and wide-mouthed (Figs.  17.4 and 17.5). The endometrial stroma and myometrium begin to become more distinct, but the thickness of the endometrium is about the same as that of the myometrium at this age (see Figs.  17.5 and 17.6). The endometrial glands remain simple in nature and are basically unchanged until late in the third trimester (Fig. 17.7). The smooth muscle cells gradually develop more eosinophilic

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Fig. 17.5  Uterine corpus at 21  weeks: higher magnification. This closer view of the early endometrial glands shows that they represent simple invaginations of the surface epithelium. The endometrial stroma in the inner half of the wall blends into the outer myometrium, which begins to have a predominantly concentric, smooth muscle fiber arrangement (H&E, 10×)

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Fig. 17.6  Endometrium at 21 weeks. This closer view shows the junction between the endometrial stroma and myometrium. The surface epithelium is stratified or pseudostratified columnar (H&E, 20×)

a

b

Fig. 17.7  Endometrium at 28 weeks (a) and 32 weeks (b). The endometrial glands remain simple and scanty (H&E, 10×)

Fig. 17.8  Uterine corpus at 35 weeks. The endometrial glands begin to branch and occupy the full depth of the mucosa. The endometrial stroma is well demarcated from the myometrium, and the myometrium has become thicker than the endometrium; compare with Fig. 17.4 (H&E, 2×)

cytoplasm and are clearly spindle-shaped, in contrast to the more rounded endometrial stromal cells, which have less cytoplasm (Figs.  17.8 and 17.9). The fascicular nature of the myometrium becomes more prominent during the third trimester, and the myometrium becomes thicker than the endometrium (see Figs.  17.8 and 17.10). Under the influence of maternal hormones, the endometrium of the nearterm fetus may develop rapidly and may have a transient proliferative appearance or, less commonly, a weak secretory appearance at birth (see Figs.  17.8, 17.9, 17.10, and 17.11). These changes regress during the first 2 weeks after birth (Figs. 17.12 and 17.13) [6].

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Fig. 17.9  Endometrium at 35 weeks. This closer view of the endometrium shows the glands with simple branches reaching the myometrium (left) (H&E, 10×)

Fig. 17.10  Uterine corpus at 38 weeks: low magnification. This cross section of the corpus shows a thick wall primarily occupied by the well-­ developed myometrium (H&E, 1×)

Fig. 17.11  Uterine corpus at 38  weeks: higher magnification. This closer view of the endometrium demonstrates glands with mild secretory activity, a feature that can be observed in term fetuses (H&E, 10×)

Fig. 17.12  Uterine corpus 18 days after birth: low magnification. This cross section of the corpus shows a thinner endometrium after postnatal withdrawal of maternal hormones. Compare with Figs. 17.10 and 17.11 (H&E, 1×)

Uterine Cervix

Fig. 17.13  Uterine corpus 18  days after birth: higher magnification. There are very rare, shallow glands and a single layer of low columnar epithelial cells on the surface. Compare with Figs.  17.10 and 17.11 (H&E, 20×)

During the early midtrimester, the endocervical lining is similar to the endometrium (see Fig. 17.3). Toward the end of the midtrimester, a few glands can be seen in the endocervical canal (Fig.  17.14), and the endocervical glands develop a more complex branching pattern earlier than the endometrium (Fig.  17.15). Mucus production becomes evident during the third trimester (Fig.  17.16) and continues to be progressively more abundant toward the end of gestation as the glands enlarge, become longer, and exhibit a more complex branching pattern (Figs.  17.17, 17.18, and 17.19). The vaginal portion of the cervix is known as the ectocervix or exocervix. It is covered by stratified squamous epithelium, which is identical to the mucosa of the vaginal fornices. Late in gestation, as the result of high maternal estrogen levels,

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Fig. 17.14  Uterine cervix at 23 weeks. A few glands can be seen in the endocervical canal, as shown. The glands begin branching relatively early, but otherwise the overall appearance of the endocervical wall at this age is similar to that of the corpus of the uterus. Compare with Fig. 17.4 (H&E, 4×)

Fig. 17.15  Uterine cervix at 24 weeks. The cervical surface and glandular epithelium is low columnar, as shown. Mucus production is not evident at this age. Note the more complex branching pattern of glands compared to the straight endometrial glands at the same gestational age (H&E, 20×)

Fig. 17.16  Uterine cervix at 32 weeks. At this age, mucus production by endocervical glands is scanty, as shown here (H&E, 10×)

Fig. 17.17  Uterine cervix at 32 weeks. A few glands can be seen in the endocervical canal (Endoc). The glands are typically slanted, almost parallel to the lumen, and pointing to the fundus. The lips of the external os (Ext Os) are completely covered by stratified squamous epithelium (H&E, 1×). Vag vagina

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Fig. 17.18  Uterine cervix at 35 weeks. Endocervical glands become more numerous and branching, particularly near the external os, as shown here. Mucus production increases, and the glands may become distended. The endocervical epithelium extends beyond the external os and covers the entire convexity of the portio vaginalis; compare with Fig. 17.17 (H&E, 2×)

E. D. Ruchelli and D. S. Huff

Fig. 17.19  Uterine cervix at 38 weeks. Mucus production is abundant at term, and the lumen of the endocervical glands becomes distended, as shown here (H&E, 1×)

fetuses at birth (see Fig. 17.18) [6]. After birth, the epithelial junction usually moves back into the endocervical canal and remains in that position until near menarche. In some fetuses, particularly near term, a large portion of ­ectocervix may be covered by columnar epithelium, sometimes including endocervical glands (Fig. 17.20). The replacement of the stratified squamous epithelium by columnar epithelium causes a grossly visible, sharply demarcated patch of brighter pink mucosa around the os. This normal variant has been erroneously called congenital erosion; it does not represent a true erosion, but rather the gross appearance of the thinner columnar epithelium.

References

Fig. 17.20  Uterine cervix at 37 weeks. In some neonates, all or part of the vaginal portion of the cervix is covered by columnar epithelium; this appearance is called congenital erosion. The squamocolumnar junction is at the vaginal fornix (arrow) (H&E, 1×)

the squamous epithelium matures and accumulates glycogen in its upper layers in a fashion similar to the vaginal mucosa. The external os is the anatomical opening of the endocervical canal into the ectocervix; the opening is usually slit-like, side-to-side, at birth. The squamocolumnar junction between the endocervical and ectocervical mucosa does not always lie at the external os. In fact, this junction shifts throughout life [7]. The junction tends to “move” outward in the latter part of pregnancy; it resides in the ectocervix in two thirds of

1. Jirásek JE. An atlas of human prenatal developmental mechanics. New York: Taylor & Francis; 2004. 2. O’Rahilly R, Muecke EC.  The timing and sequence of events in the development of the human urinary system during the embryonic period proper. Z Anat Entwicklungsgesch. 1972;138:99–109. 3. Yasuda Y. Genital system. In: Nishimura H, editor. Atlas of human prenatal histology. Tokyo: Igaku-Shoin; 1983. p. 147–69. 4. Valdes-Dapena MA.  Histology of the fetus and newborn. Philadelphia: Saunders; 1979. p. 469–77. 5. Felix W.  The development of the urinogenital organs. In: Keibel F, Mall FP, editors. Human embryology. Philadelphia: Lippincott; 1912. p. 752–979. 6. Atkins KA, Hendrickson MR, Kempson RL. Normal histology of the uterus and fallopian tubes. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 1071–117. 7. Wright TC, Ferenczy A. Anatomy and histology of the cervix. In: Kurman RJ, editor. Blaustein’s pathology of the female genital tract. New York: Springer; 2002. p. 207–24.

18

Vagina Eduardo D. Ruchelli and Dale S. Huff

Introduction During the first trimester and part of the second, the vagina undergoes complex embryologic changes that result in significant histological transformations. The histological appearance of the fetal vagina is also affected by placental and maternal hormonal influences that are unique to prenatal life. Those histological features are the focus of this chapter.

Embryology The developing vagina is derived from the fused paramesonephric ducts and the paired sinovaginal bulbs. At the end of the eighth postfertilization week, the fused paramesonephric ducts join the posterior wall of the urogenital sinus at the site of the sinual (Müllerian) tubercle [1]. The tubercle is a thickened plaque on the posterior wall of the urogenital sinus between the orifices of the mesonephric ducts. It is composed of epithelium derived from the urogenital sinus, mesonephric ducts, and paramesonephric ducts together with the underlying mesoderm. The fused paramesonephric ducts each have a lumen, except at their caudal ends, where they join the posterior wall of the urogenital sinus. At approximately the 11th or 12th postmenstrual week, two solid cords of epithelium from the urogenital sinus, the sinovaginal bulbs, arise at the posterolateral junctions of the mesonephric ducts with the sinual tubercle [2]. They grow rostrally and join the solid caudal ends of the paramesonephric ducts to form the solid vaginal plate. The rostral four fifths of the vagina derives from the paramesonephric ducts, and the caudal one fifth derives from the urogenital sinus through the sinovaginal bulbs [3]. At the

E. D. Ruchelli (*) · D. S. Huff Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]; [email protected]

same time, the common wall between the fused paramesonephric ducts disappears, and the single uterovaginal canal forms. The anterior and posterior vaginal fornices appear and establish the transition between the uterine cervix and the vagina. The short urethral portion of the vesicourethral canal, immediately rostral to the orifices of the mesonephric ducts and the sinual tubercle, begins to rapidly elongate caudally to form the female urethra and to displace the sinual tubercle more and more caudally. By the 20th postmenstrual week, the urethral orifice and the tubercle are at the introitus, and the tubercle differentiates into the hymenal ring [3].

Histology The vaginal wall consists of three layers: the mucosa, muscularis propria, and adventitia. Starting at 16 postmenstrual weeks, the pseudostratified columnar vaginal epithelium differentiates into stratified squamous epithelium [2]. After the squamous epithelium replaces the Müllerian columnar epithelium, it proliferates and progressively occludes the developing vagina (Figs. 18.1 and 18.2) [4]. Desquamation of the maturing epithelium leads to canalization of the vaginal plate (Fig. 18.3). By the 18th–20th week, the development of the vagina is complete, and the original Müllerian columnar epithelium of the uterovaginal canal is entirely replaced by stratified squamous epithelium from the urogenital sinus [5, 6]. Starting at 16 weeks gestation, the squamous epithelium begins to mature and accumulate glycogen, under the influence of placental estrogens (Figs. 18.4, 18.5, 18.6, 18.7, and 18.8). The abundant intracytoplasmic glycogen in the superficial layers of the fetal vaginal epithelium will disappear after birth, when estrogen levels drop. Without hormonal stimulation, the cells atrophy, so postnatally the vaginal epithelium remains in an atrophic state until puberty (Fig. 18.9). The squamous epithelium rests on the lamina propria. During the early midtrimester, the lam-

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Fig. 18.1  Vagina at 18 weeks gestation. This cross section illustrates how the lumen at this age is occluded by proliferating squamous epithelium, which in the previous weeks has replaced the original Müllerian columnar epithelium of the uterovaginal canal (hematoxylin and eosin (H&E), 4×)

E. D. Ruchelli and D. S. Huff

Fig. 18.2  Vagina at 18 weeks gestation. The lamina propria (LP) consists of cellular mesenchyme, and the muscularis propria (MP) is beginning to become distinct (H&E, 20×)

Fig. 18.3  Vagina at 23 weeks gestation. Maturation of the epithelium under estrogen effect is accompanied by extensive desquamation, which leads to canalization of the vaginal plate (H&E, 2×) (Compare Fig. 18.1)

ina propria blends into the muscularis propria, from which it is almost indistinguishable (see Fig. 18.2). As the mesenchyme of the lamina propria matures, it becomes less cellular and more collagenized (see Fig. 18.7). The muscularis propria of the vagina is continuous with that of the uterus, so it is not surprising that the smooth muscle fascicles in both organs become more distinct during the third trimester (see Figs. 18.6 and 18.7). The outer layers of muscle of both uterus and vagina run longitudinally, extending to the region of the hymenal ring [5], whereas the inner muscle layer has a spiral-like course (see Fig. 18.8). The adventitia is a thin layer of connective tissue that merges with the stroma of the adjacent structures.

Fig. 18.4  Vagina at 23 weeks gestation. The superficial layers of the squamous epithelium are rich in glycogen, and the cells show abundant clear cytoplasm surrounding small, pyknotic nuclei. Notice the cells desquamating into the lumen (top). The underlying cellular lamina propria (LP) is still poorly demarcated from the muscularis propria (MP) (H&E, 10×)

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Fig. 18.5  Vagina at 25  weeks gestation. The vaginal mucosa has an undulating appearance, in contrast to the flat surface of the cervix. Notice the urethra (top) displacing the muscularis propria along the anterior wall of the vagina (H&E, 2×)

Fig. 18.6 Vagina at 25  weeks gestation. The muscularis propria becomes more distinct, but an inner layer is not yet evident (H&E, 10×)

Fig. 18.7  Vagina at 32 weeks gestation. The epithelium thickens and accumulates abundant glycogen in the latter part of gestation. The lamina propria (LP) becomes less cellular, more collagenized, and more distinguishable from the muscularis propria, which now has an inner layer (IL) and an outer layer (OL) (H&E, 4×)

Fig. 18.8  Vagina at 32 weeks gestation. The outer layer (OL) of the muscularis is continuous with the longitudinal fibers of the uterus. The inner muscle layer (IL) has a spiral-like configuration (H&E, 10×)

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Fig. 18.9  Vagina at 18 days of postnatal life. After birth, estrogen levels drop, and the epithelium adopts an atrophic appearance, with cells devoid of intracytoplasmic glycogen (H&E, 4×; inset, 20×) (Compare the inset with Fig. 18.7)

References 1. O’Rahilly R, Muecke EC.  The timing and sequence of events in the development of the human urinary system during the embryonic period proper. Z Anat Entwicklungsgesch. 1972;138: 99–109. 2. Yasuda Y. Genital system. In: Nishimura H, editor. Atlas of human prenatal histology. Tokyo: Igaku-Shoin; 1983. p. 147–69.

3. Jirásek JE. An atlas of human prenatal developmental mechanics. New York: Taylor & Francis; 2004. 4. Forsberg JG.  Cervicovaginal epithelium: its origin and development. Am J Obstet Gynecol. 1973;115:1025–43. 5. Bean SM, Veras E, Bentley RC, Robboy SJ. Vagina. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 1059–69. 6. Bulmer J.  The development of the human vagina. J Anat. 1957;91:490–509.

19

External Genitalia Michael K. Fritsch

Introduction The external genitalia of fetuses are not commonly examined histologically. After birth, the external genitalia are examined grossly and usually designated as male, female, or ambiguous genitalia. Histologic examination of the external genitalia at the time of autopsy may be important in cases of cloacal abnormalities (more common in females) and in bladder outlet obstruction disorders, such as prune belly syndrome (more common in males). Therefore, understanding the histology of normal external male and female genitalia can be helpful in those rare cases when the external genitalia are abnormally formed. In addition, at some institutions, the male foreskin from routine circumcision surgery in a newborn is a common specimen for gross and/or histologic examination.

Embryology The external genitalia of both sexes are similar up until about the 9th week of development, and they are not fully differentiated until the 12th week [1–7]. External genitalia primordia first appear early in the fourth week with the formation of the genital tubercle from proliferating mesenchyme, appearing at the cranial portion of the cloacal membrane. The genital tubercle will elongate to form a primordial phallus, which eventually will become the penis or clitoris. Shortly after the formation of the genital tubercle, the labioscrotal swellings and urogenital folds appear. The labioscrotal folds are destined to give rise to the scrotum in males and the labia majora in females. The urogenital folds border the urethral groove and in females will develop into the labia minora.

M. K. Fritsch (*) Department of Pathology and Laboratory Medicine, University of Wisconsin, Madison, Madison, WI, USA e-mail: [email protected]

At 9  weeks of development, the male and female external genitalia begin to show distinct morphological maturation. Male external genitalia development from week 9 is initiated by testosterone production [1, 4–8]. In males, testosterone is produced from the fetal testes and converted to dihydrotestosterone (DHT) by the enzyme 5-alpha-reductase in the tissues destined to become the external genitalia. DHT acts locally to create the final masculinized morphology of the external genitalia. The primordial phallus will enlarge and elongate to form the penis. On the ventral surface of the developing penis, the urethral folds will form the lateral aspects of the urethral groove. Eventually the urethral folds fuse to form the spongy urethra along the ventral surface of the penis, and this is followed by surface ectoderm fusion in the median plane to form the penile raphe. At the tip of the glans penis, an ectodermal cord of cells forms, which will grow toward the spongy urethra. Once the cord becomes canalized, the penile urethra becomes complete, and the orifice is at the tip of the glans penis, rather than on the shaft. At about 12 weeks of development, a circular ingrowth of ectoderm begins to surround the glans penis and continues to grow to cover the glans by about 16  weeks. This extra layer of skin covering the glans penis is referred to as the prepuce (foreskin) and will remain fused to the glans throughout development and into early childhood, so that complete retraction of the foreskin is not normally possible until late childhood in most individuals. Also during the 12th week of development, the mesenchyme of the penis will give rise to the highly vascularized erectile tissue. The erectile tissues consist of the corpus spongiosum surrounding the penile urethra and extending into the glans penis and two laterally placed columns of erectile tissue, the corpora cavernosa. Also beginning by the ninth week of development in males, the labioscrotal swellings will grow toward each other and eventually fuse to form the scrotum. The site of fusion is visible on the skin surface as the scrotal raphe. The molecular events controlling external genitalia development are complex [9–11]. Testis descent into the scrotum is complex and proceeds slowly throughout

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intrauterine life, with the testes reaching the inguinal canal by about 6 months of development and then entering the canal by the seventh month and being completely descended into the scrotum by the end of the eighth month of development in most individuals. In some males, final testicular descent may occur in the first 3 months of extrauterine life [12]. In females, the absence of testosterone and the presence of estrogen result in the final morphology of the female external genitalia [2–7]. This is substantiated by the presence of female external genitalia or ambiguous genitalia seen in males with inactivating mutations in 5-alpha-reductase or the androgen receptor genes. The primordial phallus becomes the clitoris in females. The clitoris can be quite prominent up until 20  weeks of development, often necessitating careful inspection of the entire external genitalia to assign the correct sex. Mesenchymal-derived erectile tissue will develop within the clitoris. These crura of the clitoris are paired structures homologous to the corpora cavernosa in the penis. Similar to the penis, a small growth of ectoderm will surround the clitoral glans and form the prepuce of the clitoris, also referred to as the clitoral hood. This skin merges with the most superior fusion of the labia minora. The urogenital folds will give rise to the labia minora. The labia minora will form the lateral borders of the vaginal vestibule. The urethra and vaginal introitus open anteriorly within the vaginal vestibule. Most of the labioscrotal folds remain unfused and form the labia majora. The labioscrotal folds will fuse anteriorly to form the mons pubis and will fuse posteriorly to form the posterior labial commissure [13].

M. K. Fritsch

Fig. 19.1  Longitudinal low-power view of the male external genitalia (penis and scrotum) at 17 weeks of gestation. CC corpus cavernosum, CS corpus spongiosum, U urethra, G glans penis, S scrotum (hematoxylin and eosin (H&E), 2×)

Histology The Male External Genitalia The male external genitalia are composed of the penis and the scrotum (Fig. 19.1). The penis can be anatomically divided into the distal portion, composed of the glans, foreskin, and coronal sulcus, and the shaft, containing the erectile tissue and urethra. The arteries of the penis are branches from the internal pudendal artery (a branch of the iliac artery). These arteries branch extensively, and thick-walled helicine or perforating branches eventually open directly into the vascular spaces of the erectile tissue. The deep dorsal vein is the primary vein draining the erectile tissues. The deep dorsal vein eventually connects to the vertebral veins. The obstruction of venous outflow plays a key role in maintaining an erection. The penis contains a rich network of lymphatics, including within the foreskin and glans penis. The major nerves innervating the penis originate from the sacral and lumbar plexuses, with the dorsal nerve of the penis providing the primary somatosensory innervation to the penis, including the shaft, urethra, and glans [1, 8] (Fig. 19.2).

Fig. 19.2  Dorsal vein (DV) and large nerve (N) of the penis in a stillborn male at 22 weeks of gestation. Hemorrhage seen around vessels is common in stillbirths. CC corpus cavernosum, TA tunica albuginea surrounding both corpora cavernosa (H&E, 10×)

Distal Penis The glans penis is the bulbous distal end of the penis and is largely composed of an expanded corpus spongiosum covered by the foreskin (Fig. 19.3). The urethral meatus exits at the tip of the glans penis. The corpus spongiosum is the central erectile tissue surrounding the urethra in the shaft of the penis; the corpus spongiosum expands to fill most of the glans. The corpus spongiosum represents specialized venous sinuses lined by endothelial cells and contains abundant c­ onnective tissue, elastic fibers, and only sparse smooth muscle cells (Fig. 19.4). The proximal lateral bor-

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a

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Fig. 19.4  Erectile tissue of the glans penis at 18 weeks of gestation. On the left, the surface epithelium is separating from the glans, a common artifact in postmortem examinations (H&E, 20×)

Fig. 19.3  Longitudinal sections of the glans penis (G) at 14 weeks of gestation (a) and 18 weeks of gestation (b). CS corpus spongiosum, F foreskin (H&E, 4×)

der of the corona of the glans penis consists of an elevated rim that surrounds approximately 80% of the circumference, being interrupted on the ventral aspect by the mucosal fold of the frenulum (Fig. 19.5). The narrow circumferential recess just proximal to the glans corona is the coronal sulcus (Fig. 19.6). The surface of the glans after separation of the foreskin in early childhood is a partially keratinized, stratified squamous epithelium (5–6 cell layers thick), usually being more keratinized following circumcision. In the fetus, the glans is fused to the foreskin. The epithelium of the glans does not contain melanocytes, and neither adnexal structures nor glandular structures are present. The lamina propria is poorly defined fibrovascular tissue that blends into the corpus spongiosum. As a continuation of the skin of the penile shaft, the foreskin extends and eventually completely covers the glans by 15–18 weeks of gestation (see Fig. 19.3b). The outer surface of the foreskin is keratinizing epithelium, and

Fig. 19.5  Transverse section of the distal glans penis demonstrating the ventral frenulum (asterisk), the foreskin (F), and the navicular urethra (N, fossa navicularis) lined by squamous epithelium at 22 weeks of gestation (H&E, 2×)

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5  months of gestation [14]. The dermis contains developing adnexal structures, connective tissue, blood vessels, and nerves. The dartos fascia represents a discontinuous layer of smooth muscle fibers, connective tissue, and elastic fibers that extend from the penile shaft into the foreskin. The dartos fascia is found in all locations of the male external genitalia, including the foreskin, along the penile shaft, and in the scrotum (Fig. 19.8). The fourth layer of the foreskin is the lamina propria, composed of fibrovascular tissue and nerves. The fifth and innermost layer of the foreskin is the mucosal squamous epithelium, which has a variable thickness, no melanocytes, and no associated adnexal structures. This fifth layer is fused to the glans epidermis throughout fetal development. Occasional intraepithelial nerves may be seen [1, 8]. Fig. 19.6  Longitudinal section showing the coronal sulcus (C) of the distal penis at 19 weeks of gestation. CC corpus cavernosum, F foreskin, U urethra (H&E, 2×)

Fig. 19.7  Interface between the glans penis, containing the corpora spongiosum (CS), and the foreskin (F), with fused intervening squamous epithelium (asterisk) at 18 weeks of gestation. The dartos muscle fibers of the foreskin are thin, single, smooth muscle cells at this age and are difficult to see in this photomicrograph (H&E, 10×)

the inner surface is mucosal, with minimal keratinization after s­eparation from the glans. During fetal development and usually until mid-childhood, the glans, corona sulcus, and foreskin are fused by a shared, immature-appearing epithelial plate composed of a cuboidal to columnar basal layer on each side and a few layers of squamous epithelial cells in between. A normal foreskin is composed of five layers, including the outer surface epidermis, dermis, dartos layer, lamina propria, and inner squamous mucosa (Fig. 19.7). In newborns and thereafter, the outermost layer is keratinized, stratified squamous epithelium with scattered melanocytes. The melanocytes are difficult to see by conventional H&E staining in the fetal epithelium, and pigment does not begin to be expressed and transferred to keratinocytes until after

Penile Shaft The external surface of the penile shaft is composed of skin with a nonkeratinizing, stratified squamous epithelium until mid-gestation, when a thin layer of keratin develops that will persist thereafter. The basal epithelial layer is usually hyperpigmented in adults but not during fetal development, and there are scattered adnexal structures (Fig.  19.9). This layer of the skin of the penis directly extends to the foreskin. Below the epidermis and dermis lies the dartos fascia, similar to that seen in the foreskin; it is composed of discontinuous smooth muscle fibers arranged both transversely and longitudinally, with supporting connective tissue (see Fig. 19.8b). Below the dartos layer is the deep fascia of the penis (also known as Buck’s fascia), extending from the penile root to the coronal sulcus. Buck’s fascia is a continuous fibrovascular sheath encasing all three erectile tissues, the two corpora cavernosa and the corpus spongiosum. Buck’s fascia is not well demarcated histologically, especially in the fetal penis, where it is composed of loose connective tissue (including abundant elastic fibers to allow for expansion during an erection) and contains nerves, veins, and arteries (Fig.  19.10). The deep dorsal vein lies within Buck’s fascia and superficial to the tunica albuginea on the dorsal aspect of the penile shaft between the two corpora cavernosa (Fig.  19.11). The deep dorsal vein is not as prominent a structure as the superficial dorsal vein visible near the penile surface. The deep dorsal vein becomes partially compressed during an erection and helps maintain blood within the erectile tissues. Below Buck’s fascia but only encasing the two corpora cavernosa (each being completely surrounded) is the well-defined connective tissue layer referred to as the tunica albuginea, composed of collagen and rich in elastic fibers (Fig. 19.12). The ventral surface of the corpus spongiosum is not enveloped in the tunica albuginea. The corpora cavernosa are paired erectile tissue structures lying dorsal to the urethra and composed of thin, compressed, endothelial-lined vascular spaces with surrounding connective tissue and smooth muscle cells (Fig. 19.13). During an

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Fig. 19.9  Surface histology of penile shaft at 22  weeks of gestation. A adnexal structures, D dartos smooth muscle fibers, E epidermis (H&E, 10×)

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Fig. 19.10  Transverse section of mid-penile shaft at 22 weeks of gestation. B Buck’s fascia, CC corpora cavernosa, CS corpus spongiosum, TA tunica albuginea, U urethra (H&E, 2×)

Fig. 19.8  Dartos smooth muscle fibers at various locations and at different ages. (a) Intermediate-power view of foreskin (F) fused to the glans in a fetus at 18 weeks of gestation, with the individual smooth muscle fibers of the dartos layer visible (asterisk). CS corpus spongiosum. (b) A portion of the penile shaft surgically removed at 12 years of age, with the epidermal surface defined at the lower left, and discontinuous smooth muscle bundles of the well-developed dartos layer in the deep connective tissue (asterisk). (c) High-power view of scrotum at 18 weeks of gestation, with a few discontinuous smooth muscle cells deep in connective tissue (asterisk). The scrotal skin surface epithelium is to the left, and a portion of a primitive adnexal structure is present (H&E, a 10×; b 4×; c 20×)

Fig. 19.11  Longitudinal section of proximal penile shaft at 19 weeks of gestation. CC corpus cavernosum, CS corpus spongiosum, D deep dorsal vein, TA tunica albuginea, U urethra (H&E, 2×)

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Fig. 19.12  Transverse section of mid-penile shaft at 22 weeks of gestation. The tunica albuginea (TA) is a well-defined connective tissue layer that completely surrounds each corpus cavernosum (CC) but not the corpus spongiosum (CS). U urethra (H&E, 4×)

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erection, these vascular spaces become distended by blood. The corpus spongiosum is the other erectile tissue that directly surrounds the urethra on the ventral aspect of the penile shaft. The corpus spongiosum is composed of endothelial-­lined vascular spaces with a thin layer of surrounding smooth muscle cells; the spaces are separated from each other by connective tissue trabeculae (Fig. 19.14). In the early second trimester, the erectile tissue of the corpus spongiosum and corpora cavernosa appears hypercellular, and vascular spaces are difficult to appreciate by H&E staining. There are more elastic fibers and fewer smooth muscle cells in the corpus spongiosum than in the corpora cavernosa. The erectile tissues are fed by afferent arteries and drain into the deep dorsal vein via efferent veins. The proximal corpora cavernosa are connected by tapered crura that insert into the ischiopubic bone. The corpora cavernosa end bluntly at the distal end of the penile shaft (see Fig.  19.6). The corpus spongiosum extends into the glans penis and expands to occupy most of

a

a

b

b

Fig. 19.13  High-power view of cellular corpus cavernosum with mostly collapsed vascular channels at 13  weeks of gestation (a) and 22 weeks of gestation (b) (H&E, 20×)

Fig. 19.14  High-power view of corpus spongiosum at 13 weeks of gestation (a) and 22 weeks of gestation (b). The immature corpus spongiosum is cellular, with mostly collapsed vascular spaces. Note the increased connective tissue separating the vascular spaces of the more mature corpus spongiosum in (b), compared with the corpora cavernosa at the same age shown in Fig. 19.13b. The urethra (U) is also seen in (b) (H&E, 20×)

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the glans. Finally, the urethral epithelium varies in composition, being a nonkeratinizing, squamous epithelium for the distal portion of the penile urethra (referred to as the fossa navicularis), which is continuous with the epidermis of the glans penis (see Fig. 19.5). The majority of the penile urethra is referred to as the pendulous and bulbous urethra and is lined by surface columnar epithelium and a basal layer composed of stratified to pseudostratified columnar epithelium (Fig. 19.15). The distal penile urethra has numerous recesses referred to as Morgagni lacunae (see Fig. 19.15), which extend into small, single-layered, intraepithelial mucin-secreting periurethral glands (referred to as Littre glands). Littre glands

also line the lateral walls of the bulbous penile urethra. Littre glands contain clear mucinous cytoplasm. The two bulbourethral (Cowper) glands (Fig. 19.16) are located at the base of the penis and open into the proximal membranous urethra (a short segment of urethra just distal to the prostatic urethra).

Fig. 19.15  High-power view of penile urethra in mid-shaft at 22 weeks of gestation, showing multiple layers including a cuboidal to columnar surface epithelium and a lacuna of Morgagni with scattered intraepithelial mucin cells (glands of Littre). CS corpus spongiosum, M Morgagni lacuna, U urethra (H&E, 20×)

Fig. 19.16  Intermediate-power view of Cowper gland at 23 weeks of gestation with mild autolytic changes. Cowper glands are tubuloalveolar exocrine glands composed of mucinous acini (cells with clear cytoplasm) with intercalated ducts (cells with more dense cytoplasm) (H&E, 10×)

Fig. 19.17  The scrotum at 18  weeks of gestation. Low-power view demonstrating mostly loose, edematous fibrovascular tissue; in the median plane, the site of labioscrotal fusion is defined by slightly more dense collagen (asterisk) (H&E, 2×)

Fig. 19.18  The scrotum at 18 weeks of gestation. Intermediate-power view of scrotal skin with adnexal structures is shown. The dartos smooth muscle fibers cannot be seen in this photomicrograph (H&E, 10×)

Scrotum In early to mid-gestation, the fetal scrotum consists of nonkeratinized, stratified squamous epithelium overlying a loose fibrovascular dermis, scattered dartos smooth muscle fibers, and then loose connective tissue, often with edema and/or extravasated red blood cells in stillborn fetuses delivered vaginally (Figs. 19.17 and 19.18). By term, the skin surface

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will be a keratinizing, stratified squamous epithelium. The final layers of the scrotum are not established until the testes have completely descended, by the eighth to ninth month of development. The scrotum is seldom sampled at this late gestational age and usually only in cases with markedly abnormal development and therefore without the expected mature scrotal cellular layers. Following complete descent of the testes at term, the histologic layers of the scrotum consist of an external surface of skin composed of keratinizing, stratified squamous epithelium and underlying dermis with adnexal structures. Intimately within the dermis lies the tunica dartos, a continuation of the dartos fascia of the penile shaft and foreskin. The tunica dartos of the scrotum contains more smooth muscle fibers than the foreskin or penile shaft, but in fetuses this layer may not be easily seen at low power, depending upon the gestational age. Immediately below this layer lies a connective tissue layer referred to as the external spermatic fascia, which is derived from the superficial aponeurosis of the external oblique muscle, brought into the scrotum by testicular descent. Next is the cremaster muscle, which is composed of skeletal muscle fibers derived from the internal oblique and transverse skeletal muscles during testicular descent. Immediately subjacent to this is another connective tissue layer called the internal spermatic fascia, derived from the fascia of the transverse muscle. This is adherent to the parietal layer of the tunica vaginalis (derived from the processus vaginalis of the peritoneum) and lined by mesothelium. There is a virtual space and then mesothelium with an underlying thin layer of visceral tunica vaginalis immediately covering a thicker connective tissue layer, the tunica albuginea directly surrounding the testis [1, 15].

The Female External Genitalia The external genitalia of female fetuses are seldom studied histologically, although it is critical to be familiar with the gross anatomy in order to accurately assign anatomic sex and in order to identify pathologic conditions, including disorders of sex development. Most of the female external genitalia structures have homologous male counterparts based on embryonic origins, but the function may be significantly different in the female than in the male. The female external genitalia, referred to as the vulva, consists of the mons pubis, labia majora, labia minora, clitoris, and the vulvar vestibule, which contains the urethral meatus, the vaginal introitus, and the major and minor vestibular glands [2, 4, 13, 16, 17]. The most anterior portion of the vulva consists of the mons pubis, which lies superior to the vulvar vestibule and laterally divides into the labia majora. The two labia majora are derived from the labioscrotal folds, which do not fuse in females; their counterpart in males is the scrotum. Medially, the interlabial sulcus separates the labia majora from the

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labia minora (which lie medial to the labia majora). The labia minora form the lateral borders of the vulvar vestibule. The labia minora are derived from the urogenital folds and contain erectile tissue that is homologous to the male counterpart, the corpus spongiosum, but debate continues as to whether the labia minora erectile tissue has the same embryonic origins as the corpus spongiosum in males. In females, the amount of erectile tissue in the labia minora is much less than in the male. Anteriorly, the labia minora split, with the most anterior portion becoming continuous with the prepuce of the clitoris. Immediately below the clitoris, the labia minora can fuse to form the clitoral frenulum. Posteriorly, the labia minora fuse to form the posterior fourchette. This site of fusion may be poorly defined in some individuals. The clitoris is derived from the primordial phallus; the male counterpart includes the corpora cavernosa of the penile shaft and the glans penis. The vestibule is the portion of the vulva exterior to the hymen and extending to the frenulum of the clitoris anteriorly, to the fourchette posteriorly, to the labia minora anterolaterally, and to Hart’s line posterolaterally. Hart’s line represents the anatomic outline of the vulvar vestibule, including the medial aspects of the labia minora and the most inferior posterior portions of the labia minora and fourchette. The vulvar vestibule contains a number of structures, including the urethral opening and paired periurethral Skene ducts (the Skene gland being homologous to the male prostate). Also within the vestibule are the vaginal introitus and the associated hymen, with the major paired vestibular ducts draining the Bartholin glands (the male counterpart being the Cowper glands). The Bartholin gland ducts open beneath the hymen. Posterior to the fourchette lies the perineum [2, 13, 16, 17]. The arteries feeding the vulva include the superficial and deep pudendal arteries, derived from the femoral artery, and the internal pudendal artery, from the internal iliac artery. The deep artery of the clitoris and dorsal artery of the clitoris are branches of the internal pudendal artery. The venous drainage of the vulva is mostly via branches to the bilateral internal iliac veins. The vulvar lymphatics drain mostly to the inguinal lymph node chains, with some drainage to the femoral lymph nodes. The major nerves supplying the vulva include the anterior labial nerve (a branch of the ilioinguinal nerve) and the posterior labial nerve (a branch from the pudendal nerve). The dorsal nerve of the clitoris is a terminal branch of the pudendal nerve [2, 13, 16, 17].

Vulvar Vestibule The epithelium lining the vestibule is nonkeratinizing, stratified squamous epithelium and is derived from endoderm, rather than the ectodermally derived epidermis of the skin of the labia and mons pubis. The urethral orifice lies in the vestibule and contains transitional epithelium merging with stratified squamous epithelium at the opening (Figs.  19.19 and 19.20). The hymen represents the distal vagina and is

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Fig. 19.19  Low-power view of an oblique coronal section of the vulva at 21 weeks of gestation. C clitoris, Lm labia minora, P clitoral prepuce, U urethra, V vestibule (H&E, 2×)

Fig. 19.20  Oblique coronal section of the vulva at 21 weeks of gestation at slightly higher power. The urethral opening into the vulvar vestibule (transitional epithelium merging into squamous epithelium) can be seen. U urethra, V vestibule (H&E, 4×)

Fig. 19.21  Oblique coronal section of the vulva at 21 weeks of gestation, demonstrating the histology of the hymen. The vaginal introitus (Va) is separated from the more anterior vestibule (V) by the hymen (H), shown as a bridge of tissue with squamous epithelium on both sides and a fibrovascular core (H&E, 4×)

Fig. 19.22  Higher-power view of the oblique coronal section of the vulva at 21 weeks of gestation, demonstrating the histology of the hymen (H), especially the nonkeratinizing squamous epithelium and the fibrovascular core. V anterior vestibule, Va vaginal introitus (H&E, 10×)

the posterior or proximal aspect of the vestibule. The lining of the hymen is reminiscent of the vagina, a nonkeratinizing, stratified squamous epithelium, which can be glycogenated in term newborn females, owing to the effects of estrogen. The epithelium rests on highly vascularized connective tissue (Figs. 19.21 and 19.22). Scattered nerve endings can be found throughout this region. Also opening into the vestibule are the major and minor vestibular gland orifices. The paired periurethral Skene ducts open just below the urethral orifice. The Skene glands are composed of mucinous, columnar epithelium and drain via ducts lined with transitional epithelium. The major vestibular glands (Bartholin glands) are composed of acini lined by mucinous, columnar epithe-

lium, which merges with transitional epithelium with a surface layer of columnar cells lining the ducts; this becomes squamous epithelium at the ostia, which empty at the lower end of the vaginal introitus (Fig. 19.23). The minor vestibular glands consist of superficial, mucus-secreting columnar epithelium that directly connects to the mucosal surface of the vestibule. These minor vestibular glands are considered homologous to the male Littre glands. From personal experience, Littre glands are difficult to identify in fetuses in early and mid-gestation. Recently, small eccrine-like glands have been described in the sulcus between the clitoral prepuce and the clitoral glans [18] and have been detected in female fetuses after 15 weeks of gestation (Figs. 19.24 and 19.25).

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Fig. 19.23  Oblique coronal section of the female vulva at 21 weeks of gestation, demonstrating a Bartholin gland (BG) and its corresponding duct (D) situated below the lowest portion of the vaginal introitus (H&E, 4×)

Fig. 19.24  Low-power view demonstrating a small eccrine gland (EG) located just below the clitoral sulcus (CS) in a fetus at 21  weeks of gestation. C clitoris, P clitoral prepuce (H&E, 4×)

Fig. 19.25  Higher-power view of a small eccrine gland (EG) located just below the clitoral sulcus (CS) in a fetus at 21 weeks of gestation. N large nerves associated with the clitoris (H&E, 10×)

Fig. 19.26  A low-power view of an oblique coronal section of the clitoris (C), clitoral prepuce (P), and mons pubis (M) in a fetus at 21 weeks of gestation. Note the artifactual separation of the clitoris and prepuce. Ec erectile tissue of crura (homologous to corpora cavernosa), V vestibule (H&E, 2×)

Clitoris The clitoris is composed of two crura and a clitoral glans (Figs. 19.26, 19.27, and 19.28). The clitoral glans is covered by squamous mucosa without dermal appendages. Similar to the male glans penis, the clitoral prepuce is fused to the glans clitoridis during fetal development and at birth. The external surface of the clitoral prepuce is externally lined by stratified squamous epithelium. Below the clitoral epidermis, there is a very thin layer of connective tissue that merges with the

erectile tissue composed of small, slit-like vascular spaces lined by endothelial cells. Numerous nerves are found near the clitoris, which contains many nerve endings. The two crura, one on either side of the clitoral glans, are composed of erectile tissue homologous to the corpora cavernosa of the penis (Figs. 19.29, 19.30, and 19.31). A female homologue of the male corpus spongiosum does not exist within the clitoris, but erectile tissue is found in the two labia minora.

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Fig. 19.27  An intermediate-power view of the clitoris (C) and clitoral prepuce (P) in a fetus at 14 weeks of gestation. Note that the epidermis of the clitoris is fused to the prepuce (asterisk). Ec erectile tissue of crura (homologous to corpora cavernosa) (H&E, 10×)

Fig. 19.28  An intermediate-power view of the clitoris (C) fused to the prepuce (P) (asterisk) in a fetus at 21 weeks of gestation (H&E, 10×)

Fig. 19.29  The clitoral glans and crura. Low-power view of the clitoris (C) and the erectile tissue of both crura (Ec, homologous to corpora cavernosa) at 14 weeks of gestation (H&E, 2×)

Fig. 19.30  The clitoral glans and crura. High-power view of the female erectile tissue within the crura at 14 weeks of gestation (H&E, 20×)

Labia minora The paired labia minora lie on either side of the vestibule and are lined by stratified squamous epithelium, nonkeratinizing on the vestibular surface and thinly keratinizing on the lateral surfaces in late fetal life. In later life, this epithelium contains increased pigment, but this is not appreciated in fetuses, compared with the surrounding vestibular epithelium. Most of the labia minora are without adnexal structures (no hair

follicles, eccrine, or apocrine glands), but sebaceous glands are present on the lateral aspects (again, often sparse in midgestation fetuses). The epithelium rests on vascularized connective tissue with abundant elastic fibers and virtually no adipose (Fig. 19.32). This relatively highly vascularized tissue is considered erectile tissue similar to the corpus spongiosum, although the vascular density is less than that seen in the corpus spongiosum in males.

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Fig. 19.31  The clitoral glans and crura. High-power view of the female erectile tissue within the crura at 21 weeks of gestation (H&E, 20×)

Fig. 19.32  Labia minora (Lm) at 21 weeks of gestation. ILS interlabial sulcus, V vestibule (H&E, 4×)

Fig. 19.33  Labia majora (LM) and interlabial sulcus (ILS) at 21 weeks of gestation. C clitoris, Lm labia minora, M mons pubis, V vestibule (H&E, 2×)

Fig. 19.34  Mons pubis (M) at 21 weeks of gestation. C clitoris (H&E, 10×)

Labia Majora The labia majora lie lateral to the labia minora, separated by the interlabial sulcus (Fig.  19.33). In fetuses prior to the third trimester, the labia majora are lined by nonkeratinizing, stratified squamous epithelium, which will eventually keratinize and contain numerous adnexal structures, including hair follicles, sebaceous glands, apocrine glands, and eccrine glands. The medial aspect of the labia majora (the inner ­portion of the interlabial sulcus) does not contain hair follicles. Below the epithelium lies an elastic fiber-rich dermis. The labia majora are rich in nerve endings and sensory receptors. They merge with the mons pubis (Fig.  19.34), which has similar histology but contains a greater amount of deep adipose tissue, best appreciated after puberty.

References 1. Velazquez EF, Barreto JE, Cubilla AL. Penis and distal urethra. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2011. p. 1027–42. 2. Wilkinson EJ, Hardt NS. Vulva. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2011. p. 1045–58. 3. Fritsch MK, Cajaiba MM. The female reproductive system. In: Husain AN, Stocker JT, Dehner LP, editors. Stocker and Dehner’s pediatric pathology. 4th ed. Philadelphia: Wolters Kluwer; 2016. p. 869–70. 4. Moore KL, Persaud TVN, Torchia MG.  Urogenital system. In: The developing human clinically oriented embryology. 10th ed. Philadelphia: Elsevier; 2016. p. 267–70. 5. Marshall FF.  Embryology of the lower genitourinary tract. Urol Clin N Am. 1978;5:3–15. 6. Yamada G, Satoh Y, Baskin LS, Cunha GR.  Cellular and molecular mechanisms of development of the external genitalia. Differentiation. 2003;71:445–60.

19  External Genitalia 7. Sajjad Y.  Development of the genital ducts and external genitalia in the early human embryo. J Obstet Gynaecol Res. 2010;36: 929–37. 8. Yiee JH, Baskin LS. Penile embryology and anatomy. Sci World J. 2010;10:1174–9. 9. Yamada G, Suzuki K, Haraguchi R, Miyagawa S, Satoh Y, Kamimura M, et al. Molecular genetic cascades for external genitalia formation: an emerging organogenesis program. Dev Dyn. 2016;235:1738–52. 10. Cohn MJ.  Development of the external genitalia: conserved and divergent mechanisms of appendage patterning. Dev Dyn. 2011;240:1108–15. 11. Blaschko SD, Cunha GR, Baskin LS.  Molecular mechanisms of external genitalia development. Differentiation. 2012;84:261–8. 12. Bay K, Main KM, Toppari J, Skakkebaek NE. Testicular descent: INSL3, testosterone, genes and the intrauterine milieu. Nat Rev Urol. 2011;8:187–96.

209 13. Sacher BC. The normal vulva, vulvar examination, and evaluation tools. Clin Obstet Gynecol. 2015;58:442–52. 14. Holbrook KA, Vogel AM, Underwood RA, Foster CA. Melanocytes in human embryonic and fetal skin: a review and new findings. Pigment Cell Res. 1988;1(suppl 1):6–17. 15. Chaux A, Cubilla A, Hakim S, Pernick N.  Penis and scrotum. PathologyOutlines.com. Revised 22 May 2018. 16. Roychowdhury M.  Vulva: general, normal, anatomy/histology. PathologyOutlines.com. Revised 3 Oct 2017. 17. Yeung J, Pauls RN. Anatomy of the vulva and the female sexual response. Obstet Gynecol Clin N Am. 2016;43:27–44. 18. van der Putte SC, Sie-Go DM.  Development and structure of the glandopreputial sulcus of the human clitoris with a special reference to glandopreputial glands. Anat Rec (Hoboken). 2011;294:156–64.

Part V Endocrine System

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Adrenal Gland Linda M. Ernst and Lily Marsden

Introduction Structurally, the fetal adrenal gland is significantly different from the adult adrenal gland. One of the obvious differences noted on fetal dissection is that the fetal adrenal gland is proportionately large, appearing nearly as large as the kidney in the early midtrimester. In adults, the normal adrenal gland weight is about 5% of the normal kidney weight, and the adrenals combined represent 0.02% of total adult body weight, but throughout fetal life, the adrenal/kidney weight ratio and adrenal/body weight ratio are much higher (Table 20.1) [1, 2]. The bulk of the fetal adrenal gland is largely due to the presence of the fetal adrenal cortex, or fetal zone, which produces dehydroepiandrosterone (DHEA) and DHEA sulfate. These weak androgens are important precursors of placental estrogens. Later in gestation, the fetal adrenal gland will also produce aldosterone and cortisol. Therefore, the fetal adrenal gland is an organ of immense importance for the maintenance of pregnancy and fetal homeostasis. The hormone production also promotes organ maturation late in gestation and may assist in the timing of labor [3]. After birth, the fetal zone of the adrenal gland involutes, and the adrenal weight is markedly reduced.

Embryology The adrenal glands develop as bilateral structures within the intermediate mesoderm in intimate association with the dorsal root mesentery medially and the mesonephros and gonad laterally [4, 5]. The first cells to populate that mesoL. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected] L. Marsden Utah State Office of the Medical Examiner, Salt Lake City, UT, USA e-mail: [email protected]

Table 20.1 Adrenal weight and proportions of kidney and body weight Gestational age, week 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40

Adrenal weight, combined (g) 0.1 0.15 0.22 0.33 0.48 0.65 0.83 1.03 1.23 1.43 1.64 1.83 2.02 2.19 2.35 2.5 2.7 3.0 3.3 3.7 4.1 4.6 5.1 5.6 6.1 6.6 7.1 7.4 7.7

Kidney weight (%) 62.5 71.4 62.9 54.1 49.5 48.9 47.4 45.2 42.6 41.3 39.0 36.5 35.7 33.6 313 29.1 27.8 27.5 26.8 27.0 27.0 27.3 27.6 27.9 28.1 28.3 28.6 28.4 28.2

Body weight (%) 5 5 4 4 4 4 4 4 4 4 4 4 3 3 3 3 3 3 3 3 3 3 3 3 3 3 3 3 3

Data from Hansen et al. [1] with permission from Springer and Siebert [2] with permission from Elsevier

dermal area in approximately the fifth week of gestation are derived from the overlying mesothelial/celomic lining. These mesothelial cells proliferate and penetrate into the underlying tissue, forming the first cells of the adrenal gland, the fetal cortical cells. As the cells differentiate, they become large, polygonal cells with abundant eosinophilic cytoplasm.

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Within a short period, a second layer of cells, which also originate from the mesothelial lining cells, surrounds the developing fetal cortex. These cells will form the definitive cortex; as they differentiate, they will be generally smaller and less eosinophilic than the fetal cortical cells. The position of the developing adrenal glands relative to adjacent structures shifts caudally, whereas that of the kidneys shifts cranially, and the two meet to take their final positions. The cells of the adrenal medulla are derived from neural crest cells. Initially, the neuroblastic cells, derived from the neural crest, form collections along the aorta that will become the paravertebral sympathetic ganglia. Nerve fibers from the sympathetic ganglia penetrate the developing adrenal gland, and neuroblastic cells migrate along these tracts. The final destination of these neural crest derivatives is the central portion of the gland, but migrating neuroblastic-type cells can be seen singly or in small groups within the fetal cortex throughout fetal life and even into the neonatal period [6, 7].

Histology

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Fig. 20.1  Adrenal gland at 39 weeks of gestation. In this low-power view, note the thin capsule. The definitive zone of the cortex is the most superficial layer, with the more closely packed, darkly stained cells. The much thicker fetal zone of the cortex has the more eosinophilic appearance. In the center, large, thin-walled vessels are noted, surrounded by medullary cells (H&E, 4×)

General Overview The fetal adrenal gland is an encapsulated organ composed of two main areas—the outer cortex and the central medulla— similar to the adult organ. However, the cortex of the fetal adrenal gland is much thicker than the adult gland and consists of at least two histologically distinct layers: the definitive zone and the fetal zone (Fig. 20.1). It is the fetal zone, recognized by its large eosinophilic cells, that contributes to the large size of the fetal adrenal gland; it will undergo involution after birth.

The Capsule The capsule is thin and not very prominent in the second trimester, consisting only of thin layers of collagen and elastic tissue. By the late third trimester, the capsule is only slightly thicker.

The Cortex The adrenal cortex is composed of two histologically distinct layers, seen at different stages of fetal life in Figs. 20.2, 20.3, 20.4, 20.5, and 20.6. The most superficial layer of the cortex, located directly beneath the capsule, is known as the definitive zone, adult zone, or neocortex. It is recognizable by its basophilic appearance on hematoxylin and eosin (H&E)-stained sections. This definitive zone is thin (10–20 cells thick) and in the fetal adrenal represents only a small

Fig. 20.2  Adrenal gland at 12 weeks of gestation. Note the thin capsule. The definitive zone of the cortex is the most superficial layer, with the more closely packed, darkly stained cells. The deeper layer with the more eosinophilic appearance is fetal zone. Small clusters of migrating neuroblasts on their path to the medulla are seen within the fetal cortex (arrows) (H&E, 10×)

fraction of the entire cortical layer. The layer is composed of nonoverlapping, polygonal cells with round nuclei and scant eosinophilic to pale, sometimes vacuolated, cytoplasm (Figs.  20.7, 20.8, and 20.9). The tightly packed cells with scant cytoplasm give an overall basophilic appearance to the definitive cortex. During fetal life, the stratification that is present in adult adrenal cortex, into zona glomerulosa, fasciculata, and reticularis, is not visible, though near term, the

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Fig. 20.3  Adrenal gland at 17 weeks of gestation. Overall, the organization and appearance of the definitive zone of the cortex (the most superficial layer) and the deeper fetal zone have not changed since 12 weeks (H&E, 10×)

Fig. 20.4  Adrenal gland at 20 weeks of gestation. Low-power view of the superficial definitive zone of the cortex and the deeper fetal zone (H&E, 10×)

Fig. 20.5  Adrenal gland at 26  weeks of gestation. Adrenal cortex showing the distinction between the superficial definitive zone and the deeper fetal zone. Note the nature of the cortical cells arranged in cords with vascular sinusoids between them (H&E, 10×)

Fig. 20.6  Adrenal gland at 35 weeks of gestation. Low-power view of the adrenal cortex showing the distinction between the superficial definitive zone and the deeper fetal zone. Note the nature of the cortical cells arranged in cords with vascular sinusoids between them (H&E, 10×)

beginnings of this zonation may be seen, as the zona glomerulosa becomes organized into small clusters distinct from the cords of the zona fasciculata (Fig. 20.10). All three layers of the definitive cortex are well-developed within the first few months of life (Fig. 20.11). The fetal zone, also known as provisional cortex or transitional cortex, constitutes the remainder and majority of the adrenal cortex in fetal life. The fetal zone of the adrenal gland is not present in most mammals, but is seen in humans and a few nonhuman primates [3, 8, 9]. It is distinguished histologically from the definitive zone by its large size and eosinophilic appearance (see Figs.  20.1, 20.3, and 20.10).

The epithelial cells of the fetal zone are large, polygonal cells with round nuclei and abundant eosinophilic, sometimes vacuolated, cytoplasm (Fig. 20.12). These cells are generally arranged in columns separated by sinusoidal vascular spaces, although sometimes the fetal cortical cells appear so tightly packed that the sinusoidal spaces are difficult to appreciate. CD31 immunohistochemistry can be used to highlight the sinusoidal spaces (Fig. 20.13). The fetal zone represents the majority of the adrenal gland in fetal life but begins to involute rapidly after birth. Involution of the fetal cortex involves loss of the eosinophilic fetal cortical cells and is typically associated with some hemorrhage within the fetal cortex

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Fig. 20.7  Adrenal gland at 12 weeks of gestation. The capsule is composed of a few layers of collagen and elastic tissue. Note that there is no distinction between separate layers of the definitive cortex. Cells of the definitive zone are small, with hyperchromatic, round nuclei, arranged in cords (H&E, 40×)

Fig. 20.8  Adrenal gland at 20 weeks of gestation, showing the junction between the definitive and fetal zones. Although there is no distinction of layers in the definitive cortex, a small pseudoglandular space is present (H&E, 20×)

Fig. 20.9  Adrenal gland definitive zone at 35 weeks of gestation. The definitive zone is still arranged in cords, but now newly forming small clusters are suggested superficially, the beginnings of the zona glomerulosa (H&E, 20×)

Fig. 20.10  Adrenal gland cortex at 39 weeks of gestation. In this low-­ power view, note the early organization of the definitive zone with clusters of cells in the zona glomerulosa. The lower two thirds of the image is occupied by the eosinophilic-appearing fetal zone (H&E, 10×)

(Figs.  20.14 and 20.15). Loss of this layer of the adrenal gland leads to a greater than 50% decrease in the gland size during the first months of neonatal life [3]. The involutional process and reorganization are generally completed within 4–6 weeks after birth [7]. A third zone, known as the transitional zone, has been described using immunohistochemical studies [10]. This zone lies between the definitive zone and the fetal zone. Its role is to produce cortisol late in gestation, and it will become the zona fasciculata in the infant [3, 10]. This zone appears to have a similar thickness (10–20 cells) as the definitive zone, and it contains tightly packed cells

that are only slightly larger than those in the definitive zone. These features may make the transitional zone hard to recognize as a distinct layer on routine H&E-stained sections.

The Medulla The fetal adrenal medulla is not the well-formed chromaffin cell mass that is seen in the center of the adult adrenal gland. As reviewed in the embryology section, the cells of the adrenal medulla are neural crest derived, and in fetal

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Fig. 20.11  Adrenal gland cortex in a 17-month-old child. Note the well-developed organization of the definitive cortex, similar to an adult adrenal gland. The fetal cortex is no longer present (H&E, 10×)

Fig. 20.12  Adrenal gland at 12 weeks of gestation. Higher-power view of the fetal zone. Larger, polygonal cells with abundant eosinophilic cytoplasm and focal fine vacuolation make up the fetal cortex. Nuclei are uniformly round (H&E, 40×)

Fig. 20.13  Fetal adrenal gland stained with CD31, which highlights the endothelial cells lining the sinusoidal vascular spaces between the cortical cells (CD31 immunohistochemistry, 10×)

Fig. 20.14  Involution of fetal cortex/zone. Adrenal gland from a preterm infant who lived for 6 weeks postnatally. The fetal zone (outlined by arrows) is markedly reduced in thickness and cellularity. Congestion and hemorrhage are also present (H&E, 10×)

life these neuroblastic cells migrate into the adrenal gland from the cortex inward. Their route of migration can be seen in sections of the fetal adrenal gland as single neuroblastic cells or small groups within the cortex. Neuroblastic nodules are unencapsulated groups of densely packed cells with high nuclear/cytoplasmic ratios, scanty cytoplasm, and nuclei with dense, granular chromatin. Therefore, they are easily differentiated from the large eosinophilic cells of the fetal zone of the cortex. Turkel and Itabashi [7] provide a detailed histological study of the size, location, and histological appearance of neuroblastic nodules in fetal adrenal glands from 10 to 30 weeks of gestation. Neuroblastic cells are seen in both superficial/subcapsular and deep locations. Superficial groups are seen at the youngest gestational ages

and decline with increasing gestational age. Superficial neuroblasts are rare after 20 weeks of gestation, but deep groups are seen throughout gestation (Figs.  20.16 and 20.17), including the presence of even a few nodules at the time of birth [11]. Later in gestation, the neuroblastic nodules are present in the center of the gland, surrounding central veins of the medulla (Fig. 20.18). The size of the neuroblastic nodules also varies over gestational age, with the size of nodules increasing until the peak size is attained between 17 and 20 weeks. The size of the neuroblastic nodules decreases thereafter, with the smallest nodules noted late in gestation. Differentiation of the neuroblastic cells to chromaffin cells is likely the reason for the decrease in number and

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size of the neuroblastic nodules over gestation. Chromaffin cells appear at the periphery of the neuroblastic nodules and can be recognized as cells that are larger than the neuroblasts, with larger nuclei showing a finely granular chromatin pattern and cytoplasm containing brownish granules (Fig. 20.19). Therefore, chromaffin cells are seen in greater numbers in later gestational ages. In the past, there has been controversy about how to deal with neuroblastic cells seen in the adrenal gland after birth. Is this a normal variant? Are these developmental remnants? Does it represent a precursor to malignancy? Is it a frank malignancy?

Fig. 20.15  Involution of fetal cortex/zone. Higher-power view of the adrenal gland from a preterm infant who lived for 6 weeks postnatally. The involuting fetal zone (FZ) is seen between the definitive zone (DZ) and medulla (M). The polygonal epithelial cells of the fetal zone show degenerative/involutional changes. Congestion and hemorrhage are also present (H&E, 20×)

Fig. 20.16  Adrenal gland at 17 weeks of gestation. This section from near the center of the gland shows irregular clusters of neuroblastic cells, characterized by their small, round, hyperchromatic nuclei and scant cytoplasm, located within the fetal zone of the cortex. Note the adjacent small nerve twig (arrow) close to the cluster of neuroblastic cells (H&E, 10×)

Fig. 20.17  Adrenal gland at 20  weeks of gestation. The clusters of neuroblastic cells have migrated through the cortex and come to rest in the center of the gland (H&E, 10×)

Fig. 20.18  Adrenal gland at 39 weeks of gestation. In this low-power view of the central region of a term adrenal gland, note that the neuroblastic cells are seen mostly surrounding the central adrenal vein (H&E, 10×)

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Fig. 20.19  Adrenal gland at 39  weeks of gestation. In this higher-­ Fig. 20.20  Nodule of adrenal cortical tissue adjacent to the capsule at power view of the center of a term adrenal gland, medullary cells appear 35 weeks of gestation. Note that it has components of both definitive less neuroblastic and take on a more chromaffin-like appearance, with and fetal cortex (H&E, 10×) more abundant, granular cytoplasm (H&E, 20×)

The presence of incidentally encountered microscopic nodules of neuroblastic cells with cytological features of neuroblastoma in the adrenal glands of children led to the concept of in situ neuroblastoma of the adrenal gland [12]. Beckwith and Perrin [12] noted that these lesions were seen only microscopically, had no associated metastases, and were otherwise clinically insignificant. The prevalence of the lesions at autopsy suggested to the authors that most in situ neuroblastomas may actually regress or differentiate into normal or hyperplastic chromaffin cells. Shimada [13] also noted that, to date, no clonal proliferation has been demonstrated in these lesions and that the photomicrographs provided of in situ neuroblastoma in the original manuscript had the morphologic characteristics of favorable histology neuroblastoma, a lesion now known to be in the process of regression or undergoing age-appropriate maturation.

Special Considerations Heterotopic Adrenal Tissue Small nodules of cortical tissue are frequently seen adjacent to the adrenal gland and are generally not considered to represent an abnormality (Fig.  20.20). They may be a reflection of the plane of section and/or irregularity of the gland. However, the finding of normal-appearing adrenal tissue in an abnormal location is not infrequent in fetal autopsy material (Fig.  20.21). The most frequent locations are adjacent to the kidney, gonads, gonadal vessels, fallopian tube, epididymis, and vas deferens. The intimate association of the developing adrenal gland with the mesonephros and gonad accounts for the most common locations of adrenal heterotopias. In our experience, the heterotopic adrenal tissue usually consists only of cortical tissue. These small nodules of

Fig. 20.21  Heterotopic adrenal tissue in a perigonadal location. Two nodules of normal-appearing adrenal tissue (arrows) are noted between the fallopian tube (FT) and ovary (OV). Note that this adrenal tissue consists of cortical tissue from the definitive zone and the fetal zone (H&E, 4×)

adrenal cortex contain both fetal and definitive cortex and are likely to be embryonic remnants that have followed an aberrant migratory course. In most instances, these adrenal heterotopias have no clinically significant effects on the fetus. More remote sites for heterotopic adrenal tissue have been reported, such as the liver [14], central nervous system [15–17], lung [18], and placenta [19, 20].

Adrenal Cytomegaly The fetal cortex occasionally contains cells with markedly enlarged nuclei at least three times the size of the adjacent fetal cortical cells. They also appear hyperchromatic, with

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nuclear atypia and occasional intranuclear cytoplasmic pseudoinclusions (Figs.  20.22 and 20.23). The presence of these cells is not specific for a single condition; it has been described in patients with Beckwith-Weidemann syndrome [21, 22], the cytomegalic type of congenital adrenal hypoplasia [23], trisomy 13 and in infants of diabetic mothers. Adrenal cytomegaly also may be seen as an unexpected finding without any clinical correlation.

L. M. Ernst and L. Marsden

 denoid Change in the Definitive Zone A of the Cortex The definitive cortical zone in fetal life frequently is not homogeneous and shows cystic spaces within the layer. These round to irregular spaces are lined by the resident cortical cells, which surround a slightly basophilic-appearing, proteinaceous fluid (Fig. 20.24). This has been referred to as adenoid change or pseudofollicular change, and its pathogenesis is uncertain. It may represent cytolysis of cortical cells that experience oxidative stress from hypoxic ischemic injury [24, 25], but this finding should be interpreted with caution because it also may represent a normal developmental process, especially in very immature fetuses [26, 27].

Fig. 20.22  Adrenal cytomegaly. Low-power view of fetal adrenal gland at 22 weeks of gestation showing adrenal cytomegaly within the fetal cortical cells. Note the patchy enlargement of nuclei at least three times larger than normal adjacent fetal cortical nuclei (H&E, 10×)

Calcifications Calcifications are generally not a feature of the normal adrenal gland (Fig.  20.25). When present, they are thought to represent a consequence of adrenal hemorrhage. Though the etiology of adrenal hemorrhage in prenatal life has not been fully elucidated, when found incidentally in infants, it has been attributed to a history of anoxia/hypoxia during delivery, large-for-gestational-age size, prolonged labor, difficult extraction, and birth trauma during either delivery or postnatal resuscitation [28–30]. This would suggest that episodes of severe intrauterine stress leading to hypoxia may explain adrenal hemorrhage during fetal life [28, 31]. Hypoxia causes redistribution of blood to the central nervous system, heart, and adrenal glands, resulting in congestion, and when combined with hypoxically damaged endothelial cells, this congestion can lead to hemorrhage and subsequent develop-

Fig. 20.23  Adrenal cytomegaly. Higher-power view of fetal adrenal gland at 22 weeks of gestation showing adrenal cytomegaly within the fetal cortical cells. Note the intranuclear cytoplasmic pseudoinclusion (H&E, 20×)

Fig. 20.24  Adenoid change in the fetal adrenal gland. Adrenal gland at 18 weeks of gestation showing definitive adrenal cortex with adenoid change. Note the round to irregular spaces containing basophilic material (H&E, 20×)

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Fig. 20.25  Adrenal gland with loss of nuclear basophilia, but evident calcification seen in the fetal cortex. No associated mass or other pathologic changes are noted (H&E, 4×)

ment of calcifications [32]. Adrenal calcifications have also been reported in a case of congenital heart disease (attributed to circulatory insufficiency and heart failure), in the setting of storage diseases such as Wolman disease, and rarely in association with tumors such as neuroblastoma [28, 31, 33]. When no mass or other adrenal disease is identified, it can be presumed that calcifications represent the resolution of previous adrenal hemorrhage.

 atty Change in the Fetal Zone of the Cortex F The adrenal cortex plays an important role in steroid production and requires transport of lipid to function. Lipid is transported to the adrenal gland from body stores via the adrenal arteries, circulates through the sinusoids, and then exits via the central adrenal vein [34]. Under normal conditions, lipids are imported and metabolized by fetal adrenal cells, but accumulation of lipid vacuoles is not a prominent feature in the fetal adrenal cortical cells. Under hypoxic stress, h­ owever, the transport and metabolism of lipid can be impaired, leading to accumulation of the lipid within adrenal cortical cells [34]. The distribution pattern of the excess lipid (fatty change) may assist in determining the chronicity of the hypoxic injury, and a grading system has been proposed [35–37]: • Type I (acute): no or minimal lipid accumulation bordering the central vein • Type II (subacute): more extensive lipid accumulation away from the central vein • Type III (chronic): lipid accumulation throughout the entire fetal zone

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Fig. 20.26  Fatty change in the fetal adrenal gland. Adrenal gland at 19 weeks of gestation showing fetal adrenal cortex with fatty change. Note the round, lipid-filled vacuoles within the cytoplasm of the fetal cortical cells (arrows) (H&E, 20×)

Some authors have used immunostains for adipophilin to assist in demonstration of intracellular lipid [34]; but lipid accumulation is generally fairly easy to recognize on H&E-­ stained sections of fetal tissue as the presence of clear, round, lipid vacuoles within the eosinophilic cytoplasm of the fetal cortical cells (Fig. 20.26). Fatty change recognized in fetal adrenal cortical cells most likely represents abnormal accumulation of lipid and is generally not a normal finding in the fetal adrenal gland.

Extramedullary Hematopoiesis Occasional islands of extramedullary hematopoiesis may be found in the normal fetal adrenal gland. These likely represent secondary areas of extramedullary expansion. If more than occasional, they should be interpreted as evidence of hypoxic stress or fetal anemia.

References 1. Hansen K, Sung CJ, Huang C, Pinar H, Singer DB, Oyer CE. Reference values for second trimester fetal and neonatal organ weights and measurements. Pediatr Dev Pathol. 2003;6:160–7. 2. Siebert J.  Perinatal, fetal and embryonic autopsy. In: Gilbert-­ Barness E, editor. Potter’s pathology of the fetus, infant, and child. 2nd ed. Philadelphia: Mosby Elsevier; 2007. p. 726–7. 3. Rainey WE, Rehman KS, Carr BR. Fetal and maternal adrenals in human pregnancy. Obstet Gynecol Clin N Am. 2004;31:817–35. 4. Sadler TW, Langman J. Langman’s medical embryology. 11th ed. Philadelphia: Wolters Kluwer Lippincott Williams & Wilkins; 2010. 5. Schoenwolf GC, Larsen WJ. Larsen’s human embryology. 4th ed. Philadelphia: Elsevier/Churchill Livingstone; 2009.

222 6. Hervonen A. Development of catecholamine-storing cells in human fetal paraganglia and adrenal medulla. A histochemical and electron microscopical study. Acta Physiol Scand Suppl. 1971;368:1–94. 7. Turkel SB, Itabashi HH. The natural history of neuroblastic cells in the fetal adrenal gland. Am J Pathol. 1974;76:225–44. 8. Mesiano S, Jaffe RB. Developmental and functional biology of the primate fetal adrenal cortex. Endocr Rev. 1997;18:378–403. 9. Pepe GJ, Albrecht ED. Regulation of the primate fetal adrenal cortex. Endocr Rev. 1990;11:151–76. 10. Suzuki T, Sasano H, Takeyama J, Kaneko C, Freije WA, Carr BR, Rainey WE. Developmental changes in steroidogenic enzymes in human postnatal adrenal cortex: immunohistochemical studies. Clin Endocrinol. 2000;53:739–47. 11. Coupland RE. The natural history of the chromaffin cell—twenty-­ five years on the beginning. Arch Histol Cytol. 1989;52:331–41. 12. Beckwith JB, Perrin EV.  In situ neuroblastomas: a contribu tion to the natural history of neural crest tumors. Am J Pathol. 1963;43:1089–104. 13. Shimada H.  In situ neuroblastoma: an important concept related to the natural history of neural crest tumors. Pediatr Dev Pathol. 2005;8:305–6. 14. Vestfrid MA.  Ectopic adrenal cortex in neonatal liver. Histopathology. 1980;4:669–72. 15. Drut R, Quijano G, Altamirano ME, Jones MC, Maffessoli OB.  Vascular malformation and choroid plexus adrenal heterotopia: new findings in Beckwith-Wiedemann syndrome? Fetal Pediatr Pathol. 2006;25:191–7. 16. Kepes JJ, O’Boynick P, Jones S, et  al. Adrenal cortical adenoma in the spinal canal of an 8-year-old girl. Am J Surg Pathol. 1990;14:481–4. 17. Mitchell A, Scheithauer BW, Sasano H, Hubbard EW, Ebersold MJ. Symptomatic intradural adrenal adenoma of the spinal nerve root: report of two cases. Neurosurgery. 1993;32:658–61; discussion 661–2 18. Marchevsky AM. Lung tumors derived from ectopic tissues. Semin Diagn Pathol. 1995;12:172–84. 19. Guschmann M, Vogel M, Urban M.  Adrenal tissue in the placenta: a heterotopia caused by migration and embolism? Placenta. 2000;21:427–31. 20. Labarrere CA, Caccamo D, Telenta M, Althabe O, Gutman R.  A nodule of adrenocortical tissue within a human placenta: light microscopic and immunocytochemical findings. Placenta. 1984;5:139–43. 21. Beckwith J.  Macroglossia, omphalocele, adrenal cytomegaly, gigantism, and hyperplastic visceromegaly. Birth Defects. 1969;5:188–96.

L. M. Ernst and L. Marsden 22. Wiedemann HR.  Familial malformation complex with umbili cal hernia and macroglossia–a “new syndrome”? J Genet Hum. 1964;13:223–32. 23. Mamelle JC, David M, Riou D, Gilly J, Trouillas J, Dutruge J, Gilly R. Congenital adrenal hypoplasia of cytomegalic type. Recessive, sex-linked form. Apropos of 3 cases [in French]. Arch Fr Pediatr. 1975;32:139–59. 24. de Sa DJ.  Adrenal changes in chorioamnionitis. Arch Dis Child. 1974;49:149–51. 25. de Sa DJ.  Stress response and its relationship to cystic (pseudofollicular) change in the definitive cortex of the adrenal gland in stillborn infants. Arch Dis Child. 1978;53:769–76. 26. Lacson A, Desa DJ.  Endocrine glands. In: Gilbert-Barness E, editor. Potter’s pathology of the fetus, infant, and child. 2nd ed. Philadelphia: Mosby Elsevier; 2007. p. 1630. 27. Rodin AE, Hsu FL, Whorton EB.  Microcysts of the permanent adrenal cortex in perinates and infants. Arch Pathol Lab Med. 1976;100:499–502. 28. Varan B, Saygili A, Tokel K, Atalay H, Mercan S, Cemil T. Incidental findings of bilateral adrenal calcification in a patient with congenital heart disease. South Med J. 2002;95:784–5. 29. Naidech HJ, Chawla HS. Bilateral adrenal calcifications at birth in a neonate. AJR Am J Roentgenol. 1983;140:105–6. 30. Abdu AT, Kriss VM, Bada HS, Reynolds EW. Adrenal hemorrhage in a newborn. Am J Perinatol. 2009;26:553–7. 31. Ryan SP, Gaisie G, Krill DE Jr. Adrenal calcification: an incidental finding or not? Pediatr Radiol. 1997;27:821–3. 32. Nissenbaum M, Jequier S. Enlargement of adrenal glands preceding adrenal hemorrhage. J Clin Ultrasound. 1988;16:349–52. 33. Shenoy P, Karegowda L, Sripathi S, Mohammed N. Wolman disease in an infant. BMJ Case Rep. 2014;2014:bcr2014203656. 34. Taweevisit M, Atikankul T, Thorner PS. Histologic changes in the adrenal gland reflect fetal distress in hydrops fetalis. Pediatr Dev Pathol. 2014;17:190–7. 35. Delprado WJ, Baird PJ. The fetal adrenal gland: lipid distribution with associated intrauterine hypoxia. Pathology. 1984;16:25–9. 36. Delprado WJ, Baird PJ. The fetal adrenal gland: definitive cortex cystic change, lipid patterns, and their relationship to fetal disease and maturity. Pathology. 1984;16:312–7. 37. Becker MJ, Becker AE.  Fat distribution in the adrenal cortex as an indication of the mode of intrauterine death. Hum Pathol. 1976;7:495–504.

21

Thyroid Gland Linda M. Ernst

Introduction By the beginning of the fetal period, the thyroid gland has attained its proper anatomical position anterior to the laryngeal and tracheal structures and has acquired its bilobed shape, with the two lobes connected by a narrow isthmus. However, the histological appearance of the fetal thyroid gland is quite different from that of the mature adult thyroid gland, especially early in the second trimester. Therefore, the tissues of the fetal thyroid gland may be overlooked or mischaracterized. This chapter reviews the histological changes in the thyroid gland over the fetal period and discusses the two main functional components of the thyroid gland: the thyroid follicles, which produce colloid, and the parafollicular C cells, which produce calcitonin.

Embryology The primordium of the thyroid gland begins to form in the posterior portion of the primitive tongue in the fourth week of gestation as an endodermal mass at the apex of the foramen cecum, a depression or shallow diverticulum in the midline of the sulcus terminalis [1, 2]. After formation, the endodermal mass begins migrating caudally in the midline anterior to the pharynx and remains connected to the foramen cecum by the thyroglossal duct. By approximately the fifth week of gestation, the thyroglossal duct begins to break down, and the thyroid mass becomes isolated from the tongue. As the thyroid mass descends, it begins to develop a bilobed shape. It migrates anterior to the hyoid bone and larynx to reach its final destination in front of the trachea by the seventh week of gestation. By this time, the thyroid gland has also acquired L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected]

its final shape, with two lateral lobes connected inferiorly by a thin isthmus. This endodermal tissue derived from the foramen cecum gives rise to the numerous follicles of the thyroid gland. The fetal thyroid follicular cells begin to produce colloid as early as 10–12 weeks of gestation [3–5]. The parafollicular C cells of the thyroid gland migrate from an anatomically and embryologically distinct source, the ultimobranchial bodies, derived from the somewhat controversial fifth pharyngeal pouch (immediately inferior to the fourth pharyngeal pouch). The cells that populate the paired ultimobranchial bodies are derived from the neural crest [6], and once populated by these cells, the ultimobranchial bodies detach from the pharynx and migrate caudally to become embedded into the upper lateral aspect of the thyroid gland.

Histology The fetal thyroid gland is best examined by sampling sequential transverse sections of the “neck block,” which includes all the soft tissues of the neck, posterior to the strap muscles, from the soft palate to the trachea (Figs. 21.1 and 21.2). This allows for examination of the laryngeal tissues, pharyngeal tissues, and thyroid gland resting anterior to the thyroid cartilage. A thin, fibrous capsule surrounds the thyroid gland and is sometimes closely approximated to the strap muscles of the neck. On low-power histological examination, the fetal thyroid gland frequently appears more cellular than the adult thyroid gland because thyroid follicles are generally smaller in the fetus, and colloid is less abundant. In addition, the fetal thyroid gland undergoes postmortem autolytic changes relatively quickly, which can lead to a collapse of the glandular architecture and the appearance of a nearly solid sheet of epithelium (Fig. 21.3). In well-preserved second-trimester and third-trimester fetal thyroid tissue, tightly packed, round follicular elements containing colloid can be appreciated throughout the gland.

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_21

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Fig. 21.1  Fetal neck structures including the thyroid gland. This anterior view of fetal neck organs shows the horseshoe-shaped fetal thyroid gland resting over the thyroid and cricoid cartilages

a

Fig. 21.3  Autolysis of the fetal/neonatal thyroid gland. A frequent problem in fetal autopsy specimens is rapid autolysis of the thyroid gland, with a collapse of the glandular architecture and nearly solid appearance of the gland, which does not reflect the normal expected

Fig. 21.2  Fetal neck structures after serial sectioning in the transverse plane. Note the beefy red thyroid gland anterior to the laryngeal cartilages. All of these tissue sections can be submitted for histological examination

b

follicular development. (a) Thyroid gland from a 27-week live-born infant. (b) Thyroid gland from a 32-week live-born infant (hematoxylin and eosin (H&E), 10×)

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Thyroid Follicles The follicles of the thyroid gland produce colloid, which contains the thyroid hormones necessary for proper fetal development, especially brain development. Thyroid ­follicles are

Fig. 21.4  Low-power view of well-preserved fetal thyroid gland at 15  weeks gestation. At low power, early fetal thyroid has a cellular, almost solid appearance, but on close inspection, the follicular development can be seen, with the largest follicles at the periphery of the gland. Note the thin, fibrous thyroid capsule. This section also includes a segment of laryngeal cartilages posterior to the gland (H&E, 4×)

a

Fig. 21.5  Higher-power views of the fetal thyroid gland at 15 weeks gestation. Note the small follicular formation throughout the gland. At the periphery, the follicles are slightly larger, and intrafollicular col-

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formed by predominantly cuboidal epithelial cells. With increased secretory activity, the follicular cells may become slightly taller, with more visible apical cytoplasm, which appears lightly eosinophilic. The lipofuscin pigment sometimes seen in adult thyroid follicular cells [7] is not identified in fetal follicular cells. Brown pigment within the fetal thyroid gland should raise suspicion for extrahepatic iron deposition, as seen in neonatal hemochromatosis, and should be confirmed by Prussian blue stain. The follicular cells rest their base on the follicular basement membrane. Their nuclei are round to oval, and a small nucleolus sometimes can be seen. The thyroid follicles change significantly in size and amount of colloid throughout gestation. Although fetal colloid production begins around 11 weeks gestation, intraluminal colloid is not a prominent feature of the fetal thyroid gland in the late first trimester to early second trimester (Figs. 21.4, 21.5, 21.6, and 21.7), and hormone production is limited until 18–20 weeks gestation [8]. Lightly stained eosinophilic secretions are seen within the follicles in the previable stage (Figs.  21.8 and 21.9). In the early third trimester, colloid becomes more prominent within follicles (Figs. 21.10 and 21.11). In general, the thyroid follicles in the fetus appear larger toward the periphery of the gland, and this may represent a progressive maturation of the follicles from center to periphb

loid is scanty. Also note the rich capillary network in the interfollicular interstitial tissues and the delicate fibrous capsule (H&E, (a) 10×; (b) 20×)

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ery. Histochemical studies have shown that thyroid colloid stains with periodic acid-­Schiff and toluidine blue stains [9]. The colloid in the larger follicles demonstrates slightly more toluidine staining than small ones [9]. By near term

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(Figs.  21.12 and 21.13) to term (Figs.  21.14 and 21.15), there is enough colloid present to distend some of the follicles, especially at the periphery of the gland.

C Cells

Fig. 21.6  Low-power view of the fetal thyroid gland at 18 weeks gestation (H&E, 4×)

a

The parafollicular C cells, responsible for calcitonin production, are not easily recognized with routine H&E stains, especially in the fairly cellular-appearing fetal thyroid gland. When recognized, C cells are described as polygonal, with a paler eosinophilic cytoplasm than the follicular cells, and a round to oval nucleus with a central nucleolus [7]. To aid in their recognition, C cells can be highlighted by staining their secretory granules with the Grimelius silver nitrate argyrophil method. Calcitonin immunohistochemistry is also a ­convenient method to highlight the C cells; it distinguishes small groups of cells or single cells within the deep, central region of the middle third of the lateral lobes (Fig. 21.16). C cells are closely associated with the thyroid follicle (Figs. 21.17 and 21.18), and by ultrastructural analysis, most C cells are found within the basement membrane of the follicle and are separated from the colloid by the follicular cell [10].

b

Fig. 21.7  Higher-power views of the fetal thyroid gland at 18 weeks gestation. Note the small follicular formation throughout the gland. At the periphery, the follicles are slightly larger. Intrafollicular colloid is slightly more prominent than at 15 weeks gestation (H&E, (a) 10×; (b) 20×)

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Stroma The fetal thyroid gland is composed overwhelmingly of closely packed follicles, but between the follicles, there is a small amount of interstitial fibrous tissue and numerous blood vessels. Inflammatory cells are usually not a feature of the fetal or neonatal thyroid stroma.

Special Considerations Intrathyroidal Inclusions

Fig. 21.8  Low-power view of the fetal thyroid gland at 21 weeks gestation (H&E, 4×)

a

In the 1970s, Carpenter and Emery [11] examined hundreds of fetal, neonatal, and pediatric thyroid glands. These authors described numerous inclusions of nonthyroidal tissue within the thyroid gland, including parathyroid, thymus, cartilage, ciliated epithelium, fat, and striated muscle. The

b

Fig. 21.9  Higher-power views of the fetal thyroid gland at 21 weeks gestation. Follicular formation and intrafollicular colloid continue to increase (H&E, (a) 10×; (b) 20×)

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Fig. 21.10  Low-power view of the fetal thyroid gland at 26 weeks gestation (H&E, 4×)

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presence of most of these ectopic tissues within the thyroid gland was considered a normal finding by the authors, related to the pharyngeal origin of the thyroid gland and its descent with thymus and parathyroid glands. Striated muscle may represent disorganization of cervical tissues or, as the authors suggest, a possible association with myopathies. The pyramidal lobe is often admixed with skeletal muscle. From the same large review of thyroid glands, Sagreiya and Emery [12] also introduced the concept of “perinatal thyroid discharge,” which is characterized by the reduction or loss of colloid (not postmortem-related), vacuolation or detachment of epithelial cells, irregular and pyknotic nuclei, and nuclear knots. The authors felt that these changes represented pathologic changes and did not consider them normal or related to autolysis. Studies of this type have not been repeated, however, and it can be extremely difficult to sort out autolytic changes from true pathologic changes in the examination of the thyroid gland of the stillborn fetus.

b

Fig. 21.11  Higher-power views of the fetal thyroid gland at 26  weeks gestation. Follicular formation and intrafollicular colloid continue to increase (H&E, (a) 10×; (b) 20×)

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Ultimobranchial Body Remnants

Fig. 21.12  Low-power view of the fetal thyroid gland at 34 weeks gestation (H&E, 4×)

a

Also known as “solid cell nests,” ultimobranchial body remnants are small, well-defined collections of cells found in the same location as the C cells, interspersed between thyroid follicles. They are considered to contain two cell types: “main cells” and “C cells.” The main cells are more abundant in solid cell nests and consist of polygonal epithelial cells with abundant eosinophilic cytoplasm demonstrating a squamoid appearance (Fig. 21.19). These main cells are positive for p63. The C cells of the solid cell nests are less conspicuous, but do mark with calcitonin immunohistochemistry. The nature of the solid cell nest is still controversial; some investigators believe that the main cells may represent a stem cell population and suggest a link between solid cell nests and thyroid carcinoma [7, 13–15].

b

Fig. 21.13  Higher-power views of the fetal thyroid gland at 34  weeks gestation. Follicular formation and intrafollicular colloid continue to increase (H&E, (a) 10×; (b) 20×)

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Fig. 21.14  Fetal thyroid gland at term. This low-power view shows nearly all follicles with colloid and some distension of follicles (H&E, 1×)

Fig. 21.16  C cells within the lateral lobe of thyroid gland. This low-­ power view of the lateral lobe of the thyroid gland at term shows calcitonin stain-positive cells within the middle portion of the lobe (calcitonin immunohistochemistry, 1×)

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Fig. 21.15  Fetal thyroid gland at term. This higher-power view shows the details of the predominantly cuboidal follicular cells with oval nuclei and a characteristic coarsely granular chromatin pattern. Focal colloid scalloping is noted (H&E, 20×)

Fig. 21.17  C cells in the fetal thyroid gland. A higher-power view of term thyroid gland shows darkly stained parafollicular C cells. Note how intimately associated the C cells are with the basement membrane of the thyroid follicle and how they occur as single cells or in small clusters (calcitonin immunohistochemistry, 20×)

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Fig. 21.18  C cell in the fetal thyroid gland. This close-up view is of one C cell, which appears to be intrafollicular (calcitonin immunohistochemistry, 60×)

References 1. Schoenwolf GC, Larsen WJ. Larsen’s human embryology. 4th ed. Philadelphia: Elsevier/Churchill Livingstone; 2009. 2. Sadler TW, Langman J.  Langman’s medical embryology. 11th ed. Philadelphia: Wolters Kluwer Lippincott Williams & Wilkins; 2010. 3. Zoeller RT, Crofton KM.  Thyroid hormone action in fetal brain development and potential for disruption by environmental chemicals. Neurotoxicology. 2000;21:935–45. 4. Park SM, Chatterjee VK. Genetics of congenital hypothyroidism. J Med Genet. 2005;42:379–89. 5. Contempré B, Jauniaux E, Calvo R, Jurkovic D, Campbell S, de Escobar GM. Detection of thyroid hormones in human embryonic cavities during the first trimester of pregnancy. J Clin Endocrinol Metab. 1993;77:1719–22. 6. De Felice M, Di Lauro R. Thyroid development and its disorders: genetics and molecular mechanisms. Endocr Rev. 2004;25:722–46. 7. Carcangiu M. Thyroid. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 1185–207.

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Fig. 21.19  Solid cell nests (also known as ultimobranchial body remnants) in the thyroid. Note the interfollicular cluster of polygonal cells with squamoid features. C cells are not easily recognized (H&E, 20×) 8. Fisher DA, Greuters A.  Disorders of the thyroid in the newborn and infant. In: Sperling MA, editor. Pediatric endocrinology. 3rd ed. Philadelphia: Saunders/Elsevier; 2008. p. 198–226. 9. Shepard TH, Andersen H, Andersen HJ. Histochemical studies of the human fetal thyroid during the first half of fetal life. Anat Rec. 1964;149:363–79. 10. Hazard JB.  The C cells (parafollicular cells) of the thyroid gland and medullary thyroid carcinoma. A review. Am J Pathol. 1977;88:213–50. 11. Carpenter GR, Emery JL. Inclusions in the human thyroid. J Anat. 1976;122:77–89. 12. Sagreiya K, Emery JL. Perinatal thyroid discharge. A histological study of 1225 infant thyroids. Arch Dis Child. 1970;45:746–54. 13. Cameselle-Teijeiro J, Preto A, Soares P, Sobrinho-Simoes M. A stem cell role for thyroid solid cell nests. Hum Pathol. 2005;36:590–1. 14. Burstein DE, Unger P, Nagi C, Wang BY.  Thinking “out of the nest”—a reply to “a stem-cell role for thyroid solid cell nests [letter]”. Hum Pathol. 2005;36:591–2. 15. Burstein DE, Nagi C, Wang BY, Unger P.  Immunohistochemical detection of p53 homolog p63 in solid cell nests, papillary thyroid carcinoma, and Hashimoto’s thyroiditis: a stem cell hypothesis of papillary carcinoma oncogenesis. Hum Pathol. 2004;35:465–73.

22

Parathyroid Gland Linda M. Ernst and Lily Marsden

Introduction Studies in mammals have shown that the fetus maintains a plasma calcium concentration that is higher than that of the mother. This high fetal calcium level is essential for the proper development and mineralization of the fetal skeleton, and the fetal parathyroid glands play an important role in this process. Hormones elaborated by the fetal parathyroid gland, including parathyroid hormone-related protein, are required to maintain the fetal calcium gradient created through active placental transport of calcium by syncytiotrophoblast cells [1]. This chapter focuses on the histological features of the fetal parathyroid glands.

Embryology The parathyroid glands are derived from the third and fourth pharyngeal pouches, beginning in the fifth week of gestation. Although one might expect the more cephalad pharyngeal pouch to give rise to the superior parathyroid gland and the more caudal pouch to give rise to the inferior gland, the reality is actually reversed. The two paired inferior parathyroid glands are derived from the bilateral dorsal buds of the third pharyngeal pouches, whereas the two superior parathyroid glands originate from the fourth pharyngeal pouches. The nodular primordia of the parathyroid glands separate from the pharyngeal pouches, descend into the neck, and take their usual positions on the dorsal

L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected] L. Marsden Utah State Office of the Medical Examiner, Salt Lake City, UT, USA e-mail: [email protected]

aspect of the thyroid. Descent is typically completed by approximately 7 weeks of gestation [2, 3].

Histology Parathyroid glands can be difficult to identify grossly in fetuses, even with the aid of a dissecting microscope, but they can be identified histologically if the eviscerated neck organs are serially sectioned in the transverse plane and entirely submitted from the base of the tongue to just below the thyroid gland (see Chap. 21, Figs. 21.1 and 21.2). The parathyroid gland is surrounded by a thin connective tissue capsule from which septa extend into the parenchyma of the gland, conveying blood vessels, nerves, and lymphatics. The fetal and neonatal parathyroid glands are composed of only one parenchymal cell type, the chief cell [2, 4]. Oxyphil cells are not present until later in childhood (4.5–7 years of age) [2]. The chief cells within the parenchyma of the fetal parathyroid gland are polygonal cells with well-defined cell borders, a key feature in differentiating them from the lymphocytes of small cervical lymph nodes in this anatomic location (Figs.  22.1 and 22.2) or adjacent thyroid gland (Fig. 22.3), especially when the tissues are autolyzed. The parenchymal cells of the parathyroid gland are arranged in sheets, cords, and trabeculae. In contrast to the adult parathyroid gland, the fetal gland has no stromal fat, and the connective tissue septa consist mainly of a vascular network with very little supporting collagen in the perivascular stroma (Figs.  22.4 and 22.5). The cytoplasm of the chief cells is clear to lightly eosinophilic (Figs. 22.6, 22.7, and 22.8), and the chromatin of the nuclei has a vague “salt and pepper” appearance (Fig.  22.9). Ultrastructural studies have shown that the chief cells are rich in glycogen particles, with occasional secretory granules, lipid bodies, and multivesicular bodies [4]. Immunostaining with antiserum to bovine parathyroid

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hormone has identified immunoreactive p­ arathyroid hormone in fetuses as early as at 10 weeks of gestation [5]. With careful microscopic inspection of the cross sections of the neck organs, it is not unusual for a parathyroid gland to be found posterior to or within the thyroid gland (Fig.  22.10) or with an associated undescended thymic lobule (Figs.  22.11, 22.12, and 22.13), given the common embryonic origin of the thymus and the inferior parathyroid glands. Other sites where parathyroid glands can be found include within the thymus gland proper and in the pericardium or in paratracheal or paraesophageal tissues.

Fig. 22.1  Parathyroid gland versus lymph node. Low-power view of two nodules within the cervical tissues. At low power it may be difficult to distinguish parathyroid gland from lymph node or ectopic thymic tissue, especially in the setting of any significant autolysis. In this image, the nodule on the left is a parathyroid gland and the nodule on the right is a lymph node. Thymic tissue is above. See Fig.  22.2 for a higherpower view of cellular details (hematoxylin and eosin (H&E), 4×)

a

Fig. 22.2  Parathyroid gland versus lymph node. Higher-power view for comparison of parathyroid gland (a) and lymph node (b). (a) The parathyroid gland shows a trabecular arrangement of epithelial cells with

b

lightly eosinophilic cytoplasm and well-defined cell borders. (b) The lymph node shows a more diffuse population of small lymphocytes without significant cytoplasm or well-defined cell borders ((a, b) H&E, 20×)

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Fig. 22.3 Parathyroid gland versus autolyzed thyroid tissue. Comparison of parathyroid gland (a) and thyroid gland with autolysis (b). (a) The parathyroid gland shows a trabecular arrangement of epithelial cells with lightly eosinophilic cytoplasm and well-defined cell

borders. (b) The thyroid gland, when autolyzed, can have a collapsed follicular architecture and resemble the trabecular arrangement of cells in the parathyroid gland ((a) H&E, 20×; (b) H&E, 10×)

Fig. 22.4  Medium-power view of the fetal parathyroid gland at 18 weeks of gestation. This small parathyroid gland is located posterior to the thyroid lobe (above it). In contrast to the thyroid gland, the parathyroid gland is usually well preserved, as it undergoes autolysis more slowly. The cytoplasm of the parenchymal cells is clear and the nuclei are round and centrally placed. Cell borders are well demarcated and no stromal fat is present (H&E, 20×)

Fig. 22.5  Lower-power view of the fetal parathyroid gland at 24 weeks of gestation. The gland shown here is larger than the one at 18 weeks (see Fig. 22.4) and is also found adjacent to the thyroid gland (THR) in the perithyroidal fat and connective tissue. Note the thin fibrous capsule and rich vascular network, which are visible at low power (H&E, 10×)

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Fig. 22.6  Higher-power view of the fetal parathyroid gland at 24 weeks of gestation. The parenchymal cells, chief cells, are fairly monotonousappearing polygonal cells arranged in cords and trabeculae. Cell borders are well defined, and the cytoplasm appears clear to lightly eosinophilic (H&E, 20×)

Fig. 22.7  High-power view of the fetal parathyroid gland at 24 weeks of gestation. Note the thin, fibrous capsule and detail of the parenchymal chief cells (H&E, 40×)

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Fig. 22.8  Fetal parathyroid gland at 39 weeks of gestation. The gland is larger still, and the parenchymal cells appear more tightly packed than in mid gestation; compare with Fig. 22.6 (H&E, 20×)

Fig. 22.9  High-power view of the fetal parathyroid gland at 39 weeks of gestation (H&E, 40×)

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Fig. 22.10  Intrathyroidal parathyroid gland (arrow) identified within the thyroid parenchyma at 34 weeks of gestation (H&E, 10×)

Fig. 22.11  Term gestation with identification of parathyroid gland adjacent to an ectopic, undescended thymic lobule above it (H&E, 4×)

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Fig. 22.13  Parathyroid gland embedded within an ectopic, undescended thymic lobule. A higher magnification of parathyroid gland (black arrows) within the cervical thymic lobule is shown. The white arrow denotes a Hassall’s corpuscle (H&E, 10×) Fig. 22.12  Ectopic, undescended thymic lobule with embedded parathyroid gland. This is a low-power view of an ectopic thymic lobule in the cervical region, intimately associated with parathyroid gland tissue, indicated by arrows (H&E, 4×)

References 1. MacIsaac RJ, Heath JA, Rodda CP, Moseley JM, Care AD, Martin TJ, Caple IW. Role of the fetal parathyroid glands and parathyroid hormone-related protein in the regulation of placental transport of calcium, magnesium and inorganic phosphate. Reprod Fertil Dev. 1991;3:447–57. 2. Valdes-Dapena MA.  Histology of the fetus and newborn. Philadelphia: Saunders; 1979.

3. Schoenwolf GC, Larsen WJ. Larsen’s human embryology. 4th ed. Philadelphia: Elsevier/Churchill Livingstone; 2009. 4. Nakagami K, Yamazaki Y, Tsunoda Y.  An electron microscopic study of the human fetal parathyroid gland. Z Zellforsch Mikrosk Anat. 1968;85:89–95. 5. Leroyer-Alizon E, David L, Anast CS, Dubois PM. Immunocytological evidence for parathyroid hormone in human fetal parathyroid glands. J Clin Endocrinol Metab. 1981;52: 513–6.

23

Pituitary Gland Linda M. Ernst

Introduction Despite its small size, the fetal pituitary gland produces a variety of important hormones. Normal development of the human fetal pituitary gland is essential for fetal homeostasis, as well as for the appropriate differentiation and maturation of several organ systems required for extrauterine survival [1]. This chapter reviews the histological features of the adenohypophysis (the anterior lobe of the pituitary gland) and the neurohypophysis (the posterior lobe of the pituitary gland). In addition, some features unique to the fetal and neonatal pituitary gland, such as laminated calcifications within the anterior lobe, are discussed.

The posterior wall of the cyst remains thin and is in close contact with the infundibulum; it will become the pars intermedia. The epithelium of the anterior wall of the cyst proliferates to form the rounded mass of the anterior lobe of the pituitary gland, the pars distalis (Fig.  23.1). A small group of cells from the posterior portion of the anterior lobe grow up the stalk of the infundibulum and are known as the pars tuberalis. The lumen of the original Rathke’s pouch becomes flattened in the anteroposterior direction and becomes a slit-like space

Embryology The pituitary gland is formed from two different ectodermally derived structures: Rathke’s pouch and a portion of the diencephalon known as the infundibulum. These distinctive embryologic origins correlate with the two distinct histological compartments of the pituitary gland, the adenohypophysis (anterior lobe), derived from Rathke’s pouch, and the neurohypophysis (posterior lobe), derived from the infundibulum [2–4]. Rathke’s pouch begins in the third week of gestation as an outgrowth of ectoderm from the stomodeum, the primitive mouth of the embryo. The epithelial outgrowth develops a lumen and forms an epithelium-lined diverticulum. This pouch extends dorsally toward the infundibulum, which is the small ventral extension of the floor of the diencephalon; it will become the neurohypophysis. At approximately 6 weeks gestation, Rathke’s pouch loses its connection with the surface ectoderm of the mouth and becomes an epithelium-­lined cyst. L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected]

Fig. 23.1  Low-power view of the entire fetal pituitary gland at 17 weeks gestation. The gland, removed with portions of the cartilaginous sella turcica (upper right), is shown here. The slit-like opening of Rathke’s pouch (RP) is visible. The most anterior portion of the anterior lobe or pars distalis (PD) is the rounded mass anterior to Rathke’s pouch. The thin rim of similar-appearing cells of the anterior pituitary on the opposite side of Rathke’s pouch is known as the pars intermedia (PI). The posterior lobe (PL) is adjacent to the PI and has a generally paler staining appearance than the anterior lobe (hematoxylin and eosin (H&E), 2×)

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between the pars distalis anteriorly and the pars intermedia posteriorly. Its remnants become the microscopic glandular structures of the pars intermedia [2–4]. The anterior lobe of the pituitary gland is responsible for elaboration of the pituitary hormones: growth hormone (GH), adrenal corticotropin hormone (ACTH), prolactin (PRL), thyroid-stimulating hormone (TSH), follicle-­stimulating hormone (FSH), and luteinizing hormone (LH). Differentiation of the ectodermal cells from Rathke’s pouch into the various hormone-producing cells depends on gradients of growth factors, such as fibroblast growth factor 8, which emanates from the ventral diencephalon, and bone morphogenetic protein 2, which emanates from the ventral pituitary organizing center. These overlapping gradients result in the expression of specific transcription factors that drive differentiation and the production of specific hormones [5–7]. The posterior lobe of the pituitary gland forms by the proliferation of neuroglial cells at the distal end of the infundibulum. The infundibulum remains the stalk of the pituitary gland, and nerve fibers from the hypothalamus also grow into the posterior lobe via the infundibular stalk.

L. M. Ernst

a

b

Histology General Overview Sampling the fetal pituitary gland for histological examination can be difficult because the tissues of the gland are extremely soft and are easily compressed if they are grasped by forceps. After removing the brain, the pituitary gland can be removed intact by excising it along with the bony tissues of the sella turcica. Using this technique, the surrounding cartilaginous and bony tissues of the sella turcica protect the pituitary gland and serve as a “handle” to grasp the gland. The entire gland with the rim of the cartilaginous and bony tissue can be submitted in toto for histologic examination after brief decalcification. Because of their separate embryologic origins, the anterior and posterior lobes of the pituitary gland have quite distinct histological appearances in the fetus. The anterior lobe of the pituitary gland appears as an ovoid, amphophilic mass of polygonal epithelial cells that are separated by a slit-like cavity from the thin layer of similar-appearing cells, forming the pars intermedia in the posterior aspect of the anterior lobe. Although well demarcated from the anterior lobe, the posterior lobe is in direct contact with the anterior lobe and stands in contrast, with its pale staining and fibrous appearance (Figs. 23.1, 23.2, 23.3, and 23.4). In routine sections of the fetal pituitary gland, the posterior lobe may not be readily identified, because it makes up a smaller proportion of the gland and the gland often has a random orientation. Deeper sections may be needed to assess the posterior pituitary gland.

Fig. 23.2  Higher-power views of the anterior lobe of the pituitary gland at 17 weeks gestation. (a) The anterior lobe is composed of the pars distalis and pars intermedia, separated by residual lumen of Rathke’s pouch. (b) This close-up view is of the cells lining the lumen of Rathke’s pouch. Note the cuboidal to columnar, palisading epithelial cells. ((a) H&E, 4×; (b) H&E, 20×)

Fig. 23.3  Low-power view of the pituitary gland at 26 weeks gestation. Note that the gland is larger, but the overall structure of the anterior and posterior lobes is identical (H&E, 2×)

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Fig. 23.4  Closer view of the junction between anterior and posterior pituitary lobes at 26  weeks gestation. Note the residual lumen of Rathke’s pouch and the lining epithelial cells. Glandular structures are formed within the pars intermedia. The posterior lobe (upper left) has a pale, fibrillary appearance (H&E, 4×)

Anterior Pituitary The anterior lobe of the pituitary gland is composed of epithelial cells arranged in cords and nests with an intervening delicate capillary network. The epithelial cells have round, slightly eccentrically placed nuclei and fairly abundant cytoplasm, which is characterized by an affinity for either acidophilic dye (acidophils), basophilic dye (basophils), or neither (chromophobes) on H&E staining. The acidophil cells appear to be the most numerous in the fetal pituitary gland and have brightly eosinophilic cytoplasmic granules. As a general rule, acidophil cells are considered to secrete GH and PRL and are located primarily in the rounded lateral projections of the pars distalis. Basophil cells are also commonly present and have dark bluestained cytoplasmic granules. Basophils that secrete ACTH and TSH are located primarily in the central region of the pars distalis, referred to as the mucoid wedge because the basophil cells are also positive using the periodic acid–Schiff method [8]. Basophil cells that secrete the gonadotropins FSH and LH are scattered throughout the pars distalis. Both acidophils and basophils are generally arranged in clusters throughout the gland (Fig.  23.5). Although chromophobe cells can represent up to 50% of cells in the adult pituitary gland [8], they are not very prominent in the fetal pituitary. They are seen as only small clusters of cells with poorly stained, inconspicuous cytoplasm. Immunohistochemical studies in rats indicate that ACTH is the first pituitary hormone, detectable on embryonic day 15, and that the other hormones become detectable in a predictable pattern over the next few days [9]. Immunohistochemical and ultrastructural studies of human fetal pituitary glands [10–12] have shown that ACTH-­ producing cells are seen as early as 6–8  weeks gestation.

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Features characteristic of somatotrophs, which secrete GH, are seen at 8–9  weeks, and cells producing most other hormones are identifiable by 15–16 weeks gestation. Lactotrophs, PRL-producing cells, are somewhat variable in number in the first and second trimesters but are relatively numerous near term. In general practice, immunohistochemical stains for the pituitary hormones can be performed on the human fetal pituitary gland and will reveal the presence of all the anterior pituitary hormones near term. The staining pattern for each hormone is different, varying in intensity and the number of cells stained, but is similar to the pattern for adult control pituitary glands. The hormones with the most intense and widespread staining in the anterior lobe are ACTH (Fig. 23.6) and GH (Fig. 23.7). A unique histological feature of the anterior lobe of the fetal and newborn pituitary is the presence of ovoid, calcified concretions (Fig.  23.8). These laminated calcifications have been noted in a majority of fetal pituitaries from mid-­ gestation to term, but they generally disappear within the first 6 months of life [13, 14]. Although the etiology of these calcifications is uncertain, hypotheses include a relationship to hormonal activity of the fetal pituitary gland or developmental processes associated with cell death [13, 14]. By adulthood, the lumen of Rathke’s pouch between the pars distalis and pars intermedia is no longer visible, but in the fetal and neonatal pituitary gland, the space may be present and lined by cuboidal to columnar epithelial cells (see Fig. 23.2). Persistence of a so-called Rathke cleft cyst (Fig. 23.9) is identified as an incidental finding in up to one third of autopsy cases [15, 16]. If large enough, the Rathke cleft cyst in adults and children can become symptomatic and require surgical intervention. Embryonic epithelial remnants of the obliterated stalk connecting Rathke’s pouch to the embryonic oral cavity may also persist in the submucosa of the nasopharynx as the “pharyngeal pituitary” or in the sphenoid bone along the embryonic path of the stalk; these embryonic rests may be functional, may be the origin of ectopic pituitary adenomas, and have been described with central nervous system malformations such as anencephaly, holoprosencephaly, and neural tube defects [17–20].

Posterior Pituitary The main cell type seen in the posterior pituitary is the pituicyte, which is best classified as a subtype of the glial cell. Pituicytes have ovoid to elongate nuclei, and most have fibrillary cytoplasmic processes (Fig. 23.10; see also Figs. 23.1, 23.3, and 23.4). In the adult pituitary gland, the posterior pituitary shows immunoreactivity for glial fibrillary acidic protein, vimentin, and S100 protein, but in fetal life, only very few glial fibrillary acidic protein-positive fibers are seen (Fig. 23.11). The morphologic features of pituicytes can

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Fig. 23.5 (a, b) Views of the anterior lobe from the pars distalis region at 26 weeks gestation. Cells of the anterior lobe are arranged in small clusters, cords, and nests with intervening capillary network. Acidophil

cells with eosinophilic cytoplasm are easy to distinguish from the basophil cells, with their dark, granular cytoplasm. Chromophobe cells are inconspicuous (H&E: a 20×; b 40×)

Fig. 23.6 Anterior pituitary gland at 35  weeks gestation. Immunohistochemical stain for ACTH shows numerous anterior pituitary cells with positive staining (ACTH immunohistochemistry, 20×)

Fig. 23.7 Anterior pituitary gland at 35  weeks gestation. Immunohistochemical stain for GH shows numerous anterior pituitary cells with positive staining (GH immunohistochemistry, 20×)

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Fig. 23.8 (a, b) Anterior lobe of the pituitary gland at 35 weeks gestation. The overall architecture and cellular components are similar to 26 weeks gestation (as in Fig. 23.5). Acidophil and basophil cells are easily distinguished by their cytoplasmic features. Chromophobe cells

are still inconspicuous. Note the ovoid, darkly basophilic concretions present in the anterior lobe. These calcified concretions, which appear partially laminated, are commonly seen in fetal and newborn pituitary glands, until approximately 6 months of age (H&E: a 20×; b 40×)

Fig. 23.9  Rathke cleft cyst in a 1-year-old infant. Note the large cystic space involving the anterior pituitary gland. The cyst is lined by cuboidal to columnar epithelial cells, which can have cilia (H&E, 1×)

Fig. 23.10  Posterior lobe of the pituitary gland at 35 weeks gestation. The posterior pituitary occupies most of the field, with a small amount of anterior pituitary on the left for comparison. The posterior pituitary is less cellular, and its pituicytes have ovoid to elongated nuclei and fibrillary cytoplasm (H&E, 20×)

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Fig. 23.11  Posterior lobe of the pituitary gland at 35 weeks gestation. Stain for glial fibrillary acidic protein (GFAP) shows only a few positive-­staining fibers (GFAP immunohistochemistry, 40×)

vary, with at least five different types described in the adult pituitary [8]. The second major constituent of the posterior lobe is the nonmyelinated axons of neurosecretory neuronal cell bodies originating in the supraoptic and paraventricular nuclei of the hypothalamus. These axons store the two secretory products of those nuclei, vasopressin or antidiuretic hormone and oxytocin. These axons are easily identifiable in silver-stained sections of adult glands, but they are not as easily identifiable, even with silver stains, in the fetal pituitary. The pituicytes surround the neurosecretory axons, and a major function of the posterior lobe is to release the stored hormones into the circulation. The posterior pituitary also has a rich vascular network. Salivary gland rests have been described in a small percentage of normal pituitary glands in children and adults; their presence may explain the occurrence of salivary gland neoplasms within the sella turcica [21]. The rests are typically found either on the surface of the neurohypophysis or within it. They are characterized by a cluster or clusters of small, rounded glandular structures lined by a simple cuboidal to columnar epithelium (Fig.  23.12). The nuclei of the epithelium are usually basally located, and the cytoplasm is granular and strongly PAS-positive [8]. Two possible mechanisms for the formation of these rests are the transfer of salivary glands from the primitive mouth during the formation of Rathke’s pouch or the induction of pituitary epithelium by heterotopic submandibular mesenchyme [21].

L. M. Ernst

Fig. 23.12  Salivary gland rests in the posterior lobe/neurohypophysis of the pituitary gland in a 1-year-old child. This image shows a small cluster of glandular structures lined by a simple columnar epithelium with basally placed nuclei and pale, eosinophilic granular cytoplasm. Note the fibrillary background of the neurohypophysis (H&E, 20×)

References 1. Ng PC. The fetal and neonatal hypothalamic-pituitary-adrenal axis. Arch Dis Child Fetal Neonatal Ed. 2000;82:F250–4. 2. Valdes-Dapena MA.  Histology of the fetus and newborn. Philadelphia: Saunders; 1979. 3. Sadler TW, Langman J.  Langman’s medical embryology. 11th ed. Philadelphia: Wolters Kluwer Lippincott Williams & Wilkins; 2010. 4. Schoenwolf GC, Larsen WJ. Larsen’s human embryology. 4th ed. Philadelphia: Elsevier/Churchill Livingstone; 2009. 5. Dasen JS, O’Connell SM, Flynn SE, Treier M, Gleiberman AS, Szeto DP, et al. Reciprocal interactions of Pit1 and GATA2 mediate signaling gradient-induced determination of pituitary cell types. Cell. 1999;97:587–98. 6. Nogami H, Hisano S.  Functional maturation of growth hormone cells in the anterior pituitary gland of the fetus. Growth Hormon IGF Res. 2008;18:379–88. 7. Treier M, Gleiberman AS, O'Connell SM, Szeto DP, McMahon JA, McMahon AP, Rosenfeld MG. Multistep signaling requirements for pituitary organogenesis in vivo. Genes Dev. 1998;12:1691–704. 8. Beatriz M, Lopes S, Pernicone PJ, Scheithauer BW, Horvath E, Kovacs K. Pituitary and sellar region. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 343–72. 9. Watanabe YG, Daikoku S.  An immunohistochemical study on the cytogenesis of adenohypophysial cells in fetal rats. Dev Biol. 1979;68:557–67. 10. Asa SL, Kovacs K, Horvath E, Losinski NE, Laszlo FA, Domokos I, Halliday WC.  Human fetal adenohypophysis. Electron microscopic and ultrastructural immunocytochemical analysis. Neuroendocrinology. 1988;48:423–31.

23  Pituitary Gland 11. Asa SL, Kovacs K, Laszlo FA, Domokos I, Ezrin C. Human fetal adenohypophysis. Histologic and immunocytochemical analysis. Neuroendocrinology. 1986;43:308–16. 12. Asa SL, Kovacs K, Singer W. Human fetal adenohypophysis: morphologic and functional analysis in  vitro. Neuroendocrinology. 1991;53:562–72. 13. Barson AJ, Symonds J. Calcified pituitary concretions in the newborn. Arch Dis Child. 1977;52:642–5. 14. Groisman GM, Kerner H, Polak-Charcon S. Calcified concretions in the anterior pituitary gland of the fetus and the newborn: a light and electron microscopic study. Hum Pathol. 1996;27:1139–43. 15. Naylor MF, Scheithauer BW, Forbes GS, Tomlinson FH, Young WF.  Rathke cleft cyst: CT, MR, and pathology of 23 cases. J Comput Assist Tomogr. 1995;19:853–9. 16. Zada G, Ditty B, McNatt SA, McComb JG, Krieger MD. Surgical treatment of Rathke cleft cysts in children. Neurosurgery. 2009;64:1132–7; author reply 1137–8.

247 17. Melchionna RH, Moore RA. The pharyngeal pituitary gland. Am J Pathol. 1938;14:763–72. 18. Kjaer I, Fischer-Hansen B.  Human fetal pituitary gland in holoprosencephaly and anencephaly. J Craniofac Genet Dev Biol. 1995;15:222–9. 19. Kjaer I, Fischer Hansen B, Reintoft I, Keeling JW. Pituitary gland and axial skeletal malformations in human fetuses with spina bifida. Eur J Pediatr Surg. 1999;9:354–8. 20. Hori A, Schmidt D, Rickels E.  Pharyngeal pituitary: develop ment, malformation, and tumorigenesis. Acta Neuropathol. 1999;98:262–72. 21. Hampton TA, Scheithauer BW, Rojiani AM, Kovacs K, Horvath E, Vogt P. Salivary gland-like tumors of the sellar region. Am J Surg Pathol. 1997;21:424–34.

Part VI Hematolymphoid System

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Thymus Gland Linda M. Ernst and Chrystalle Katte Carreon

Introduction The normal thymus gland is not frequently encountered in the examination of adult histology, because of its fatty involution after puberty. In the examination of fetal and neonatal tissues, however, the thymus gland is one of the major thoracic organs identified, so an understanding of its development and histological structure is important for anyone examining fetal and neonatal tissues. This chapter reviews the embryological origin and the major histological characteristics of the thymus gland throughout gestation, including the main components of the cortex and medulla and features of thymic involution.

Embryology The thymus gland is a lymphoepithelial organ composed of two lobes of approximately equal size. It is located beneath the sternum in the upper thorax, ventral to the great arteries, innominate vein, and the superior portion of the pericardial sac. The bulk of the organ is derived from the ventral buds of the bilateral third pharyngeal pouches, which arise at the end of the fourth week after conception. These buds are initially hollow diverticula, but they become solid cords of epithelial cells as their connection to the pharyngeal wall is lost by the seventh week. The cords migrate caudally and fuse to form the normally bilobed gland located inferior to the thyroid gland, with the bulk of the gland in the anterior mediastinum [1, 2]. In the fetus and newborn, it is usual for the gland

L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected] C. K. Carreon Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]

to include thin cylinders of tissue extending superiorly from the superior edge of one or both thymic lobes into the neck region. These cervical extensions of the thymus are normal, indicative of the trail of the embryologic descent of the thymus. Additionally, ectopic lobules of thymic tissue can be found anywhere along the descent of the thymic buds from the pharynx, as discussed further below. The endoderm of the pharyngeal pouch gives rise to the epithelial elements of the thymus gland, probably including Hassall’s corpuscles, and the organ is secondarily populated with bone marrow-derived precursor T lymphocytes shortly after formation [3]. The distinction between the cortex and medulla becomes visible in the embryologic period and is well established by the second trimester.

Histology General Overview The thymus is an encapsulated organ with a lobular architecture composed of a core of medullary tissue surrounded by cortical tissue (Fig.  24.1). The capsule comprises loose connective tissue with short trabeculae that extend into the organ and carry blood vessels and nerves. The cortex is the outer, dark-appearing region of the thymus, and the medulla is the lighter, central region. The main cell types within both the cortex and medulla of the thymus are epithelial cells and lymphocytes, also known as thymocytes, derived from the bone marrow. The epithelial-reticular cells form the reticular framework or cytoreticulum of the cortex and medulla; they secrete thymic hormones such as thymulin, thymosin, thymopoietin, and thymic humoral factor [3]. The lymphocytes are tightly packed in the interstices of the epithelial reticulum. Some epithelial cells, which are tightly joined together by desmosomes, form a continuous epithelial layer beneath the capsule, around the periphery of the entire gland, and invest the vascular trabeculae. This creates the blood–­thymus

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a

Fig. 24.1  General overview of thymic architecture. Low-power view of the thymus at term. Note the lobular architecture with outer, dark-­ appearing cortex and more central, light-appearing medulla (H&E, 1.25×)

b

barrier and isolates the thymus gland from much of the outside environment. This isolation is important for development of the lymphocytes.

Capsule and Connective Tissue Trabeculae The capsule consists of a thin layer of loose connective tissue containing blood vessels. Short connective-tissue trabeculae extend into the organ, carrying blood vessels. The capsule remains thin throughout gestation (Fig.  24.2). Immunohistochemical studies have demonstrated tyrosine hydroxylase-positive nerves in association with the large blood vessels beginning at 18  weeks gestation and have shown that these nerve fibers increase in density as gestation progresses [4]. The nerve branches penetrate the septa around 20  weeks gestation and reach the corticomedullary junction by 20–23 weeks. A few nerve fibers are also demonstrable within the medulla, closely associated with Hassall’s corpuscles.

c

Cortex The cortex is recognizable as the darker, more peripheral area of the gland, which is arranged in lobules separated by connective-tissue trabeculae (Fig.  24.3). The overall architecture and histological appearance of the thymic cortex changes over gestation (see Figs. 24.2 and 24.3). When the bone marrow-derived precursor lymphoid cells first appear in the cortex, the lobules are small and widely spaced by interlobular septa; the cortex is a small cap at the tips of the lobules. With increasing gestational age, the lobules become

Fig. 24.2  Thymus gland histology over gestation. Low-power view of the thymic development at (a) 17 weeks, (b) 22 weeks, and (c) 40 weeks. Note the thin capsule with trabeculae of connective tissue creating the lobulated appearance. The cellular, dark-staining cortex surrounds the lighter-staining medulla within the lobule. In the early midtrimester, the interlobular septa are wider, and the cortex is thinner. As gestation progresses, the interlobular septa become narrower, and the cortex becomes thicker (a–c H&E, 5×)

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larger, the interlobular septa become narrower, and the lobules appear more closely packed as the cortex extends along the edges of each lobule. By the beginning of the third trimester, the lobule appears to be composed of equal thirds: a

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one third cortex on one side, one third medulla, and one third cortex on the opposite side of the lobule. By term, it is expected that approximately 85% of the thymus is composed of cortex; the remainder is the medulla [5]. Dark staining of the cortex on hematoxylin and eosin (H&E) stain is due to densely packed lymphocytes. The density of the lymphocytes within the cortex obscures the underlying epithelial-reticular framework, so cortical epithelial cells are difficult to identify. They can be recognized by cytokeratin stains and electron microscopy as larger, ovoid to polygonal cells with elongated cytoplasmic processes that appear to encircle the lymphocytes (Figs.  24.4 and 24.5) [6]. Thymic epithelial cells can be

b

Fig. 24.4  Thymic epithelial cells at 22  weeks gestation. Cytokeratin stain highlights scattered cortical epithelial cells, some with cytoplasmic processes (cytokeratin AE1/AE3 immunohistochemistry, 20×)

c

Fig. 24.3  Thymic cortex over gestation. Medium-power views of the thymic capsule, cortex, and superficial medulla over gestation at (a) 17 weeks, (b) 22 weeks, and (c) term. Note the architecture of the cortex as it changes over gestation and becomes thicker and more densely packed with lymphocytes (a–c H&E, 20×)

Fig. 24.5  Thymic epithelial cells at term. Cytokeratin stain highlights scattered cortical epithelial cells, many with cytoplasmic processes, creating the reticular network of the thymus (cytokeratin AE1/AE3 immunohistochemistry, 20×)

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subtyped using antibodies to the thymic hormones. For example, subcapsular cortical cells are positive for thymosin α1 and thymopoietin, and inner cortical cells are negative [3]. Lymphocytes in the cortex consist almost exclusively of T lymphocytes (Figs.  24.6 and 24.7), which form a sheet without distinct follicle or germinal center formation. There is some variability in the cytologic features and immunophenotype of the lymphocytes, however. Recently arrived precursor T lymphocytes from the bone marrow populate a thin layer of the periphery of the cortex beneath the capsule. These immature thymocytes are large and actively proliferate. They undergo a series of maturational steps, and at each step they acquire distinctive immu-

nophenotypic cytoplasmic and membrane markers. As the lymphocytes mature, they become smaller (Fig.  24.8) and descend through the cortex and into the medulla. The fully differentiated thymocytes exit the medulla via the blood vessels and lymphatic channels. During this complex developmental process, some lymphocytes will undergo apoptosis. Macrophages are also present in the cortex and function as both antigen-presenting cells and phagocytic cells. After ingesting the debris of apoptotic lymphocytes, the macrophages are known as tingible body macrophages (Fig. 24.9). Extramedullary hematopoiesis is common and is usually confined to the capsule, interlobular septa, and along the vessels. It may be composed of purely eosinophilic myeloid elements (see Fig. 24.10).

Fig. 24.6  Thymic lymphocytes at term. CD3 stain highlights densely packed T lymphocytes in the cortex; in the medulla, they are still abundant but slightly less packed (CD3 immunohistochemistry, 20×)

Fig. 24.7  Thymic lymphocytes at term. CD20 stain shows a near absence of B lymphocytes in the cortex and scattered B lymphocytes in the medulla (CD20 immunohistochemistry, 10×)

Fig. 24.8  Thymic cortical lymphocytes. High-power view of thymic cortex at 23 weeks gestation. The lymphocytes are slightly larger near the periphery of the lobule and are smaller near the center (H&E, 40×)

Fig. 24.9  Tingible body macrophage in thymic cortex. Note the macrophage (arrow) with abundant cytoplasm containing apoptotic debris (H&E, 60×)

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Medulla In the medulla, the same lymphoid and epithelial components are present, but there are fewer lymphocytes, so the medulla has a “lighter” appearance by H&E stain. The reticular medullary epithelial cells are the predominant cells in the medulla, as they are no longer obscured by lymphocytes, and these cells are positive for thymosin α1 [3]. The epithelial cells of the medulla are larger than those in the cortex. In addition, ovoid epithelial structures with concentric lamellated keratinization, known as Hassall’s corpuscles, are characteristic of the medulla (Figs. 24.11, 24.12, 24.13, and 24.14). These structures originate from a single medullary epithelial cell that undergoes degeneration characterized by enlargement of the cell, eosino-

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philia, and vacuolation of the cytoplasm. Adjacent epithelial cells become involved, to form a lamellated, hyaline mass. Although the majority of the medullary lymphocytes are T lymphocytes, isolated B lymphocytes are seen along connective-­ tissue trabeculae, around the small vessels at the corticomedullary junction, and in the medulla around Hassall’s corpuscles.

Special Considerations Thymic Tissue in Ectopic Sites Thymic tissue can be found at any location along the embryologic path of descent of the thymus. It is normal and not

Fig. 24.10  Extramedullary hematopoiesis in thymus. High-power view of connective-tissue trabecula in thymic cortex with extramedullary hematopoiesis consisting mostly of perivascular eosinophilic myeloid precursors (H&E, 40×)

Fig. 24.11  Thymic medulla at 17 weeks gestation. Hassall’s corpuscles are visible as a few, small lamellated structures (arrow) (H&E, 20×)

Fig. 24.12  Thymic medulla at 22 weeks gestation. Hassall’s corpuscles are easily identified in the medulla and vary somewhat in size and shape (H&E, 20×)

Fig. 24.13  Thymic medulla at term. Numerous Hassall’s corpuscles are visible in the medulla (H&E, 20×)

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Fig. 24.14  Thymic medulla at term. Higher-power view of Hassall’s corpuscles (H&E, 40×)

L. M. Ernst and C. K. Carreon

Fig. 24.15  Thymic tissue in ectopic site. An ectopic thymic lobule is shown, located in the perithyroidal soft tissues near the lower pole of the thyroid gland, visible at the upper left (H&E, 5×)

 ge-Associated, Physiologic Involution A of the Thymus Gland

Fig. 24.16  Thymic tissue in ectopic site. Ectopic thymic tissue is shown within a partially autolyzed thyroid gland (H&E, 1.25×)

uncommon to find cervical extensions of thymic tissue emanating from the most superior pole of each lobe. It also is not unusual for the cervical extensions to reach the lower pole of the thyroid gland, but it is unusual for cervical extensions to reach the level of the isthmus or higher [7]. Nodules of thymic tissue not connected to the main gland are a common finding in fetuses. They can be found near the lower pole of the thyroid gland (Fig. 24.15), along the lateral lobes of the thyroid gland, adjacent to or admixed with parathyroid gland, within the thyroid gland (Fig. 24.16), along and behind the carotid sheath, behind the pharynx, and even as far superior as the base of the cranium.

After birth, the thymus continues to develop and organize its cortical and medullary compartments, while it is being periodically seeded with bone marrow-derived hematopoietic progenitor cells. An overall increase in organ size and production of naïve T lymphocytes to populate the peripheral environment is also observed during the early postnatal period [8, 9], and the organ has been observed to reach its maximum absolute weight during puberty [10, 11]. The thymus then goes through a slow and progressive process of involution with age, known as ageassociated or physiologic thymic involution, which is considered to be one of the hallmarks of the aging immune system. Physiologic thymic involution is characterized by gross regression in thymic size and microscopic alterations that eventually culminate in thymic atrophy (Figs. 24.17, 24.18, 24.19, 24.20, 24.21, and 24.22) [9–12]. This process was initially thought to commence at puberty [10, 11], shortly after the thymus has reached its maximal growth, but the exact timing of these events has not been fully characterized, and there is evidence of involutional events beginning even in early childhood [11, 13]. The microscopic features of physiologic thymic involution include gradual changes in the ratio of thymocytes to cortical and medullary epithelial cells. In the early stages, there can be a subtle to noticeable decrease in the density of cortical lymphocytes with relative sparing of the epithelial elements, which results in a higher density of lymphocytes in the medulla (“lymphocyte inversion”). Foci of adipose tissue may be seen in interlobular septa and within the parenchyma. In the more advanced stages, there is significant cortical lymphocyte depletion, with most of

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Fig. 24.17  Thymus at 2 weeks of age. A slightly prominent “starry-­ sky” pattern in the cortex may be seen around this age, which corresponds to the recently described “transient thymic involution” that can occur in the immediate postnatal period; this must be distinguished from stress-related involution (H&E, 4×)

Fig. 24.18  Thymic corticomedullary junction at 2 weeks of age. Note the many scattered tingible body macrophages in the cortex. Hassall’s corpuscles are present in the bottom left (H&E, 20×)

Fig. 24.19  Thymus at 4 years of age. There is prominent cortical lymphocyte dropout by this age, and additional early involutional changes characterized by the presence of adipose tissue within the septa and parenchyma (H&E, 4×)

Fig. 24.20  Thymus at 4 years of age. A closer view of the corticomedullary junction demonstrates prominent tingible body macrophages in the cortex, with mature adipocytes within the septa and parenchyma (H&E, 20×)

the thymic parenchyma being replaced by abundant adipose tissue; only scattered cords of epithelial cells and sparse lymphocytes remain [10, 11]. Partly cystic and closely aggregated Hassall’s corpuscles have also been described during the later stage of involution [10]. Theoretically, the process of physiologic thymic involution can reach a point where no thymic tissue can be identified grossly, but microscopic thymic remnants are probably always present, and sectioning the pre-epicardial soft tissue may increase the likelihood of finding these elusive foci [10]. One group observed that some of the abovementioned histologic features of involution are present as early as the

first month of life and in a few subjects less than 5 years of age. Moreover, the group noticed that the changes became more evident beyond 12  years of age, which marks the beginning of thymic parenchyma replacement by adipose tissue [11]. Interestingly, another group found that a “transient thymic involution” occurs in the neonatal thymus, which was described as a temporary severe cortical depletion of immature double-positive (both CD4-positive and CD8-positive) thymocytes (Fig.  24.17), which is largely induced by an increased rate of apoptosis during this period. This t­ emporary reduction in thymocytes could be observed as early as 1 day

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Fig. 24.21  Thymus at 17 years of age. In contrast to the involutional changes noted in early childhood, this thymus exhibits a more significant amount of fat, which comprises at least 60% of the total parenchyma. The thymic lobules are represented by scattered small, irregular islands of lymphocytes and epithelial cells (H&E, 2×)

Fig. 24.22  Thymus at 17 years of age. The remnants of thymic lobules show extensive lymphocyte depletion and are composed of an admixture of lymphocytes and epithelial cells but without distinct cortical and medullary regions. There is abundant intervening mature fat separating the lobules. A Hassall’s corpuscle is noted (arrow) (H&E, 20×)

Fig. 24.23  Stress involution of the thymus, mild. Thymus gland at 32 weeks gestation is shown with starry-sky appearance of the thymic cortex due to depletion of cortical lymphoid cells (H&E, 10×)

Fig. 24.24  Stress involution of the thymus, moderate. Thymus gland at 30  weeks gestation is shown with more severe cortical lymphoid depletion, cortical thinning, and blurring of the distinction between cortex and medulla. Also note that Hassall’s corpuscles appear more crowded (H&E, 10×)

after birth, with recovery of the normal proportion of the thymocyte subset toward the end of the first month of postnatal life. This occurrence is thought to be a physiologic event, not a form of stress response [11, 14].

 tress-Associated Involution of the Thymus S Gland The histologic changes of thymic involution described above, excluding the adipose tissue replacement, are sometimes seen in the fetal thymus gland and can be a useful marker of stress in utero. A stressful in utero environment, such as those caused by congenital infections, malnutrition, or severe placental pathology, can lead to the production of glucocorti-

coids that have lytic effects on the cortical thymocytes [15]. Therefore, thymic stress involution, also known as acute thymic involution, is characterized by lysis or apoptosis of the many dark-staining cortical lymphocytes (Figs. 24.23, 24.24, and 24.25). The process is progressive and produces increasingly severe histologic abnormalities over time: (1) a starrysky appearance of the cortex due to unmasking of tingible body macrophages and epithelial cells, (2) a less basophilic appearance of the cortex, (3) narrowing of the cortex, (4) blurring of the corticomedullary distinction, (5) disappearance of the cortex, (6) crowding of Hassall’s corpuscles in the medulla, and (7) reduction in thymic weight [5, 16]. Recent evidence suggests that the changes of acute thymic involu-

24  Thymus Gland

Fig. 24.25  Stress involution of the thymus, severe. Thymus gland from a neonate born at 33 weeks gestation who survived for 1 week and ultimately died of sepsis. There is severe, nearly complete cortical lymphoid depletion with marked reduction of the distinction between cortex and medulla and more prominent crowding of Hassall’s corpuscles (H&E, 10×)

tion may be therapeutically halted and/or reversed [17]. As discussed in Chap. 37, caution should be used when interpreting changes of stress involution in the setting of intrauterine fetal death, as loss of nuclear basophilia in all organs is characteristic of fetuses retained in utero after death.

References 1. Valdes-Dapena MA. Thymus. In: Histology of the fetus and newborn. Philadelphia: Saunders; 1979. p. 45–58. 2. Sadler TW, Langman J.  Head and neck. In: Langman’s medical embryology. 11th ed. Philadelphia: Wolters Kluwer Lippincott Williams & Wilkins; 2010. p. 272.

259 3. Pearse G. Normal structure, function, and histology of the thymus. Toxicol Pathol. 2006;34:504–14. 4. Anagnostou VK, Doussis-Anagnostopoulou I, Tiniakos DG, Karandrea D, Agapitos E, Karakitsos P, Kittas C.  Ontogeny of intrinsic innervation in the human thymus and spleen. J Histochem Cytochem. 2007;55:813–20. 5. Singer D. Disorders of the lymphoid tissue and immune system. In: Wigglesworth J, Singer D, editors. Textbook of fetal and neonatal pathology. 2nd ed. Malden: Blackwell Science; 1998. p. 436–7. 6. Suster S, Rosai R.  Thymus. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 541–62. 7. Gorlin R, Cohen M.  The orofacial region. In: Wigglesworth J, Singer D, editors. Textbook of fetal and neonatal pathology. 2nd ed. Malden: Blackwell Science; 1998. p. 743–4. 8. Chinn IK, Blackburn CC, Manley NR, Sempowski GD.  Changes in primary lymphoid organs with aging. Semin Immunol. 2012;24:309–20. 9. Rezzani R, Nardo L, Favero G, Peroni M, Rodella LF. Thymus and aging: morphological, radiological, and functional overview. Age. 2014;36:313–51. 10. Suster S, Rosai J.  Histology of the normal thymus. Am J Surg Pathol. 1990;14:284–303. 11. Raica M, Cîmpean AM, Encică S, Cornea R. Involution of the thymus: a possible diagnostic pitfall. Romanian J Morphol Embryol. 2007;48:101–6. 12. Gui J, Mustachio LM, Su D-M, Craig RW. Thymus size and age-­ related thymic involution: early programming, sexual dimorphism, progenitors and stroma. Aging Dis. 2012;3:280–90. 13. Hale LP. Histologic and molecular assessment of human thymus. Ann Diagn Pathol. 2004;8:50–60. 14. Varas A, Jiménez E, Sacedón R, Rodríguez-Mahou M, Maroto E, Zapata AG, Vicente A.  Analysis of the human neonatal thymus: evidence for a transient thymic involution. J Immunol. 2000;164:6260–7. 15. Glavina-Durdov M, Springer O, Capkun V, Saratlija-Novaković Z, Rozić D, Barle M.  The grade of acute thymus involution in neonates correlates with the duration of acute illness and with the percentage of lymphocytes in peripheral blood smear. Pathol Stud Biol Neonate. 2003;83:229–34. 16. van Baarlen J, Schuurman HJ, Huber J. Acute thymus involution in infancy and childhood: a reliable marker for duration of acute illness. Hum Pathol. 1988;19:1155–60. 17. Ansari AR, Liu H.  Acute thymic involution and mechanisms for recovery. Arch Immunol Ther Exp. 2017;65:401–20.

25

Spleen Linda M. Ernst

Introduction The histological appearance of the spleen changes dramatically during fetal life. Much of the architecture of the spleen is not fully formed in the embryonic period, so significant development of the red and white pulp occurs during the second and third trimesters of fetal life, with additional changes occurring postnatally. Unfortunately, there are only a few histological studies of the human fetal spleen in the literature, and although they provide some insight into prenatal splenic development, more comprehensive analyses are needed to fully understand the changes occurring in this complex organ during fetal life. Nevertheless, a working knowledge of the normal splenic appearance at certain gestational ages can be helpful in recognizing abnormal development of the spleen that has been retarded or accelerated by a pathologic process. This chapter provides representations of the normal splenic architecture throughout the second and third trimesters.

Embryology The spleen is derived from condensation of mesoderm within the dorsal mesogastrium around the fifth week of gestation. This mesoderm forms the capsule, connective tissue, and reticular framework of the spleen. The splenic mass takes its final position in the left upper quadrant of the peritoneal cavity after growth of the dorsal mesogastrium and rotation of the stomach. This rotation also establishes the splenorenal (lienorenal) ligament. The gastrosplenic (gastrolienal) ligament is formed by the portion of the dorsal mesentery that remains between the spleen and stomach [1, 2].

L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected]

At the end of the embryonic period, although the splenic anlage is formed and has assumed its appropriate anatomic position, splenic development is quite primitive. There is no distinct architectural organization at this time, and the parenchyma of the splenic anlage consists predominantly of hematopoietic precursors and a few macrophages. Almost no lymphocytes are present at this stage [1, 2]. The following sections review the histologic changes in the capsule, the vascular tree, and the white and red pulp in the fetal and neonatal period.

Histology Capsule The fetal spleen is an encapsulated organ, and the capsule is composed of longitudinally arranged collagen and elastic fibers. In early gestation, the capsule is a thin layer, but with increasing gestational age the capsule becomes slightly thicker. Trabeculae of connective tissue extend downward from the capsule, but they generally do not contain blood vessels and extend only a short distance into the parenchyma.

Vascular Tree The splenic artery enters the spleen at the hilum and gives rise to branches that become increasingly smaller, with a decreasing collagenous cuff also containing veins and lymphatic channels. In the mature spleen, the central arteriole is identified because it no longer travels with an accompanying vein, and its collagenous adventitia is replaced by lymphoid tissue [3, 4]. Prior to significant lymphoid colonization of the fetal spleen, these central arterioles or their precursors can be seen as small, muscularized arterioles surrounded by delicate reticulin fibers (Fig. 25.1). In the mature spleen, the central arterioles terminate in a capillary bed formed within

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L. M. Ernst Table 25.1  Summary of splenic lymphoid and sinus development: information regarding lymphocytes and splenic sinuses at various gestational ages Gestational age 0–12 weeks 14–18 weeks 19–23 weeks 23+ weeks

After birth

Fig. 25.1  Fetal spleen at 16 weeks gestation. Note the early reticulin condensation around the small arteriole (reticulin stain, 40×)

Lymphocytes None Scattered B and T lymphocytes

Splenic sinuses Not formed Not formed

Lymphocyte condensations begin around arterioles Increasing number of B lymphocytes around arterioles Primary follicle formation begins Formation of germinal centers with antigen exposure Formation of mantle and marginal zones

Beginning formation Present

Present

a

the lymphoid tissue of the white pulp. At the termination of these capillaries, the lining cells are replaced by macrophages, the so-called “sheathed” end of the capillary; blood flowing through this capillary then passes into the splenic sinuses [3, 4]. The splenic sinuses eventually merge with the veins that are traveling along with the larger arteries to the hilum.

White Pulp The white pulp of the spleen is essentially the lymphoid component of the organ, which in the adult spleen contains mature T-lymphocyte and B-lymphocyte compartments. The B-cell areas include the germinal center, mantle zone, and marginal zone; the T-cell area is the less well-organized parafollicular zone, but T cells are also a component of the periarteriolar clusters. In the fetal spleen, these lymphoid compartments are just beginning to develop and do not achieve the histological architecture that is recognizable in the mature, adult spleen, even by the end of full-term gestation. Table 25.1 summarizes the time-dependent changes in the splenic lymphoid development and the development of the splenic sinuses. Colonization of the spleen by lymphoid tissue is usually not visible until approximately 14–18 weeks gestation (Figs. 25.2, 25.3, and 25.4); at first, it consists of scattered T lymphocytes and B lymphocytes [5, 6]. At about 19–20 weeks gestation, lymphocytes begin to form small clusters around splenic arterioles (Fig. 25.5). In this early stage, there is no specific organization of lymphocytes into lymphoid follicles, and a mixture of B lymphocytes and T lymphocytes is present within the lymphoid aggregates (Fig.  25.6). It has been shown that the early aggregates

b

Fig. 25.2  Fetal spleen at 16 weeks gestation. Note the absence of conspicuous lymphoid aggregates at this early gestational age. (a) Very little lymphoid tissue appears to be present on H&E stain. Note the thin, fibrous capsule. (b) Reticulin stain highlights reticulin within the fibrous capsule and the very short trabeculae extending into the parenchyma. Also note that there is very little reticulin tissue within the splenic parenchyma at this gestational age, with only small concentric condensations of reticulin around arterioles (a H&E, 10×; b reticulin stain, 10×)

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a

b

Fig. 25.4  Fetal spleen at 16 weeks gestation. This higher-power view shows the CD20+ cells aggregating around the central arteriole (CD20 immunohistochemistry, 20×)

a

Fig. 25.3  Fetal spleen at 16 weeks gestation. These images show the complement of CD3+ T lymphocytes (a) and CD20+ B lymphocytes (b). Note that the B lymphocytes are beginning to form small aggregates, and scattered T lymphocytes are also present (a CD3 immunohistochemistry, 10×; b CD20 immunohistochemistry, 10×)

of B lymphocytes accumulate around the more peripheral branches of the arterioles, whereas T lymphocytes are seen more centrally surrounding the larger arteriole trunks [6]. By the end of the second trimester, B lymphocytes begin to form small, primary follicular clusters (Fig.  25.7), including the presence of follicular dendritic cells within the follicle [6]. These primary follicles can be recognized on hematoxylin and eosin (H&E) stain by the presence of an eccentric proliferation of small lymphocytes adjacent to the periarteriolar lymphoid cuff (Figs.  25.8 and 25.9). Steiniger et al. [6] have also demonstrated that these first fetal B-cell follicles do not arise from the initial B-cell clusters along the peripheral arterioles but that they form more centrally, where the periarteriolar T-cell sheaths are present. These primary follicles enlarge throughout the third trimester (Figs.  25.10, 25.11, 25.12, 25.13, 25.14, and 25.15) [7].

b

Fig. 25.5  Fetal spleen at 21 weeks gestation. Early lymphoid aggregates are beginning to become visible on H&E stain (a). Note that the capsule is slightly thicker and that reticulin stain highlights lengthening trabeculae extending into the parenchyma (b) (a H&E, 10×; b reticulin stain, 10×)

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a

b b

Fig. 25.6  Fetal spleen at 21 weeks gestation. These images show the complement of CD3+ T lymphocytes (a) and CD20+ B lymphocytes (b). Note that the B-lymphocyte aggregates are larger and more well-formed than at 16 weeks gestation. Also, slightly more scattered T lymphocytes are present (a CD3 immunohistochemistry, 10×; b CD20 immunohistochemistry, 10×)

Germinal centers are not generally seen in the fetal spleen and may even be absent in the early neonatal period until sufficient antigen exposure and time are allowed for germinal center formation. Therefore, in summary, by the end of normal term gestation (Figs. 25.16 and 25.17), splenic lymphoid tissue consists of primary follicles (B-lymphocyte zones) and the early formation of periarteriolar lymphoid sheaths (T-lymphocyte zones), but the reactive secondary follicles and the formation of splenic mantle and marginal zones are absent. After birth, there is a continued lag time before the secondary follicles, mantle zone, and marginal zone lymphocytes are seen in the spleen. Secondary follicles form in the neonatal period, after antigen exposure, and contain a mixture of small B lymphocytes and large, transformed B lymphocytes producing a characteristically paler appearance than the primary follicles on H&E stain (Fig. 25.18). In addition, follicular dendritic cells (CD21+, CD23+, and

Fig. 25.7  Fetal spleen at 23 weeks gestation. On the H&E-stained section (a), the lymphoid tissue is becoming more prominent. The reticulin stain (b) highlights the slightly increased thickness of the capsule and the increasing number and thickening of reticulin fibers extending into the parenchyma (a H&E, 10×; b reticulin stain, 10×)

Fig. 25.8  Fetal spleen at 24 weeks gestation. This higher-power view shows one of the lymphoid aggregates (H&E, 20×)

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a

b

Fig. 25.10  Fetal spleen at 32 weeks gestation. This low-power view shows the presence of clearly recognizable lymphoid aggregates at this gestational age (H&E, 4×)

a

Fig. 25.9  Fetal spleen at 24 weeks gestation. These images show the complement of CD3+ T lymphocytes (a) and CD20+ B lymphocytes (b). Note that the B-lymphocyte aggregates are well-formed and are associated with scattered T lymphocytes, which also begin to form small clusters (a CD3 immunohistochemistry, 10×; b CD20 immunohistochemistry, 10×)

Fig. 25.11  Fetal spleen at 32 weeks gestation. Note the eccentric proliferation of lymphocytes (primary follicle) adjacent to a central arteriole (H&E, 20×)

b

Fig. 25.12  Fetal spleen at 32 weeks gestation. These images show the complement of CD3+ T lymphocytes (a) and CD20+ B lymphocytes (b). Note that B-lymphocyte aggregates now are seen adjacent to clusters of periarteriolar T lymphocytes (a CD3 immunohistochemistry, 4×; b CD20 immunohistochemistry, 4×)

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a

Fig. 25.13  Fetal spleen at 35 weeks gestation. This low-power view shows numerous primary lymphoid follicles without germinal centers (H&E, 4×)

CD35+), occasional CD4+ T cells, and macrophages are also seen within the secondary follicle [3]. In an infant autopsy series examining splenic lymphoid development, it has been shown that secondary follicles are consistently found at an age of 2.5  months or older [7]. The secondary follicle is surrounded by a more compact zone of small lymphocytes known as the mantle zone (bcl-2+, CD45RA+, IgM+, and IgD+), and the marginal zone surrounds the mantle zone [3]. The splenic marginal zone is composed of medium-sized B lymphocytes (bcl-2+, CD45RA−, and IgD−) in a dense, concentric meshwork of reticulin fibers that express smooth muscle actin [3]. The splenic marginal zone has been shown to be consistently present at an age of 8.5 months or older [7]. In addition, the lack of appropriate and timely mantle zone development has been shown to be associated with sudden infant death or infectious causes of death [7]. As the splenic B-lymphocyte regions mature, the T-cell areas become more well defined and form a nodule of CD4+ T lymphocytes around the splenic arterioles. These are usually seen at the edge of the secondary follicles and are surrounded by concentric reticulin fibers that are positive for smooth muscle actin (SMA).

Red Pulp The red pulp, the tissue intervening between the white pulp, is composed of a loose, reticular stromal tissue, also known as the cords of Billroth, which contains lymphocytes, macrophages, and other blood cells. Reticulin stains of the fetal spleen show increasing amounts of reticulin within the red pulp as the spleen matures with advancing gestation (see Figs. 25.2, 25.5, 25.7, 25.14, and 25.17). Between the splenic

b

Fig. 25.14  Fetal spleen at 35 weeks gestation. (a) A slightly higher-­ power view of the splenic capsule and a few of the underlying lymphoid follicles with H&E stain. (b) The reticulin stain highlights the capsular reticulin and increasing intraparenchymal reticulin (a H&E, 10×; b reticulin stain, 10×)

cords are the specialized sinuses of the spleen, which form as outpouchings of the peripheral venules. The sinusoids have fenestrated basement membranes and are surrounded by transversely arranged reticulin fibers. Steiniger et  al. [6] have suggested that even in the early second trimester, splenic sinuses are not yet formed, and they begin formation at 19–23 weeks gestation (see Table 25.1). The endothelium that lines the fetal splenic sinuses is recognized as weakly expressing CD34 antigen [6].

Special Considerations Accessory Spleen(s) Accessory spleens usually develop from smaller adjacent condensations of mesenchyme near the splenic hilum. Their structure and histology are typically similar to those of the main spleen.

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Fig. 25.15  Fetal spleen at 35 weeks gestation. A higher-power view of a primary lymphoid follicle is shown (H&E, 20×)

Fig. 25.16  Fetal spleen at term, showing a primary lymphoid follicle. Germinal centers are not seen (H&E, 20×)

Fig. 25.17  Fetal spleen at term, showing the mature fetal reticulin pattern (reticulin stain, 10×)

Fig. 25.18  Postnatal spleen showing focal germinal center (secondary follicle) formation (H&E, 20×)

Fetal Spleen and Hematopoiesis

poietic growth factors, such as granulocyte colony-stimulating factor and erythropoietin. The role of the fetal spleen in prenatal hematopoiesis is still a controversial topic, and designation of the spleen as a primary or secondary hematopoietic organ will require further debate and investigation.

The fetal spleen has classically been considered a hematopoietic organ, but the exact role of the fetal spleen in hematopoiesis remains controversial. It is clear that hematopoietic cells are seen in the spleen beginning in the embryonic period and throughout fetal life, but with far less density than in the fetal liver. In a study employing cell suspensions of human midtrimester fetal spleen, Calhoun et al. [8] have shown that the hematopoietic cells present are predominantly polychromatic and orthochromatic normoblasts and that the most immature myeloid and erythroid precursors are rare to absent. They note that splenic normoblasts are seen predominantly in vessels and sinuses of the spleen in histological sections. In addition, these authors showed that the midtrimester fetal spleen lacks transcripts of the typical hemato-

References 1. Sadler TW, Langman J. Langman’s medical embryology. 11th ed. Philadelphia: Wolters Kluwer Lippincott Williams & Wilkins; 2010. 2. Schoenwolf GC, Larsen WJ. Larsen’s human embryology. 4th ed. Philadelphia: Elsevier/Churchill Livingstone; 2009. 3. O’Malley DP, George TI, Orazi A, Abbondanzo SL.  Embryology and histology of the lymph node and spleen. In: Benign and reactive conditions of lymph node and spleen. Washington, DC: American Registry of Pathology; 2009.

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4. van Krieken JHJ, Oraxi A. Spleen. In: Mills SE, editor. Histology 7. Kruschinski C, Zidan M, Debertin AS, von Hörsten S, Pabst R. Age-­ dependent development of the splenic marginal zone in human for pathologists. Philadelphia: Lippincott Williams & Wilkins; infants is associated with different causes of death. Hum Pathol. 2007. 2004;35:113–21. 5. Timens W, Rozeboom T, Poppema S. Fetal and neonatal development of human spleen: an immunohistological study. Immunology. 8. Calhoun DA, Li Y, Braylan RC, Christensen RD.  Assessment of the contribution of the spleen to granulocytopoiesis and erythropoi1987;60:603–9. esis of the mid-gestation human fetus. Early Hum Dev. 1996;46: 6. Steiniger B, Ulfig N, Risse M, Barth PJ.  Fetal and early post-­ 217–27. natal development of the human spleen: from primordial arterial B cell lobules to a non-segmented organ. Histochem Cell Biol. 2007;128:205–15.

26

Lymph Nodes Michele E. Paessler

Introduction The function of the lymph nodes is to interact with and process antigens. Lymph nodes are found throughout the body and are usually located along the blood vessels. Lymph nodes are numerous in areas that drain organs and in areas that are in contact with the environment [1]. In particular, large groups of lymph nodes are found in the respiratory (hilar and mediastinal lymph nodes) and gastrointestinal (mesenteric lymph nodes) tracts. This chapter focuses on the architecture of the developing lymph node throughout the second and third trimesters.

Embryology The lymphatic vessels and lymph nodes arise from the lateral plate mesoderm. To understand the development of the lymph nodes, one must first understand the lymphatics. The lymphatic system develops as discrete spaces in the mesenchyme, which form an endothelial lining. These spaces fuse, progressively grow, and branch into the lymphatic system [2]. The lymphatic system originally is distributed along the main venous trunks. The fusion and integration of these lymphatic networks give rise to six lymph sacs [3]. The first lymph sacs appear at about the fifth week of gestation as the right and left jugular lymph sacs. Subsequently, the subclavian duct, thoracic duct, cisterna chyli, and retroperitoneal lymph sacs develop [4]. It is from these primary sacs that endothelial sprouts grow, hollow out to form a lumen, and ultimately give rise to the peripheral lymphatic vessels [4].

M. E. Paessler (*) Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]

Once the lymphatic system is established, primary lymph nodes start to develop at about 8–11  weeks gestation. The development of a lymph node starts with a lymphatic plexus in close association with strands of mesenchymal tissue. The mesenchymal cells condense at the base of the lymph sac and invaginate inward along with the endothelium [5]. In addition to the mesenchyme, the lymph sac is composed of capillaries, vascular loops, fibroblastic reticulum cells, and extracellular matrix [6, 7]. By 13 weeks gestation, the lymph sac is composed predominantly of mesenchyme, and the space between the endothelial lining and mesenchymal core develops into the marginal sinus [5]. At this point, the capillaries increase, and the intermediate sinuses develop. The connective tissue on the outer side of the marginal sinus gives rise to the lymph node capsule [6, 7]. The marginal sinus is first colonized by lymphoblasts and later repopulated by lymphocytes. A mass of smaller channels is formed by the growing lymphocytes and connective tissue; these small channels give rise to the efferent vessels. Blood vessels follow along the connective tissue framework in which the lymphocytes are proliferating. Dense lymphoid tissue continues to develop in tandem with the vascular strands, and this gives rise to the characteristic morphology seen in the medulla of the lymph node [4]. The cortex of the lymph node does not develop until much later in gestation, and mature, complete lymph nodes with germinal centers are not seen until the postnatal period [2].

Histology In the developing fetus, the lymph nodes are basically a mass of lymphoid tissue that has a capsule and sinuses. In the early weeks of gestation, the lymph nodes are small and appear to be depleted. As the gestational age increases, the lymph nodes become larger and contain more lymphocytes.

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Blood Supply Arteries enter the lymph node at the hilus and then branch and form a capillary network that supplies the paracortex and follicles. Venous drainage accompanies the arteriolar route, and the veins leave the lymph node through the hilus. The postcapillary venules are the entry site of circulating lymphocytes into the lymph node parenchyma [1].

Lymphatics Lymphatic fluid enters the lymph node through the afferent lymphatic vessel, which penetrates the capsule and then enters the subcapsular, intermediate, and medullary sinuses. Lymphatic fluid then exits the lymph node through the efferent lymphatic vessel. The intranodal sinuses are lined by sinus endothelial cells. Naïve B cells and T cells from the blood use specific receptors to recognize ligands on the high endothelial venules in the lymph nodes; they pass between these cells to enter the paracortex and then migrate to distinct B-cell and T-cell areas within the lymph node [8]. Under the influence of cytokines and chemokines, the T cells localize in the paracortex and associate with dendritic cells, which aid in their differentiation. B cells localize to follicles under the influence of cytokines secreted from follicular dendritic cells.

Lymph Nodes On gross examination, fetal lymph nodes are typically small and round- to bean-shaped. On sectioning, the lymph nodes are tan-gray in appearance. Lymph nodes (especially the mesenteric lymph nodes) can be identified grossly by the middle of the second trimester. Microscopic sections of the fetal lymph node show that the lymphoid compartments are just beginning to develop and do not achieve the histologic architecture that is recognizable in the mature, adult lymph node, even at the end of full-term gestation. The demarcation of cortex and medulla becomes more distinct with the increase in gestational weeks [5]. Early in the second trimester, lymph nodes are usually quite small, are surrounded by a capsule, and contain sinuses. Light and electron microscopy has shown that the marginal, intermediate, and medullary sinuses become developed with

Fig. 26.1  Lymph node at 19 weeks gestation. This lymph node, one of many small lymph nodes in the mesentery, is surrounded by a collagenous connective tissue capsule. Beneath the capsule, the sinus is apparent. The lymph node has a diffuse pattern and is composed of small lymphocytes. No identifiable follicles are seen (hematoxylin and eosin (H&E), 10×)

increasing gestational age [7]. By light microscopy, fetal lymph nodes have a diffuse pattern and are composed of small lymphocytes. There is often a paucity of lymphocytes when compared with adult nodes, imparting a “depleted” appearance (Fig. 26.1), but this should not be misinterpreted as a pathologic change. From the 12th week of gestation until term, T cells circulate in the periphery and settle into the growing lymph nodes [5, 9]. This influx of lymphocytes from the circulation leads to enlargement of the lymph nodes. Early in the second trimester, the fetal lymph nodes are composed predominantly of T cells that are scattered throughout the primitive lymph node (Fig. 26.2a); there is a predominance of CD4+ T cells and only rare CD8+ T cells [10]. At this point, B cells are present in the outer cortex but do not yet form primary follicles [10]. After the 17th week of gestation, clearly defined B-cell follicles can be found. Although the primary follicles are formed, they may be difficult to see on morphologic grounds alone. Immunohistochemical stains for B-cell markers highlight these small primary follicles (Fig. 26.2b). As the fetus matures and approaches term, the lymph nodes enlarge, generally become more cellular, and appear less depleted (Figs. 26.3, 26.4, 26.5, and 26.6). The demar-

26  Lymph Nodes

a

271

b

Fig. 26.2  Lymph node at 19 weeks gestation. (a) CD3 immunohistochemical stain shows that the lymph node is composed predominantly of T cells. Primary follicles cannot be recognized by H&E but become evident with immunohistochemistry. The follicles on the periphery of

the lymph node do not stain with CD3. (b) CD20 immunohistochemical stain highlights B cells in the primary follicles in the cortex. (a CD3 immunohistochemistry; b CD20 immunohistochemistry; 2.5×)

Fig. 26.3  Mesenteric lymph nodes at 34 weeks gestation. The developing lymph nodes show a capsule and a network of sinuses. No follicles are seen (H&E, 2.5×)

Fig. 26.4  Mesenteric lymph node at 34 weeks gestation. This higher-­ power view shows sinuses (H&E, 5×)

272

Fig. 26.5  Lymph node at 41 weeks gestation. The lymph node is larger and more cellular than those seen earlier in gestation; it has a capsule and sinuses. Primary follicles are not readily seen (H&E, 2.5×)

M. E. Paessler

Fig. 26.6  Lymph node at 41 weeks gestation. This higher-power view shows that the lymph node is composed predominantly of sheets of small lymphocytes (H&E, 10×)

regions in the cortex, where they are accompanied by interdigitating dendritic cells. Germinal centers are not apparent in any stage of gestation, even in the term fetus [7, 9]. Normally, germinal center formation does not occur until the postnatal period, when their formation is driven by antigenic stimuli [11].

Special Considerations Lymph Node and Hematopoiesis

Fig. 26.7  Lymph node at 41 weeks gestation. CD3 immunohistochemical stain shows that the lymph node is composed predominantly of CD3-positive T cells (CD3 immunohistochemistry, 10×)

cation of cortex and medulla becomes more apparent with increasing gestational age. The cortex is more cellular than the medulla, which has a much looser structure [5]. Both light microscopy and electron microscopy show that the marginal, intermediate, and medullary sinuses also become more developed [7]. T cells are still a major component of the lymph node (Figs. 26.7 and 26.8) and localize to T-cell

The lymph node may be a site of hematopoiesis in the fetus. Cells from the erythropoietic and granulocytic lineages may be seen. If present, they are usually located in the medulla of the lymph node.

Hemophagocytosis Hemophagocytosis is commonly seen in lymph nodes from fetal autopsies. The presence of hemophagocytosis does not imply that the patient has hemophagocytic syndrome or hemophagocytic lymphohistiocytosis. The significance of this finding is not known; it may likely represent an agonal event. A clear etiology or source of infection inciting this finding is not known.

26  Lymph Nodes

a

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b

Fig. 26.8  Lymph node at 41 weeks gestation. (a) In this higher-power view, CD3 immunohistochemical stain shows positive T cells and small nodules of negative cells that represent B-cell follicles. (b) CD20

immunohistochemical stain shows primary follicles composed of small B lymphocytes. (a CD3 immunohistochemistry; b CD20 immunohistochemistry; 20×)

References

7. Markgraf R, Gauecker B, Muller-Hermelink HK.  The development of the human lymph node. Cell Tissue Res. 1982;225: 387–413. 8. Crivellato E, Vacca A, Ribatti D.  Setting the stage: an anatomist’s view of the immune system. Trends Immunol. 2004;25: 210–7. 9. Kyriazis AA, Esterly JR.  Development of lymphoid tissues in the human embryo and early fetus. Arch Pathol. 1970;90: 348–53. 10. Asano S, Akaike Y, Muramatsu T, Mochizuki M, Tsuda T, Wakasa H.  Immunohistologic detection of the primary follicle (PF) in human fetal and newborn lymph node anlages. Pathol Res Pract. 1993;189:921–7. 11. Veereman-Wauters G.  Neonatal gut development and postnatal adaptation. Eur J Pediatr. 1996;155:627–32.

1. Van der Valk P, Meijer CJ.  Lymph nodes. In: Mills SE, editor. Histology for pathologists. 3rd ed. Philadelphia: Lippincott Williams & Wilkins; 2007. p. 763–81. 2. Arey LB.  Developmental anatomy. 7th ed. Philadelphia: W.B. Saunders; 1974. p. 370–2. 3. Valdes-Dapena MA.  Histology of the fetus and newborn. Philadelphia: W.B. Saunders; 1979. p. 65–77. 4. Patten BM. Human embryology. 2nd ed. New York: McGraw-Hill; 1953. p. 649–55. 5. Hoorweg K, Cupedo T. Development of human lymph nodes and Peyer’s patches. Semin Immunol. 2008;20:164–70. 6. Bailey RP, Weiss L. Light and electron microscopic studies of post capillary venules in developing human fetal lymph nodes. Am J Anat. 1975;143:43–58.

27

Palatine Tonsil Linda M. Ernst and Chrystalle Katte Carreon

Introduction

ond pharyngeal arches. At approximately 12-week gestation, the epithelium of the second pharyngeal pouch proliferThe palatine tonsils, representing a portion of the gut-­ ates and grows in a finger-like fashion into the surrounding associated lymphoid tissue, are usually the largest and most mesenchyme. The central cells in these epithelial columns readily sampled tonsils in the human fetus. The palatine degenerate, forming the tonsillar crypts that are secondarily tonsils are located along the lateral walls of the oropharynx, infiltrated by lymphoid tissue. The mesenchyme forms the within the tonsillar fossa created by two tonsillar pillars. remaining stromal elements of the tonsils [3, 4]. The anterior tonsillar pillar is also known as the palatoglossal arch and forms an arc of soft tissue and muscle between the tongue and the soft palate. The posterior tonsillar pillar, Histology also known as the palatopharyngeal arch, forms an arc of soft tissue and muscle between the soft palate and the posterior In the early second-trimester fetus, the palatine tonsillar tispharynx [1, 2]. Because there is little projection of the pala- sue is recognized as infoldings of stratified squamous epithetine tonsils into the pharynx during fetal life, they are easily lium in close continuity with small lymphoid tissue congeries overlooked grossly, but they can be sampled for microscopic (Figs.  27.1, 27.2, and 27.3). Investigators have shown that study by carefully removing the soft palate and tonsillar fos- as early as 14-week gestation, B-lymphoid cells are seen in sae during removal of the tongue. Serial horizontal sections association with follicular dendritic cells, representing the of the neck tissues, including the attached soft palate, will early formation of lymphoid follicles. In addition, parafollicusually provide reasonable microscopic sections of the palatine tonsils. This chapter reviews the histological changes in the palatine tonsil during fetal life.

Embryology The epithelium of the palatine tonsils is derived from the bilateral endodermal buds of the second pharyngeal pouch. The mesodermal elements are derived from the second pharyngeal membrane and nearby tissues of the first and sec-

L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected] C. K. Carreon Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]

Fig. 27.1  Palatine tonsil at 16-week gestation. This low-power view of palatine tonsil shows the finger-like extension of stratified squamous epithelium surrounded by a small collection of small lymphocytes (H&E, 10×)

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_27

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Fig. 27.2  Palatine tonsil at 16-week gestation. This higher-power view shows that the constituents of the lymphoid aggregate are small lymphocytes surrounding a delicate capillary network (H&E, 20×)

Fig. 27.3  Palatine tonsil at 17-week gestation. This low-power view shows multiple infoldings of stratified squamous epithelium surrounded by a slightly larger collection of small lymphocytes (H&E, 4×)

Fig. 27.4  Palatine tonsil at 17-week gestation. This higher-power view shows that the constituents of the lymphoid tissue are mostly small lymphocytes with a rich capillary network (H&E, 20×)

ular T cells are present in the early midtrimester [5, 6]. Using routine hematoxylin and eosin (H&E) stains, the lymphoid cells in the early midtrimester appear to be mostly small lymphocytes, with occasional larger forms seen (Figs. 27.4 and 27.5), but the formation of definitive lymphoid follicles is not well appreciated. Tonsillar crypts in early gestation are devoid of significant squamous debris. In the late second and early third trimester, the tonsillar lymphoid tissue steadily increases, and vague follicular aggregates can be seen (Figs.  27.6, 27.7, 27.8, and 27.9).

Fig. 27.5  Palatine tonsil at 17-week gestation. This high-power view shows a monotonous population of small lymphocytes with occasional larger forms (H&E, 40×)

Germinal centers are not seen. In addition, keratin debris and sloughed epithelial cells can be seen in the crypts. Although the epithelial basement membrane in fetal and neonatal tonsillar crypts appears to be intact by immunohistochemical markers, transmission electron microscopy demonstrates pores in the basement membrane that allow the passage of lymphocytes into the epithelium [7]. Therefore, it is not unusual to find numerous intraepithelial lymphocytes in the tonsillar crypt epithelium, even in the midtrimester (Fig. 27.10).

27  Palatine Tonsil

Fig. 27.6  Palatine tonsil at 21-week gestation. Note the more robust lymphoid tissue with vague nodularity surrounding an epithelial infolding with central keratin debris; compare with Figs. 27.1 and 27.3 (H&E, 4×)

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Fig. 27.7  Palatine tonsil at 21-week gestation. This higher-power view of a tonsillar crypt contains sloughed and partially keratinized squamous epithelial cells. Bacterial elements are not seen in the crypts during fetal life (H&E, 10×)

Fig. 27.8  Palatine tonsil at 27-week gestation. This low-power view shows the increasing crypt formation and an increased amount of lymphoid tissue (H&E, 4×)

Fig. 27.9  Palatine tonsil at 27-week gestation. Note the increasing crypt formation and increased amount of lymphoid tissue (H&E, 10×)

By term, the palatine tonsils have obtained significant size, and lymphoid tissue is well formed (Figs.  27.11, 27.12, and 27.13), but germinal center formation is still not normally seen. As is the case in other lymphoid tissues, germinal center formation in the tonsil normally takes place postnatally, with exposure to environmental antigens [8]. Other related lymphoid organs in the Waldeyer ring, representing the pharyngeal lymphoid ring, are the adenoids in the nasopharynx, the tubal tonsils posterior to

the opening of the pharyngotympanic or Eustachian tubes into the nasopharynx, and the lingual tonsil in the posterior tongue. Histologically, the main distinction between these different tonsillar tissues is the type of epithelium that lines the surface of the organ and the crypts or folds. The lingual tonsil (Figs. 27.14 and 27.15) shares the same stratified squamous epithelium as the palatine tonsil, whereas the adenoids and tubal tonsils have pseudostratified columnar, ciliated epithelium, as seen in the respiratory tract [1].

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Fig. 27.10  Palatine tonsil in the midtrimester with intraepithelial lymphocytes. This high-power view of palatine tonsil at 21-week gestation shows numerous lymphocytes within the tonsillar crypt epithelium (H&E, 60×)

Fig. 27.11  Palatine tonsil at term. This low-power view shows that the palatine tonsil is fairly large and displays crypt formation, along with an increased amount of lymphoid tissue (H&E, 2×)

Fig. 27.12  Palatine tonsil at term. This slightly higher-power view shows the increased amount of lymphoid tissue. Note the normal lack of germinal center formation (H&E, 4×)

Fig. 27.13  Palatine tonsil at term. This higher-power view shows tonsillar epithelium and the presence of intraepithelial lymphocytes (H&E, 20×)

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References

Fig. 27.14  Lingual tonsil at term. This is a section of the posterior tongue of a 38-week fetus. Note the skeletal muscle bundles of the tongue in the lower half of the image and the squamous epithelium at the top. Small aggregates of lymphoid tissue (arrows) are present beneath the squamous epithelium of the tongue, representing the lingual tonsil (H&E, 4×)

Fig. 27.15  Lingual tonsil at term. Higher-power view of the posterior tongue of a 38-week fetus. An aggregate of lymphoid tissue of the lingual tonsil is present deep to and abutting the squamous epithelium (H&E, 20×)

1. Pantanowitz L, Balogh K. Mouth, nose and paranasal sinuses. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 433–59. 2. Valdes-Dapena MA.  Histology of the fetus and newborn. Philadelphia: Saunders; 1979. 3. Sadler TW, Langman J. Langman’s medical embryology. 11th ed. Philadelphia: Wolters Kluwer Lippincott Williams & Wilkins; 2010. 4. Schoenwolf GC, Larsen WJ. Larsen’s human embryology. 4th ed. Philadelphia: Elsevier/Churchill Livingstone; 2009. 5. von Gaudecker B.  Development and functional anatomy of the human tonsilla palatina. Acta Otolaryngol Suppl. 1988;454:28–32. 6. von Gaudecker B, Muller-Hermelink HK. The development of the human tonsilla palatina. Cell Tissue Res. 1982;224:579–600. 7. Choi G, Suh YL, Lee HM, Jung KY, Hwang SJ.  Prenatal and postnatal changes of the human tonsillar crypt epithelium. Acta Otolaryngol Suppl. 1996;523:28–33. 8. Wittenbrink N, Zemlin M, Bauer K, Berek C.  Exposure to environmental antigens induces the development of germinal centers in premature neonates. Dev Immunol. 2002;9:177–9.

Bone Marrow

28

Michele E. Paessler

Introduction

Shortly after gastrulation, a network of extraembryonic mesodermal cells serves as the site of origin of both the vasIn the developing embryo and fetus, hematopoiesis is a cular and hematopoietic systems. Because both endothelial dynamic process that starts in the yolk sac, continues in and hematopoietic cells are derived from the same cluster the liver, and finally stabilizes in the bone marrow. The role of mesodermal cells, the existence of a common precursor of the spleen in fetal hematopoiesis is controversial and called a hemangioblast has been proposed [2]. These mesois discussed in Chap. 25. Starting at 11  weeks gestation, dermal cells have a spatial orientation, so that the peripheral hematopoiesis begins in the bone marrow and will remain cells acquire the morphology and phenotype of endothelial there throughout adult life. Hematopoiesis begins with self-­ cells, while the central cells disappear to form the first vasrenewing multipotent stem cells that give rise to committed cular lumens. The birthplace of hematopoiesis lies in the progenitor cells, which ultimately differentiate into the cells mesodermally derived clusters of primitive hematopoietic that circulate in the peripheral blood. cells, referred to as “blood islands,” which develop adjacent Because the peripheral blood elements are short-lived, to the newly formed vascular endothelium in the developing hematopoiesis must be a constant process in order to main- yolk sac blood vessels [2, 3]. Hematopoiesis begins with the tain homeostasis. Hematopoiesis is a highly complicated production of primitive red cells in the blood islands of the system that culminates in many well-orchestrated processes yolk sac shortly after implantation. The yolk sac is the priwith the cooperation of stem cells, the supportive microen- mary site of erythropoiesis from 3 to 6 weeks gestation and vironment, and numerous regulatory factors that provide maintains the viability of the embryo until hematopoiesis is stimulatory and inhibitory signals [1]. This chapter reviews established in the liver [2–4]. The blood islands are comthe structure of the normal fetal bone marrow and discusses posed almost exclusively of erythroid cells, but a few megaspecific aspects of myelopoiesis, erythropoiesis, lymphopoi- karyocytes have been identified [5]. Yolk sac erythropoiesis esis, and megakaryopoiesis. is megaloblastic and results in the production of nucleated red cells containing three embryonic hemoglobins Gower I (ζ2ε2), Gower II (α2ε2), and Portland I (ζ2γ2) and, in later Embryology embryos, hemoglobin F (α2γ2) [5]. The onset of circulation allows yolk sac-derived primitive The primary sites of hematopoiesis change during human nucleated erythrocytes to enter embryonic tissues [2]. The development, starting in the yolk sac, proceeding transiently first organ to be colonized is the liver, which is the principal to the liver, and finally stabilizing in the bone marrow. site of hematopoiesis in the embryo from weeks 6 to 22 of Interestingly, the only hematopoietic cells to be produced in gestation (see Chap. 5, Figs. 5.3 and 5.4) [2, 6]. Stem cell the same tissue at all stages of human development are T migration to the liver gives rise to hepatic hematopoiesis, in cells, which develop in the thymus. which hematopoietic foci develop in the hepatic cords during the sixth week of gestation [7]. The fetal spleen also shows foci of hematopoiesis shortly following the liver, at about the seventh week of gestation [8]. Approximately half M. E. Paessler (*) of the nucleated cells in the liver are erythroid cells; only Department of Pathology and Laboratory Medicine, The Children’s a few myeloid and megakaryocytic cells are identified. The Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA erythroid cells at this stage are normoblastic and produce e-mail: [email protected]

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n­ ucleated red cells that contain hemoglobin F. The population of erythroid cells in the liver progressively declines as the transition to the marrow approaches, but occasional erythroid cells can still be detected at the end of gestation and in the early postnatal weeks. After 22  weeks gestation, the bone marrow serves as the permanent site of hematopoiesis and remains so during the rest of gestation and postnatal life. Marrow hematopoiesis commences during the 11th week of gestation in specialized mesodermal structures termed “primary logettes,” which are loose networks of mesenchymal cells that surround a central artery [2]. The marrow cavities are formed when bone or cartilage is eroded by blood vessels and cells from the periosteum [1, 9]. Once the marrow cavities are formed, the vascular connective tissue is organized and becomes colonized by hematopoietic cells [2]. The first blood cells that differentiate in the bone marrow proper are myeloid cells, followed by erythroid cells [2]. Interestingly, the early hematopoietic precursor cells that colonize the marrow are not multipotent hematopoietic stem cells. It is thought that the hematopoiesis in the fetal bone marrow is initially established by committed progenitor cells that provide the essential myeloid and erythroid cells needed by the fetus. Multipotent self-renewing hematopoietic stem cells, in addition to their much-needed supporting stromal environment, are established later in gestation [2, 3]. After the 22nd week of gestation, the marrow becomes the predominant site of hematopoiesis. The most active site of fetal bone marrow hematopoiesis is the vertebral column, followed by the femur, pelvis, fibula/

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tibia, and humerus. Fetal bone marrow erythropoiesis is normoblastic and produces nonnucleated red blood cells that contain hemoglobins F and A.  Fetal bone marrow is also the primary site for intrauterine granulopoiesis and megakaryopoiesis.

Histology In living pediatric patients, the evaluation of bone marrow usually requires an aspirate, as well as biopsy specimens. In fetal autopsies, however, aspirates are not commonly made, so the evaluation of the bone marrow is most often made on the morphologic review of sections of bones. These sections are most often taken from the rib or vertebral bodies. Less often, the iliac crest, long bones, clavicle, temporal bone, or other bones are studied. Ideally, for optimal evaluation of the marrow, the sample should be a thin-cut, well-stained section that has not undergone excessive decalcification. Optimal cytology is usually obtained in the first 3 h postmortem, but cellular detail can be retained for up to 15  h postmortem [10]. Generally, evaluation of the bone marrow includes an estimate of the degree of cellularity, identification of the cell lineages that are present, assessment of the myeloid to erythroid (M:E) ratio, and evaluation of the maturation of the cell lines. Throughout fetal life, the bone marrow is virtually 100% cellular (Fig.  28.1). Occasional adipocytes can be seen. At birth, the marrow is approximately 80–100% cellular [1]. Another important aspect in bone marrow evaluation is the

Fig. 28.1  Bone marrow during fetal life. Note that the marrow is virtually 100% cellular in all samples from various gestational ages. Note the monotonous similarity of the bone marrow throughout the last half of prenatal life (hematoxylin and eosin (H&E), 10×)

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Fig. 28.2  Bone marrow at 19-week gestation. Maturing myeloid progenitors and small clusters of erythroid progenitors are shown. An occasional megakaryocyte is seen (H&E, 40×)

Fig. 28.3  Bone marrow at 24-week gestation. The marrow shows trilineage hematopoiesis. This field shows myeloid progenitors with pink cytoplasm and a few small erythroid clusters. The myeloid to erythroid (M:E) ratio is approximately 1 to 2:1. In addition, an occasional megakaryocyte is seen (H&E, 40×)

Fig. 28.4  Bone marrow at 32 weeks gestation. The marrow remains virtually 100% cellular, with small sheets of maturing myeloid progenitors and small clusters of erythroid progenitors. The M:E ratio is approximately 1 to 2:1. Occasional adipocytes are seen (H&E, 20×)

Fig. 28.5  Bone marrow at 41-week gestation. Periodic acid-Schiff (PAS) stain aids in the distinction between the myeloid and erythroid lineages. Myeloid cells have pink cytoplasm, whereas erythroid cells have gray cytoplasm. Megakaryocytes are also easily identified (PAS, 60×)

assessment of the M:E ratio. In the early second trimester, hematopoiesis transitions to the bone marrow from the liver, and the M:E ratio can be variable (Fig. 28.2). By the end of the second trimester, however, at about 26 weeks, the M:E ratio stabilizes at approximately 1 to 2:1 (Figs.  28.3 and 28.4). A periodic acid-Schiff (PAS) stain may be useful in differentiating erythroid cells from myeloid and lymphoid cells and for evaluating the M:E ratio. On a PAS stain, the myeloid cells have pink cytoplasm, erythroid cells have gray cytoplasm, and lymphoid cells have PAS-negative cytoplasm (Fig. 28.5).

Structure of the Bone Marrow The bone marrow is encased by cortical bone, traversed by trabecular bone, and is composed of a network of capillary-­ venous sinuses that occupy the space between the bony trabeculae. The principal blood supply to the marrow is the nutrient artery and the periosteal capillary network. The nutrient artery supplies the bony trabeculae and branches to form the capillary-venous sinus network, which is an integral component of hematopoiesis. Newly formed hematopoietic cells are released into the periphery via the capillary-venous sinus [1].

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The bone marrow microenvironment is an essential factor in hematopoiesis. A key component in the microenvironment is a heterogeneous group of cells, called stromal cells, which include adipose cells and fibroblasts, among others. These stromal cells play a key role in supporting hematopoiesis by secreting numerous proteins, adhesion molecules, and regulatory products that are necessary for stem cell survival and osteogenesis [1]. In addition to the myeloid, erythroid, lymphocytic, and megakaryocytic lineages, discussed in detail below, a variety of other cell types reside in the bone marrow, such as macrophages, mast cells, osteoblasts, and osteoclasts.

Trilineage Hematopoiesis Hematopoiesis is a spectrum of development that spans from the stem cell to a committed precursor cell that progressively acquires specific lineage characteristics and culminates in the terminally differentiated cells that are released into the circulation. This continuum of maturation is tightly ­regulated at every step by the coordinated expression of many genes [1].

Fig. 28.6 Granulopoiesis. The most immature myeloid cells are located adjacent to the bony trabeculae, and the more mature cells are located away from the trabeculae. Bone marrow at 40 weeks gestation is shown (H&E, 40×)

Myelopoiesis Granulopoiesis Granulopoiesis has a geographic distribution, in that granulocytic elements tend to mature adjacent to the bony trabeculae, where growth factors such as granulocyte colony-stimulating factor and granulocyte-macrophage colony-­stimulating factor drive maturation down the granulocytic pathway (Figs. 28.6 and 28.7). The most immature granulocyte is the myeloblast, which usually accounts for up to 3–5% of the marrow cells. The next stages of maturation are the promyelocyte, myelocyte, metamyelocyte, band, and finally the segmented form. At each successive stage, nuclear and cytoplasmic maturation changes occur. The nuclear chromatin condenses and progressively segments, while the cytoplasm develops secondary granules and becomes acidophilic. These specific nuclear and cytoplasmic changes are best appreciated on bone marrow aspirates. In tissue specimens, immature myeloid cells with moderate amounts of pink cytoplasm are seen adjacent to the trabeculae (see Fig. 28.6), but the different stages of maturation are more difficult to visualize. As the granulocytes mature, they migrate from the bony trabeculae toward the interstitium. Mature granulocytes are identified by their segmented nuclei. Granulocytes are released through the sinus wall into the circulation. In the fetal liver, hematopoiesis consists predominantly of intrasinusoidal erythropoiesis; granulopoiesis is a minor component seen in the portal tracts. The predominant site of granulopoiesis in the fetus is the bone marrow [11].

Fig. 28.7  Granulopoiesis. Immunohistochemical stain for myeloperoxidase demonstrates that the most immature myeloid progenitors are adjacent to the bony trabeculae. Bone marrow at 40 weeks gestation is shown (myeloperoxidase immunohistochemistry, 40×)

Monopoiesis The same myeloid precursor cell from which the granulocytic series differentiates also gives rise to the monocytic lineage. Monocyte differentiation consists of three stages: the monoblast, the promonocyte, and the mature monocyte. Monoblasts have open chromatin and linear condensations, discrete nucleoli, and abundant cytoplasm. As the monoblast matures into a promonocyte and mature monocyte, the nucleus shows increased folding, and the cytoplasm gains azurophilic granules. Maturing monocytes are often difficult to ascertain in postmortem fetal samples.

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Erythropoiesis The most important regulatory factor in erythropoiesis is erythropoietin (EPO). The hypoxic internal milieu of the fetus stimulates high EPO levels, which fall dramatically in postnatal life. Erythroid precursors usually mature in colonies throughout the bone marrow and do not have a specific geographic distribution like the granulocytic series. In the bone marrow, the earliest erythroid precursor is the pronormoblast, and the successive maturational stages include the basophilic normoblast, polychromatic normoblast, orthochromatic normoblast, reticulocyte, and mature erythrocyte, which is nonnucleated. Each stage shows progressive nuclear and cytoplasmic changes. The nuclear chromatin shows progressive condensation and development of a pyknotic nucleus that is ultimately extruded. The cytoplasm initially is intensely basophilic and progressively acquires hemoglobin, imparting a polychromatophilic change to the cytoplasm and eventually an eosinophilic appearance. These stages in erythroid development are best identified on bone marrow aspirates. In bone marrow sections, the erythroid precursors are identified in colonies, and the pronormoblasts are large, with open chromatin and a prominent “boxcarshaped” nucleolus. The more mature erythroid cells are smaller and show dark, round nuclei (see Figs. 28.2, 28.3, and 28.4). Erythroid precursors are the predominant lineage in the fetal liver. In contrast to the fetal liver, erythropoiesis is not the predominant lineage in the bone marrow, where the M:E ratio is approximately 1 to 2:1 after about 26 weeks of gestation [11]. It may be difficult to distinguish erythroid cells from maturing lymphoid cells (hematogones) in tissue sections. A specific immunohistochemical stain, such as glycophorin C or glycophorin A, may be necessary to assess the quantity of erythroid precursors (Fig. 28.8). A PAS stain may also help

to differentiate the two cell types (see Fig. 28.5). Red blood cells that originate from the bone marrow are usually nonnucleated, unlike the red blood cells from erythropoiesis in the fetal liver. In the third trimester of gestation, hematopoiesis transitions from the liver to the bone marrow. By the end of the third trimester and in the newborn, circulating nucleated red blood cells are rarely seen, as the primary site of erythropoiesis is the bone marrow. When the marrow is stressed, however, such as in immune-mediated hemolytic anemias, sepsis, or other clinical situations, one may find nucleated red blood cells circulating in the peripheral blood even during the late third trimester and newborn period. Because the marrow, and not the liver, is the primary source of hematopoiesis from this point on, these nucleated red blood cells are marrow-derived and are released earlier than usual in response to the ongoing stress. In sections of bone marrow from autopsies of fetuses and newborns, the nuclei of normoblasts are frequently abnormal. They may be multiple, irregular, multilobated, or have little blebs (Fig. 28.9). The changes are referred to as dyserythropoiesis or erythroid dysplasia. Dyserythropoietic cells may be mistaken for segmented neutrophils in sections stained with H&E. A PAS stain helps to make the distinction (Fig. 28.10). The dyserythropoietic changes in fetal autopsy material are almost always a nonspecific reactive change due to hypoxia or stress from other causes, but these abnormalities are very similar to those seen in some forms of familial congenital dyserythropoietic anemia. The diagnosis of congenital dyserythropoietic anemia should be made only in the very rare clinical context of profound unexplained anemia, preferably with confirmation by identifying a diagnostic genetic defect.

Fig. 28.8  Erythropoiesis. Shown is immunohistochemical stain for glycophorin C, which stains all red blood cells, including nucleated and nonnucleated forms. When hematogones are numerous (see Figs. 28.11 and 28.12), this stain may be necessary to differentiate hematogones from erythroid cells (glycophorin C immunohistochemistry, 50×)

Fig. 28.9  Dyserythropoiesis. Note the irregular and lobulated outlines of the erythroid nuclei and nuclear budding (arrows). Although congenital dyserythropoietic anemias are a possibility in neonates with anemia and dysplasia, agonal hypoxic events are the most common cause of erythroid dysplasia seen at autopsy. Term bone marrow is shown (H&E, 100×)

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Fig. 28.10  Dyserythropoiesis. This PAS stain shows sheets of red blood cell precursors with nuclear irregularities, including budding and lobations that mimic neutrophils. The PAS stain confirms that the cells in question are dyserythropoietic red blood cells and not neutrophils, which have PAS-positive cytoplasm. Note the gray-colored cytoplasm in the erythroid cells (PAS, 100×)

Megakaryopoiesis Megakaryopoiesis is regulated by many growth and regulatory factors and cytokines, thrombopoietin being one of the most important. Megakaryocytic differentiation is not associated with mitotic divisions such as the myeloid and erythropoietic lineages. Megakaryocytes mature by increasing nuclear lobations, a process termed endomitosis [1]. The most immature form is the megakaryoblast, and maturation to a platelet-forming megakaryocyte is marked by an increase in size and nuclear lobations. In addition, as the megakaryocyte matures, it expresses specific platelet glycoproteins and von Willebrand factor, among other proteins. Megakaryocytes are the most infrequent of the hematopoietic lineages and usually account for less than 1% of the marrow cells. The megakaryocytes are usually seen scattered throughout the marrow adjacent to the sinuses (see Figs. 28.2 and 28.5).

Lymphopoiesis Stem cells differentiate into committed lymphoid and myeloid progenitor cells. The myeloid progenitors give rise to the myeloid, erythroid, and megakaryocytic lineages. The lymphoid progenitors give rise to B, T, and NK cells. The development of B lymphocytes and T lymphocytes is highly dependent on the bone marrow microenvironment. T-cell precursors are derived in the bone marrow and home to the thymus, as this is the site of T-cell maturation. B-cell proliferation and differentiation occurs in the bone marrow and is dependent on numerous growth and regulatory factors such as interleukin IL-1 and IL-7. B cells and T cells can be distin-

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Fig. 28.11  Hematogones. This low-power composite image shows H&E (a) of bone marrow at 38 weeks gestation, compared with a PAX5 immunohistochemical stain (b) to show the abundance of maturing B cells (hematogones) present within the fetal bone marrow. PAX5 is a marker that is positive in all stages of B-cell development. Note that the hematogones are difficult to discern from erythroid precursors by H&E examination alone (a H&E, 10×; b PAX5 immunohistochemistry, 10×)

guished not on their morphologic features but rather on their specific immunophenotypic profile. T cells characteristically express CD3 even in the earliest stages, and B cells express different antigens at specific stages of maturation. The most immature B cells express CD19, CD79a, TdT, CD34, and HLA-DR, and often coexpress CD10. As the B cells mature, CD34, TdT, and CD10 are lost, and CD20 and surface immunoglobulin are expressed in the later stages of maturation. These immunophenotypic features are best appreciated on flow cytometric studies. Maturing B cells are often abundant in fetal and postnatal specimens, as well as in pediatric specimens; they are termed hematogones (Figs. 28.11 and 28.12). Morphologically, hematogones show a spectrum of maturation from lymphoblasts to mature B cells. These morphologic features are best seen on an ­aspirate. In tissue sections, hematogones are intermediate to small in size, with round to slightly irregular nuclei and scant cytoplasm. Hematogones

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Fig. 28.12  Hematogones. This higher-power image compares H&E (a) and PAX5 immunohistochemical stain (b) of bone marrow at 38 weeks gestation. Hematogones may be so numerous that the diagnosis of leukemia may be raised. Immunohistochemical stains for all stages of B-cell maturation are positive in hematogones, whereas in B-cell leukemias, they are positive for only one stage of maturation (a H&E, 40×; b PAX5 immunohistochemistry, 40×)

usually infiltrate through the marrow and may form small aggregates. The amount of hematogones seen in fetal and neonatal bone marrow varies, but they may comprise up to 50–60% of the cells. Hematogones often may be difficult to distinguish from maturing erythroid precursors based on morphology alone. In such cases, PAS or immunohistochemical stains should be used to distinguish erythroid elements from lymphoid elements.

Special Considerations Hemophagocytosis Macrophages are present in the marrow, and phagocytosis of cells and debris is a normal physiologic activity. An increased

Fig. 28.13  Hemophagocytosis. Note a macrophage (arrow) that has ingested several intact cells. Term bone marrow is shown (H&E, 40×)

Fig. 28.14  Hemophagocytosis. Shown is an immunohistochemical stain for CD163, hemoglobin scavenger receptor, which is a macrophage marker. Note the cytoplasmic stain in the enlarged, activated, positive macrophages. The multiple round negative spots in the cytoplasm of the cell in the middle represent phagocytic vacuoles containing phagocytosed blood cells. The single round negative image in the cell in the center right could be the nucleus (CD163 immunohistochemistry, 40×)

number of activated macrophages with phagocytosed red blood cells, granulocytes, and cellular debris are often found in fetal bone marrow (Fig.  28.13). Hemophagocytosis is easier to appreciate on aspirate slides, but it can be apparent in formalin-fixed paraffin-embedded sections. An immunohistochemical stain specific for macrophages, such as CD163, the hemoglobin scavenger receptor, is useful to highlight macrophages and help discern the hemophagocytosis (Fig. 28.14). Hemophagocytosis has many causes. Most cases are a nonspecific reactive change secondary to an underlying process such as infection and the inflammatory response.

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Morphology cannot distinguish between nonspecific reactive hemophagocytosis and familial hemophagocytic syndrome. The typical clinical picture and genetic testing are necessary for a definitive diagnosis of familial hemophagocytic syndrome.

References 1. Foucar K.  Bone marrow pathology. 2nd ed. Chicago: American Society for Clinical Pathology Press; 2001. 2. Tavian M, Peault B. Embryonic development of the human hematopoietic system. Int J Dev Biol. 2005;49:243–50. 3. Peault B.  Hematopoietic stem cell emergence in embryonic life: developmental hematology revisited. J Hematother. 1996;5:369–78. 4. Palis J, Yoder MC. Yolk-sac hematopoiesis: the first blood cells of mouse and man. Exp Hematol. 2001;29:927–36.

M. E. Paessler 5. Wickramasinghe SN. Bone marrow. In: Mills SE, editor. Histology for pathologists. Philadelphia: Lippincott Williams & Wilkins; 2007. p. 799–836. 6. Gilmour JR. Normal haemopoiesis in intrauterine and neonatal life. J Pathol Bacteriol. 1941;52:25–55. 7. Emura I, Sekiya M, Ohnishi Y. Two types of immature erythrocytic series in the human liver. Arch Histol Jpn. 1983;46:631–43. 8. Foucar K, Viswanatha DS, Wilson CS. Non-neoplastic disorders of bone marrow. In: Atlas of nontumor pathology. Washington, DC: American Registry of Pathology Press; 2009. 9. Kelemen E, Calvo W, Fliedner TM.  Atlas of human hemopoietic development. Berlin: Springer-Verlag; 1979. 10. Gilbert-Barness E, Debich-Spicer D.  Handbook of pediatric autopsy pathology. Totowa: Humana; 2005. 11. Kalpaktsoglou PK, Emery JL.  Human bone marrow during the last three months of intrauterine life. A histological study. Acta Haematol (Basel). 1965;34:228–38.

Part VII Central Nervous System and Special Sense Organs

Brain and Spinal Cord

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Patrick Shannon

Introduction

Histology

This chapter reviews the basic cytology of the fetal central nervous system and describes the histological trajectory of the major anatomical subdivisions from the beginning of the second trimester to term. Because of the astonishing complexity and anatomical density of the central nervous system, a comprehensive review of the fundamentals of neuroanatomy is beyond the scope of this text, and the reader is referred to standard works of neuroanatomy and embryology [1–3] for accounts of these specific aspects. This chapter focuses on the clinically and histologically salient points of development. The brain sections illustrated are all immersion-fixed in 5% acetic acid/formalin mixture.

Cytology of the Fetal Central Nervous System

Embryology The central nervous system first makes its appearance as a plate of ectodermal cells on the trilaminar disc of the embryo, which then successively grooves, folds, and fuses to form a closed neural tube. The tube then encloses a fluid-filled space that will be developmentally continuous with the future intracranial ventricles and the central canal of the spinal cord. This is accompanied by the folding and differential thickening of the walls of the tube such that, by late in the seventh week gestational age (GA), the cerebral vesicles, which will form the cerebral hemispheres and olfactory bulbs, are readily discerned, and the midbrain and brainstem are distinguishable from the hemispheres rostrally and the spinal cord caudally. By the end of the 11th week GA, all the major anatomical divisions of the central nervous system are identifiable.

P. Shannon (*) Department of Pathology and Laboratory Medicine, University of Toronto, Mount Sinai Hospital, Toronto, ON, Canada e-mail: [email protected]

The embryonic neural tube and ventricles are lined by a mitotically active, nestin- and vimentin-positive, ciliated, pseudostratified columnar epithelium, the neuroepithelium (Fig.  29.1). The cells are densely packed, with condensed, oval to elongated nuclei, and basal processes that extend into the adjacent tissue. Beneath this is a population of less mitotically active, primitive-appearing cells of variable thickness, which give rise to neurons and then to glial precursors. The neuroepithelium progressively recedes during fetal life at rates specific to the level of the neuraxis considered (see Fig.  29.1), and the ventricle becomes lined by postmitotic ependymal cells with open, oval nuclei. The ciliated choroid plexus is an epithelium continuous with the ventricular ependyma (Fig. 29.2); it displays a transition in morphology, from a flattened cuboidal epithelium with dense, granular amphophilic to eosinophilic cytoplasm near its junction with the ependyma to a granular, ciliated cuboidal epithelium. The stroma generally shows thin-­ walled, dilated blood vessels and can contain foci of extramedullary hematopoiesis. Particularly in the trigone of the lateral ventricles, the choroid plexus may demonstrate cleared granular cytoplasm and a hydropic myxoid stroma. Radial glia are easily the dominant well-differentiated glial component from midgestation through to the early third trimester (Fig. 29.3). They originate from the neuroepithelium and demonstrate round, generally cleared nuclei and prominent, faintly amphophilic, radially directed non-branching processes directed in parallel with their neighbors, with one process extending toward or into the subventricular zone or neuroepithelium and another directed radially toward the pial surface. The radial glia generally stain strongly for nestin and vimentin, although glial fibrillary acidic protein (GFAP) expression becomes more reliably detectable with advancing gestational age. Infratentorially, these cells are

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_29

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Fig. 29.1  Lateral ventricular lining at various gestational ages, medial and dorsal aspect of frontal horn. (a) Neuroepithelium, 13 weeks. The densely cellular, ciliated, and mitotically active neuroepithelium (arrow) lines the lateral and fourth ventricles. In the lateral ventricles, this overlies a germinal matrix composed of primitive cells with generally round, condensed nuclei. (b) By midgestation, in this area, the neuroepithelium persists, but the germinal matrix is becoming more rarefied and less cellular. (c) By 30 weeks, the epithelium is replaced largely by ependyma, although a depleted, hypocellular germinal matrix persists. (d) By 36 weeks, the germinal matrix has almost com-

pletely disappeared, although remnants remain, particularly around veins, and the ventricle is lined by a simple, mitotically quiescent ciliated ependyma (arrow). The germinal elements vary strikingly by geography. (e) In the lateral ventricle (asterisk) at 14 weeks, the neuroepithelium is underlain by the germinal matrix, which is divided into a cell-dense, more proliferative medial zone (M) and a somewhat less cellular lateral component (L). C marks the future head of the caudate and S the septal nucleus. (f) At 12–13 weeks, the neuroepithelium of the third ventricle is still mitotically active (arrow), but it does not overlie a thick, condensed germinal element as in the lateral ventricles

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Fig. 29.2 (a) Choroid plexus, fourth ventricular outflow, 26 weeks gestation. Apparently artifactual separation of the epithelium from the stroma is frequent. Many fronds contain foci of extramedullary hematopoiesis

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(arrow). (b) Choroid plexus in trigone, 22 weeks. Note the distinctly fibromyxoid stroma (arrow) and swollen, almost hydropic epithelial cells. Same magnification as (a). This should not be mistaken for specific pathology

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Fig. 29.3  Radial glia. (a) Subplate, 26 weeks gestation. The radial glial nuclei (arrows) are round and somewhat cleared, with the glial bodies resembling drinking straws in a vertical orientation. Subplate neurons (arrowheads) are recognizable by their cytoplasm. (b) 20 weeks, subplate. In sections stained immunohistochemically for nestin, the radial glia stand out sharply with a finely undulating, beaded staining pattern. At this gestation, the radial glia show very few side processes in the subplate. Only occasional nuclei are densely surrounded by nestin (arrow). Note that

endothelial cells also stain densely. A similar pattern is obtained by immunohistochemical staining for vimentin. (c) 18 weeks, midbrain. An aqueduct is marked with an asterisk. Arrow indicates the dorsal raphe; the ventral raphe is marked by an arrowhead. Note the sweeping pattern of the radial glia laterally. (d) Cerebellar cortex, 18 weeks, stained with immunohistochemistry for nestin. Glial processes extend radially toward the pial surface (asterisk), although cell bodies (arrow) may be difficult to find and typically are deeper than the earliest stainable Purkinje cells

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readily discernible in the spinal cord and brainstem raphe (see Fig. 29.3), and they persist as readily stainable elements there until term. In the cerebrum, radial glia are best appreciated in the hypocellular outer layer of the transitional cerebral mantle. In general, hemispheric radial glia become more difficult to identify as cerebral convolutions develop and axonal tracts mature; by 32 weeks gestation, they can be difficult to identify in the cerebrum. Many radial glia mature into an astrocytic phenotype, though some differentiate into neurons or oligodendroglia or act as pluripotent cells [4–7]. Not surprisingly, the earliest stages of glial differentiation may have nuclear features similar to oligodendroglia. In the cerebellar cortex, there is a specialized population of radial glia, the Bergmann glia, which has a dense layer of cell bodies located just deep to the developing Purkinje cell layer and

P. Shannon

foot processes extending through the external granular cell layer to the pia. Prior to about 30 weeks, these are difficult to distinguish on cytological grounds alone using hematoxylin and eosin (H&E) stain, but they may be seen with vimentin or nestin immunohistochemistry. Astrocytic and oligodendroglial precursors originate from the neuroepithelium; these committed glial precursors proliferate both before and after migration. In midgestation, they can be identified as alpha-beta crystallin and GFAP immunoreactive populations, typically in the dorsal and medial portions of the supratentorial germinal matrix (Fig. 29.4). Glial precursors are identifiable in the cerebral hemispheres at midgestation as cells with round, open nuclei, and nonradially directed processes demonstrating a tendency to surround and ensheathe blood vessels. Early in development, they may

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Fig. 29.4  Astrocytic development. (a) At 22  weeks, glial elements stain strongly for alpha-beta crystallin in the medial and dorsal component of the frontal germinal matrix (arrowhead). (b) Within the adjacent intermediate zone, small vessels are ensheathed with glial cells, some of which are separate from the vessels but extend a stubby foot process to the vessel (arrow). Some of these cells are probably radial glia (arrowhead), while others extend radiating processes (small arrow). (c) At 28 weeks, cells positive for glial fibrillary acidic protein

(GFAP) with clearly astrocytic morphology have begun to develop in the subplate; the parallel rows of radial glial processes can be seen in the background. These are difficult to appreciate with H&E staining, as they have a protoplasmic astrocytic phenotype, with round, open nuclei like those of radial glia and inconspicuous cytoplasm. (d) Fibrillary astrocytes are now readily appreciated in subpial areas, with irregular, dark nuclei and glassy, dense, radiating processes

29  Brain and Spinal Cord

extend a thick foot process laterally toward a blood vessel. Astrocytes, specifically, are derived from glial precursors, with a contribution from the radial glial cells. They make their first appearance at roughly 15 weeks gestation, in the spinal cord. Not surprisingly, their nuclei are difficult to distinguish from those of radial glial nuclei early on, and only in the third trimester can they be differentiated easily on H&E-stained sections in the cerebral mantle. Prior to this, in the cerebral mantle, their usual eosinophilic cytoplasm can be very difficult to discern, although immunohistochemical staining for GFAP leaves little doubt as to their morphology. They are readily discerned at midgestation in the basal ganglia, the subpial region of the basal forebrain, the hippocampal alveus, the brainstem, and the spinal cord, however. The appearance of astrocytes follows the general caudal to rostral sequence of maturation in the infratentorial compartment, and the general central to peripheral pattern in the supratentorium. In the spinal cord, brainstem, and basal ganglia, the density of GFAPpositive processes in normal maturation is such that it may prompt consideration of a pathological process. Care is generally required before assigning any such significance. Oligodendrocytes are the last of the major neuroepithelial lineages to appear; they are derived from the periventricular zone and undergo multiple morphological changes before adopting their mature, myelinating form (Fig. 29.5) [7–9]. In midgestation, their precursors are distributed in an area with poorly defined borders laterally and dorsally in the germinal matrix and staining for alpha-beta crystallin and SOX10. As these differentiate into myelination glia, they demonstrate eccentrically placed, round, and partially clear nuclei, and a few prominent, occasionally refractile, eosinophilic processes, mimicking reactive astrocytes. The consistent eccentricity of the nucleus and the geography of their prominence, the timing of which is stereotypically related to early local myelination, are key to their identification. Occasionally, myelination glia may demonstrate voluminous, vesicular cytoplasm that can be immunoreactive for GFAP and occasionally myelin basic protein (see Fig. 29.5) and may contain neutral lipid [10]. The significance of this morphology is unknown. In normal development, by the time myelin basic protein immunoreactivity is pronounced in a tract and myelin sheaths may be seen extending around processes, the nuclei have the familiar round, homogenous basophilia of oligodendrocytes. Near term, many oligodendroglia have very distinct perinuclear caps of cytoplasm in well-fixed specimens. In general, standard histochemical stains for myelin are not very useful for detecting the small amounts of immature myelin in the fetal central nervous system, and immunohistochemistry is preferred. Immunohistochemistry tends to be more sensitive for detecting myelination than histochemical stains [11], so there may be some discrepancy between immunohistochemical staining and the classic studies [12, 13] on the subject.

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Neurons have the most diverse phenotypic range and dramatic developmental trajectories. Though knowledge of precise cytoarchitectural location and gestational age is always important in evaluating the normality of central nervous system tissue, in no case is this more important than the evaluation of neuronal populations. In general, mature neurons are characterized by a vesicular, round nucleus with a prominent nucleolus, with the perinuclear cytoplasm characterized by a basophilia that may be fine and dustlike or organized into platelike aggregates. Most neuronal populations arise from cells with sparse cytoplasm and dark, homogenous chromatin. The perikaryal cell borders of most mature neurons are readily distinguished, but maturation usually involves expansion of the cytoplasm in finely dividing dendrites and projecting axons to give rise to the feltlike neuropil. At the beginning of the second trimester, the neuropil is poorly developed and consists of wisps of poorly staining processes in an abundant extracellular matrix rich in hyaluronate. With maturation, astrocytic and myelinating elements contribute, neuronal processes become more abundant, and the neuropil becomes more coarse and refractile. Neuronal populations mature at differing rates, according to the developmental sequences specific to their particular local anatomy. The neurons of the spinal cord, much of the brainstem, the thalamus, and the basal nuclei, derive directly from primitive periventricular populations early in development, and for most of fetal life, the histological changes are those of maturation of populations established in embryonic life, with eventual maturation of astrocytic elements and myelination. However, the bulk of cerebral cortical neurons arise from the ventricular zone or germinal matrix of the lateral ventricles. They then either migrate radially (as in large projection neurons) or migrate ventrally to the basal forebrain and from there spread out over the surface of the convexities before moving deeply into the layers of the cortex (as in many interneurons), migrating from the germinal matrix tangentially into the intermediate zone of the cortex, and then down into the ventricular zone before undertaking their final radial migration, or they may follow even more chaotic trajectories [14–16]. Complex migrational pathways are also present in the cerebellar cortex, where the internal granular cell layer of the cerebellar cortex derives from primitive cells in the external granular cell layer of the cerebellum. They descend through the molecular layer to take up their final position in the cerebellar cortex (see below); a process that is not complete until postnatal life [17]. Whatever the path taken, the morphological trajectory is similar in that it involves the movement of cells with oval to circular, condensed nuclei, and sparse, often bipolar processes into their definitive positions, where they mature into their more easily recognizable mature phenotype. Prior to achieving their final adult morphology, maturing neurons may express a variety of neuronal epitopes (e.g., calbindin, neurofilament light chain, calretinin, SATB6), which can

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Fig. 29.5 (a) Myelination-associated glia, 30  weeks gestation. Note the round nuclei (arrow), often with a sharp peripheral chromatin ring, and eccentrically placed eosinophilic cytoplasm, giving a general appearance of a ladybug. These astrocyte-like cells proliferate in tracts prior to myelination onset. (b) Mature oligodendrocytes, pontine tegmentum, 40 weeks. The nuclei are condensed and dark; though some have a perinuclear halo (arrowhead), others have distinct perinuclear condensations of cytoplasm (arrow). Within white matter tracts, mature oligodendrocytes are often arranged in short linear arrays. (c)

Occasionally, myelination glia can be identified by pale foamy cytoplasm (arrow) and a large nucleus with open chromatin. (d) On staining for alpha-beta crystallin or GFAP, myelination glia may demonstrate voluminous cytoplasm. (e) In early myelination, the oligodendrocyte can be seen extending a few fine processes and ensheathing axons (external capsule, 30 weeks). (f) Unmyelinated tracts (e.g., basis pontis, 23 weeks, corticofugal tracts) resolve as densely packed masses of fine eosinophilic processes that in cross section may seem to show slight turbulent swirling

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aid in their identification. The axonal tracts of the fetal nervous system have a distinct histology (see Fig. 29.5). Prior to myelination, developing axons can be identified in cross sections as eosinophilic dots, often arranged in slightly swirling bundles. These may be stained using immunohistochemistry for microtubule antigens or neurofilament light chain antibodies. Some long tracts, most strikingly the pyramidal tracts of the brainstem, can be readily highlighted by immunohistochemical staining for vimentin. The vascular supply of the embryonic central nervous system starts as a continuous thin-walled sinus that extends over the surface of the central nervous system after neural tube closure and progressively organizes to the thin-walled arteries and veins of the late embryonic period; by the end of the first trimester, large arteries can be distinguished from large veins by the organization of their media and adventitia. The meninges are divided into the pachymeninges (dura mater) and leptomeninges (arachnoid and pia mater). By the end of the first trimester, the dura can be recognized as loose, fibromyxoid tissue in continuity with the arachnoid and containing thin-walled vascular channels; it adopts an obvious multi-plied texture recognizable as dura by midgestation (Fig.  29.6). The craniospinal dura originates as continuous with the fibrous tissue giving rise to the calvarium and the periosteum of the chondrocranium. In the cranium, the dura remains continuous with the periosteum, but in the spinal column, it separates from the spinal periosteum dorsally and laterally by the end of the embryonic period. However, it maintains fibrous continuity ventrally, especially in the lumbar cord, to form a sac around the spinal cord (the thecal sac), which ends bluntly around the cauda equina in the sacrum. In the spinal column, the epidural space is largely filled by myxoid material and dilated sinusoidal vessels. Until the third trimester, mature fat is not generally present in the spinal epidural space, but adipocytes can be readily recognized in the neural foramina by midgestation. The arachnoid is the fine fibrous coating surrounding the brain and spinal cord. It has an outer fibrous leaflet, which, although continuous via fine cellular processes with the dura, readily separates from the dura on blunt dissection (see Fig. 29.6). Beneath this outer layer are the arachnoid trabeculations, a meshwork of collagenous processes through which the large fetal arteries and veins course before entering the brain. Also within the subarachnoid space are generally found numerous foamy histiocytes. The surface of the brain is covered by the pia mater, which has two components: the epipia, which is a collagenous membrane in continuity with the arachnoid trabeculae and secreted by fibroblasts, and the intima pia, which is a very delicate membrane secreted by the end feet of the radial glia and later by astrocytes. The intima pia can be stained on well-oriented specimens with periodic acid-Schiff (PAS), reticulin, or

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immunohistochemical stains for laminin or collagen IV, in which it shows as a razor-sharp demarcation between the brain and subarachnoid space. Microglia, the resident immune effector and phagocytic elements of the central nervous system, establish themselves from granulocytic precursors and initially take up residence in the subventricular embryonic human brain. By midgestation, they are strikingly nonuniformly distributed in the brain. Large numbers are present in the neuroepithelium and subventricular zone, but increases in local density near the developing internal and external capsule and adjacent to perivascular spaces are common and nonspecific (see Fig. 29.6), and foamy macrophages are readily identified in the leptomeninges. These should not be mistaken for postnecrotic changes. In the spinal cord and brainstem, scattered microglia are often noted near perivascular spaces, but individual cells infiltrate into the neuropil in parallel with the degree of local myelination. In general, as myelination extends into the central nervous system, there is an accompanying sparse population of microglial cells.

 evelopmental Anatomy and Histology D of the Fetal Central Nervous System  pinal Cord and Brainstem S In embryological development, the spinal cord is generated from the flat neuroepithelial plate by elevation and fusion of lateral and dorsal elements, such that it forms a four-sided tube surrounding a neuroepithelium-lined ventricle, which will become the central canal [2, 3] (Fig. 29.7). This canal persists into postnatal life as a continuous ependymal-lined tube, marked dorsally by a prominent raphe extending between the dorsal columns. The neural elements of the spinal cord derive from periventricular precursors in embryonic life, such that spinal cord motor neurons are readily recognizable by the end of the embryonic period; by 14  weeks, spinal cord neurogenesis is complete. Radial glia persist until term and can readily be appreciated in the dorsal raphe. Myelination begins in the dorsal columns and propriospinal tracts at roughly 16 weeks gestation and is prominent by its sparsity in the dorsolateral columns until postnatal life [12]. The spinal column elongates relative to the spinal cord in late embryonic life, such that the cord initially extends the length of the column, but by 27 weeks gestation, it extends only to the mid lumbar spine. The pons, medulla, and cerebellum (collectively, the rhombencephalon) derive from segmental cranial condensations of neuroepithelium (rostrally to caudally termed rhombomeres 1–8) as well as part of the isthmus (the narrowing of the embryonic brain rostral to rhombomere 1). In the rhombencephalon, the alar plate of the embryonic neural tube is largely membranous. The future cerebellum starts as a

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Fig. 29.6  Meninges and microglia. (a) The dura and dural reflections at the start of the fetal period are composed of cellular fibrous tissue in visible continuity with fine fibrous tissue strands of the arachnoid (arrows). The hemorrhage here is artefactual (tentorium, 24 weeks gestation). (b) Convexity dura at 40  weeks gestation is composed of a multi-plied fibrous tissue with layers of collagen fibers running at right angles to each other (cross polarized light with lambda filter). (c) The arachnoid at midgestation over the cerebellum and in some basal cisterns is fibromyxoid, containing foamy and cleared cells. (d) By contrast, the supratentorial arachnoid is generally more similar to the

neonatal anatomy in the fetal period, with a fine arachnoid at left, and a sharply defined pia (arrow) here stained with immunohistochemistry for collagen IV. (e) Microglia are nonrandomly aggregated in the fetal brain. Near midgestation (here at 17 weeks), aggregates are often found near the internal capsule (arrowhead), the external capsule (arrow), in germinal matrix or neuroepithelium (small arrows), and notably in the walls of the cavum septi pellucidi (asterisk) and vergae (immunohistochemistry for CD68). (f) A sparse scattering of resting microglia cells (arrow) is a normal feature of subcortical gray matter and myelinated white matter (spinal cord, 22 weeks, immunohistochemistry for CD68)

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Fig. 29.7  Maturation of the spinal cord. The lumbar enlargement is shown and oriented with dorsal at the top of the figure. At 13 weeks, the moth wing profile of the central gray with surrounding hypocellular tracts is obvious. By 17 weeks, the motor neurons are arranged in obvious aggregates (small arrows), as in the adult form, and the ventral roots (arrow) are well-myelinated and darker on H&E than the poorly myelinated dorsal roots (arrowhead). By 22  weeks, myelination has become obvious in the dorsal columns (arrowhead) and is expanding

the cord at that point, although the profile is still somewhat rectangular. Note also that the epidural space is occupied by myxoid connective tissue containing large blood vessels and is devoid of mature adipose tissue. Six weeks later, at 28 weeks, the cord is round, and now the dorsal roots are the same diameter as, or larger than, the ventral roots. By near term, at 35 weeks, the hypomyelination of the dorsolateral columns is still obvious (arrows), even on H&E. At 41 weeks, the morphology is that of a neonate

c­ ondensation of cells at the rostral and lateral margins, which expands laterally, medially, and dorsally to form the cerebellar hemispheres. Neuronal precursors of the rhombencephalon arise from periventricular cells from the floor of the future fourth ventricle (the ventral region) as well as from the rostral and ventrolateral margins of the alar membrane (the rhombic lip, which gives rise to some cerebellar neurons, the nucleus basis pontis, and the inferior olivary nuclear ­complex) [2, 3]. The basic brainstem nuclear organization is established by the end of the embryonic period. Primitive neuroepithelial elements are still present at the beginning of the third trimester in the lateral recess of the fourth ventricle, and rhombic lip-derived precursors giving rise to the neurons of the nucleus basis pontis can still be seen laterally in the caudal pons and lateral medulla until 18–20  weeks gestation (Fig. 29.8). These may persist until midgestation. At about 18  weeks, the inferior olivary nuclear complex is still a smooth, cup-shaped mass, but it may be beginning to acquire crenation. Myelination is largely restricted to the cranial nerve roots, medial lemniscus, and tractus solitarius [2, 3, 12]. By midgestation, olivary crenation is usually present

though incomplete, and myelination is distinct in the medial longitudinal fasciculus and inferior cerebellar peduncle. By the beginning of the third trimester, there is extensive early myelination of the medullary and pontine tegmentum including the amiculum olivae, and by term, all but the middle cerebellar peduncles and corticospinal and corticopontine tracts demonstrate poor myelination. Throughout, the corticospinal tracts can be striking in their dense staining for vimentin (personal observation). The dentate nucleus at 22  weeks starts to develop distinct crenation [17]. Myelination starts as single cells in the white matter of the vermis and flocculus and immediately around the dentate and nuclei interpositi at roughly 22 weeks gestation, but by 27–30 weeks, this same pattern is beginning to outline fiber tracts in these areas as well as the inferior and superior cerebellar peduncles. By term, the middle cerebellar peduncle is beginning to myelinate [12].

Cerebellum The cerebellum is first distinguished in the late embryonic stage as a horseshoe-shaped mass, with its apex pointing rostrally, at the rostral and lateral margins of the membranous

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Fig. 29.8  Brainstem. (a) 17 weeks open medulla, near outflow of the fourth ventricle (asterisk). Arrows indicates remnants of rhombic lip; p marks the pyramid. (b) 17 weeks, lateral half of mid pons, rotated so ventricle is marked with asterisk. Arrow indicates neuronal progenitors from rhombic lip, invading the lateral pons; bp indicates nucleus basis pontis; V is the motor nucleus of the fifth cranial nerve. (c) Deep cerebellar nuclei, 17 weeks. Arrows outline smooth, poorly delimited den-

tate nucleus laterally and nuclei interpositi medially; asterisk marks the ventricle. (d) At 22 weeks, the dentate is distinctly crenated (arrow), and the nuclei interpositi are well defined. (e) Ventral medullary tegmentum, 17 weeks. The inferior olivary nuclear complex has smooth contours, although all its elements are present. p pyramid. (f) By 22 weeks, the olive has distinct crenations

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roof of the fourth ventricle. By 14 weeks, the cerebellum has begun to form gross lobes, and the primary fissure is demonstrable in the vermis. By midgestation, the lobules of the cerebellar vermis are all formed, and elaboration of cerebellar folia continues into the third trimester [2, 17]. Although the mature cerebellar cortex is remarkably cytoarchitecturally homogenous, maturation rates vary within the fetal cerebellum, with the vermis and flocculonodular cortex maturing earlier than the neocerebellum. The cytological elements contributing to this maturation are complex (Fig.  29.9). At about 14  weeks gestation, the cerebellum is present as a mostly smooth-surfaced mass with a distinct external granular cell layer derived by migration over the surface of the cerebellum from the rhombic lip. This external granular cell

layer generates neurons that migrate from the superficial to the deep cortex, through the future Purkinje cell layer, to form the internal granular cell layer and the basket cells of the mature cerebellar cortex. The external granular cell layer also gives rise to the stellate cells of the future molecular layer. Deep to the external granular cell layer are a cell-poor marginal zone and a deep, poorly defined and large cellular zone containing the future Purkinje cells migrating from the ventricular zone and the neuronal precursors that will form the dentate and cerebellar roof nuclei. At 14  weeks gestation, individual cerebellar neuronal populations are difficult to discern, but Purkinje cells can be discerned as distinct cell types at 16–18  weeks gestation, starting in the vermis, and there is a definite developing

Fig. 29.9  Cerebellar cortical maturation, vermis. At 13  weeks gestation, the marginal zone (M) is between a thin external granular cell layer (EG) and a thick, architecturally homogenous (on H&E) layer of migrating Purkinje cell and dentate neurons. The external granular cell layer is bilaminar (best illustrated here at 30 and 35  weeks) and persists until postnatal life. The marginal zone (future molecular layer, M) is populated prior to midgestation by numerous descending external granular cell layer derivatives, but by midgestation, the marginal zone is

expanded, and Purkinje cell dendrites are easily seen. At 17  weeks, Purkinje cells (P) are discernible from other elements only on careful inspection at high power, but by 22 weeks, they form an obvious layer. Between 30  weeks and term, they gain large amounts of cytoplasm. Slightly deep to the future, Purkinje cell layer is the cerebellar granular cell layer (G), which is thin but distinct from the underlying future white matter by 20–22 weeks gestation. The lamina dissecans (LD) is immediately deep to the Purkinje cell layer and absent after 32 weeks

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molecular layer throughout. At the same time, the dentate nucleus is vaguely discernible as a smoothly contoured, deep cell mass with a discernible, hypercellular, neuron-poor hilum (see Fig.  29.8). Slightly deep to the future Purkinje cell layer is the cerebellar internal granular cell layer, which has begun to accumulate by migration from the external granular cell layer and glial cell bodies with large, open nuclei, and processes that extend to the pia can be discerned immediately deep to the Purkinje cell layer (see Fig. 29.3). By 20–22 weeks gestation, the external granular cell layer is thick; it persists until postnatal life as a bilaminar structure consisting of a proliferative outer layer with small, dark nuclei and a postmitotic deeper layer. The molecular layer is distinct, and the Purkinje cell nuclei start to become distinctly larger and the cell layer more compact. At 22 weeks, a cell-poor, axon-rich transient layer (the lamina dissecans) forms immediately deep to the Purkinje cell layer and persists until 32  weeks gestation. The cerebellar cortex thus begins as having three layers, but by midgestation it has five (external granular cell layer, marginal, Purkinje cell, lamina dissecans, internal granular cell); at birth it has four, having lost the lamina dissecans. It adopts an adult configuration months after birth, when the external granular cell layer disappears. The dentate nucleus at 22 weeks starts to develop distinct crenation. Myelination starts as single cells in the white matter of the vermis and flocculus and immediately around the dentate and nuclei interpositi at roughly 22 weeks gestation, but by 27–30 weeks, this same pattern is beginning to outline fiber tracts in these areas as well as the inferior and superior cerebellar peduncles. By term, the middle cerebellar peduncle is beginning to myelinate.

Midbrain and Thalamus The midbrain develops from neuromeres M1 and M2, with neuronal cell masses deriving directly from periventricular neuroepithelium. The nuclear architecture is established by the end of the embryonic stage, as is the distinction of the superior and inferior colliculi [2, 12, 13]. The pattern of myelination is similar to the pons, with early nerve root myelination established by midgestation. Inferior colliculus myelination and medial lemniscus myelination start at roughly the same time, with superior cerebellar peduncle and ascending auditory tract myelination starting toward the end of the second trimester. Myelination in the brachium of the superior colliculus has started by term. Cerebral Hemispheres At the time of rostral neuropore closure, the primordia of the cerebrum are only slightly longer than those of the midbrain and derive from a single prosencephalic neuromere, which also gives rise to the optic primordia. This prosencephalic neuromere expands massively and undergoes a bewilder-

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ingly complex differentiation, giving rise to the diencephalon (thalamus, hypothalamus, epithalamus, and neurohypophysis) and the telencephalon (the cerebral cortex, amygdala, olfactory bulb and tract, and basal ganglia). By the end of the first trimester, the cerebral hemispheres are the largest masses in the central nervous system, and their surface development is one of the most dramatic anatomical features of fetal life (Fig. 29.10 and Table 29.1). The basic deep nuclear organization is established in the embryonic period: the thalamic and basal nuclei originating directly from periventricular zone neural precursors. In the fetal period, the major histological changes in the deep nuclei are those of progressive neuronal maturation with a disappearance of primitive elements, the latter of which is essentially complete other than isolated rests prior to ­ 16–18 weeks gestation. Myelination begins around 30 weeks in deep tracts, and by term, myelination in the cerebrum is well established only in the ansa lenticularis, optic chiasm and tract, globus pallidus, anterior commissure, and central corona radiata. The organization of the cerebral mantle and its various cortical areas, however, undergoes substantial histological evolution. Cerebral cortical development is one of the most dramatic anatomical features of fetal life (see Figs. 29.10 and 29.11). The description that follows must be regarded as highly schematic. At the start of the fetal period, the lateral ventricles are relatively large and are lined by a mitotically active, pseudostratified, ciliated neuroepithelium (see Fig. 29.1), underlain by a subventricular zone of primitive, proliferating cells, which give rise to migrating neuronal and glial precursor cells. This subventricular population is initially continuous under the neuroepithelium and is of varying thickness, but by the beginning of the second trimester, it concentrates in the germinal eminence of the lateral ventricles, which consist of a thick, mitotically active cellular mass extending from the lateral and ventral frontal recess, over the caudothalamic groove, and arching posteriorly into the anterior and lateral trigone and continuously into the temporal horn. By midgestation, the germinal matrix in the frontal horn demonstrates two major cytological divisions: a dorsolateral, somewhat more compact aggregate of cells with open, ovoid nuclei and definite but small amounts of amphophilic cytoplasm and a ventral component of small cells with round to irregular cells that extend into the surrounding brain, often in small clusters, and tend to be more mitotically active. (This rough compartmentalization is dorsoventrally inverted in the temporal horns). The medial, ventral component prominently gives rise to interneuron precursors, which migrate tangentially over the hemispheres, followed by descent into the cerebral cortex. There is also a basally directed stream of precursors toward the olfactory bulbs, which represent outpouchings of the telencephalon and often contain ciliated neuroepithelium. This olfactory stream persists into postnatal life.

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Fig. 29.10  External cerebral development, 18–38 weeks, gross. Note the changes in the Sylvian fissure (S). At 16 weeks, the insula is a simple, smooth-edged notch. At 20  weeks, the Rolandic (central) fissure (arrow) is usually obvious, though shallow. The Sylvian fissure is wide open, the insula (i) at this point is large and is bounded superiorly by a shelflike overhanging ridge of cortex, which will become the frontal operculum. By 22 weeks, the posterior Sylvian fissure is beginning to close, and the prerolandic fissure is present; at 24  weeks, the post-­ Rolandic sulcus is discernible. At 26 weeks, the superior temporal sul-

cus (arrow) is reliably present bilaterally, and by 28 weeks, the superior frontal sulcus and inferior frontal sulcus are well defined. From 30 weeks onward, the majority of gross cortical evolution takes place by elaboration of secondary sulcation and increased definition, but the inferior temporal gyrus often is not obvious until 36  weeks (arrow). Note that at all ages, the presence of leptomeningeal congestion and subarachnoid hemorrhage, scant amounts of which are normal findings at post mortem, accentuate the sulcal development, which is usually mildly effaced by postmortem fluid shifts

At the beginning of the second trimester, the future neocortical mantle consists of five major zones (Fig. 29.12). On the outside, there is a subpial granular cell layer, containing neuronal precursors and small neurons originating by tangential migration. This granular cell layer is transient; it covers the cortical surface by 16 weeks gestation and disappears after about 30 weeks. Immediately subjacent to the subpial granular cell layer is the hypocellular marginal zone, containing ascending cortical dendrites from neurons of the cortical plate, large reelin- and calretinin-positive neurons of Cajal-Retzius, and a meshwork of calretinin-positive processes. This marginal zone will become the future cortical layer I. Deep to this is the developing cortical plate. Neurons are progressively added to the cortical plate by at least two pathways: The first is migration from the ventricular zone along radial glia (“radial migration”) in an “inside out” fashion, such that future layer VI neurons arrive in the future cortical plate first, followed by more superficial layers, with the last migrating cells being those of the future layer II. This process peaks at midgestation and is the source of large pro-

jection neurons. The other pathway is tangential migration, which can follow complex trajectories [14–16]. Initially, the cortical neurons have a simple, bipolar morphology, with scant cytoplasm, an apical dendrite, and a basal axonal projection, so that the cortical plate prior to 20 weeks gestation consists of a homogenously packed cellular layer on H&E. By midgestation, reciprocal axonal projections from the thalamus to the cortex are already established (though not complete), and at 20–22 weeks, a poorly defined pyramidal cell population can be discerned in most cortical areas (Fig.  29.13), which expresses markers corresponding to roughly layer V [18]. Differentiation of the cortical layers takes place rapidly thereafter, and by 30 weeks, a six-layered neocortex is obvious throughout, though with substantial differences in cytoarchitecture depending on cortical location. Cortical sulcation develops in an orderly fashion, with primary sulcation completed by late preterm, and secondary sulcation is usually complete by 40  weeks gestational age (see Table 29.1). Immediately deep to the cortical plate is a poorly delimited area containing neurons recognizable by

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Table 29.1  Sulcal development Gestational age Medial surface GW 8–11 Hippocampal GW 12–15 GW 16–19

GW 20–23

Callosal Parieto-occipital Calcarine Cingulate Collateral

GW 24–27

GW 28–31 GW 32–35

Marginal

Insular

GW 36–39

GW 40–44

Secondary sulci from: Callosomarginal Insular

Base/inferior Superolateral surface surface Interhemispheric Lateral fossa Lateral Olfactory

Rolandic/central Superior temporal Precentral Postcentral Superior frontal Lateral occipital Intraparietal Inferior temporal Inferior frontal Secondary sulci from: Superior and inferior frontal Superior temporal Intraparietal, occipital Secondary sulci from: Transverse and inferior temporal Cingulate Tertiary sulci from: Superior and inferior frontal Intraparietal Tertiary sulci from: Inferior temporal Occipital

Orbital

Secondary sulci from: Orbital

GW gestation week

18  weeks (subplate neurons), which express calretinin and send projections into the cortical plate and to the thalamus and are involved in the establishment of point-to-point reciprocal corticothalamic connections.

The neural elements in the region between the subplate and the ventricular zone (the intermediate zone) comprise radially and tangentially oriented axonal tracts, migrating differentiated glial and neuronal cells and radial glia. The histology and arrangement of these elements vary with the regional anatomy and overlying cortex. One schema for discussing the regional anatomy of the cerebral mantle divides the intermediate zone into six layers (see Fig.  29.12) [19]. This arrangement is obvious at low-power magnification by 15  weeks gestation, but with migration and maturation, it becomes difficult to discern by 30 weeks. Specific mention should be made of the hippocampal formation, which emerges early as a distinct cytoarchitectural unit by the beginning of the second trimester (Fig.  29.14). The fascia dentata and cornu ammonis are both discernible by 15  weeks, and the characteristic seahorse shape is well defined by 16 weeks gestation. The major cytoarchitectural divisions (cornu ammonis, subiculum, entorhinal cortex) are distinguishable by 20 weeks.

Special Considerations In a discussion of normal histology, some mention should be made of histological findings that are common but sometimes confusing (Fig. 29.15). Prominent among these are the papillae of Retzius; the midgestation brain may display prominent tufting of the superficial cortical plate into the marginal zone, giving an appearance of abnormal cortical maturation and migration. This appearance is a consequence of even mild autolysis and should not be interpreted as a migrational anomaly [20]. A second finding often questioned is the discontinuity and inhomogeneity of the external granular cell layer of the cerebellar cortex, particularly in term infants. The foci of discontinuity in this layer are often accompanied by a mounding up of the layer between the defects, occasionally yielding an almost rhythmic pattern. This histology can be seen in a very wide variety of pathologies, as well as in otherwise normal brains in well-explained sudden asphyxial deaths [21]. It tends to accompany areas in

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which the leptomeninges are at least partially separated from the superficial cerebellum, and the defects often correlate with the presence of superficial penetrating blood vessels. Particularly between 20 and 28 weeks gestation, it is often possible to find herniations of germinal matrix tissue through the neuroepithelium and into the ventricles. Such nodular excrescences may be floridly increased in cases of pathological ventriculomegaly or destructive pathology, and they rarely may obstruct the aqueduct (personal observation), but

they are of little diagnostic significance, if rare, in an otherwise normal brain. Similarly in the cerebellum, deeply situated nests of disorganized neurons with germinal elements or jumbled foci of developing cerebellar cortex can occasionally be found as isolated lesions. These represent local derangements of normal cerebellar development and may be consequent to destructive or malformative conditions, but if they are isolated and focal, they are not suggestive of any particular diagnostic entity [10].

Fig. 29.11  Coronal sections, cerebral hemispheres just posterior to the interpeduncular cistern (not to scale). At 18 weeks, the lateral ventricles (LV) are capacious, and the choroid plexus (CP) and germinal matrices (gm) are relatively large. At this gestational age, this level is near the atrial trigone, so the frontal and temporal germinal matrices are almost in continuity on a coronal section. The hippocampal formation (H) is readily appreciated, although not fully rotated. The corpus callosum in this specimen is artifactually ruptured. At 22 weeks, the germinal matri-

ces are still large and are readily demarcated from adjacent brain by their pale color; cc, corpus callosum. By 26 weeks, the superior temporal sulcus (sts) and cingulate sulcus (cs) are readily visible on coronal sections, and the germinal eminences have distinctly decreased in relative size. By 30 weeks, the Rolandic sulcus (arrow) is a deep and consistent feature. Thereafter, secondary gyration predominates the picture, and by 40  weeks, myelination is grossly obvious in the internal capsule (arrow)

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Fig. 29.12  Development of cortical mantle, 13–36  weeks, lateral occipital lobe, except where specified. Lateral ventricle on left, leptomeningeal surface on right (not to scale). At 13 weeks, six layers of the mantle, extending from the ventricular zone (VZ) to the marginal zone (MZ) are obvious, but by 17 weeks, the intermediate zone (IZ) has massively expanded, and the interaction between cellular migration and axonal tract orientation produces a distinct lamination, with layers designated 1–6. In the occipital lobe, layer 2 is inconspicuous, and there is a very dense layer 5. By contrast, in the frontal convexity at 17 weeks, layer 6 is expanded, without a striking layer 5, and there is a layer 2.

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The subplate (SP) is distinguished by a population of well-­differentiated, radially directed neurons. By 22  weeks, the lamination is even more obvious, and the ventricular zone is relatively smaller; by 28 weeks, the detailed lamination pattern has largely disappeared. The region beneath the subplate can be divided into an inner intermediate zone (IZi) and an outer intermediate zone (IZo), based on the degree of cellularity and the presence of interlacing fiber bundles. By 36 weeks, the ventricular zone is almost completely depleted and the intermediate zone, which will become the future subcortical white matter, has hugely expanded. CP cortical plate

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Fig. 29.13  Cerebral neocortex (images not to scale). (a) At 22 weeks gestation, the prefrontal cortex demonstrates a distinct subpial granular cell layer (arrowhead) with a deep, more optically opaque layer of apical dendrites projecting up from the superficial cortical plate (arrow). Artefactual separation in the cell-poor region between these two features is common. The cortical plate does not show obvious lamination at low power at this stage, although careful search on H&E will demonstrate a vague deep layer of pyramidal neurons. The deep margin of the cortical plate is poorly defined. (b) By 40 weeks gestation, there is no obvious

myelin, and lamination can be distinguished at low power. The subpial granular cell layer is no longer present, and the cortex is readily demarcated from the future subcortical white matter, even in the absence of myelination. (c) The future primary visual cortex at 22 weeks (same case as a) also shows a distinct subpial granular cell layer. The arrowhead here indicates a large neuron of Cajal-Retzius; such neurons are present over the entire neocortex. The cortical plate, however, shows at low-power early lamination in a hypocellular area (asterisk). (d) By 40 weeks, the cytoarchitecture of the primary visual cortex is discernible

308 Fig. 29.14 Hippocampal development. All three sections taken in coronal plane at the level of lateral geniculate nucleus (LGN). The developing cytoarchitecture of the hippocampal formation is perhaps most easily demonstrated in the posterior portion, where the seahorse morphology is most obvious. (a) At 15 weeks, the alveus (a) is already a distinct fiber bundle, and the fascia dentata (fd) and cornu ammonis (CA) are readily discerned. However, the future entorhinal cortex (EC) is still thin, and the temporal horn of the ventricle capacious (asterisk) and the hippocampus are still somewhat vertically oriented. (b) By 22 weeks, the entorhinal cortex is increasing in thickness, so the hippocampal formation is beginning to assume a more horizontal orientation, similar to that seen at term (c). CA1–CA4 subfields of the cornu ammonis, S subiculum

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Fig. 29.15 (a) Papillae of Retzius. The distortion of the marginal zone can be very difficult to classify as artifact on the basis of local histology alone. A careful search will identify an adjacent cleft in the marginal zone in many cases. (b) Cerebellar granular cell defects. These are frequent findings in term infants and may be eye-catching. Note the asterisk where small leptomeningeal blood vessels carry the same shape as the subjacent defects (arrowheads) in the granular cell layer. (c) Nodular excrescences. The herniation of a small blood vessel (arrowhead) with the germinal elements into the ventricle is nearly constant

when sought. An excrescence may contain occasional pigmented macrophages. (c) Nodular excrescences. The herniation of a small blood vessel and the germinal elements into the ventricle are nearly constant when sought. An excrescence may contain occasional pigmented macrophages. (d) Cerebellar dysplasia: isolated foci of disorganized cortical architecture are common, especially in the flocculus and nodulus. Similar foci can be found near the dentate, with large neurons intermixed with primitive germinal tissue

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References 1. Mai JK, Paxinos G, editors. The human nervous system. 3rd ed. Oxford: Academic Press; 2011. 2. O’Rahilly R, Müller F.  The embryonic human brain. An atlas of developmental stages. 3rd ed. Hoboken: Wiley; 2006. 3. Feess-Higgins A, Larroche JC.  Development of the human fetal brain: an anatomical atlas. Paris: Masson Editor; 1988. 4. Hartfuss E, Galli R, Heins N, Götz M.  Characterization of CNS precursor subtypes and radial glia. Dev Biol. 2001;229:15–30. 5. Brunne B, Zhao S, Derouiche A, Herz J, May P, Frotscher M, Bock HH.  Origin, maturation, and astroglial transformation of secondary radial glial cells in the developing dentate gyrus. Glia. 2010;58:1553–69. 6. Hansen DV, Lui JH, Parker PR, Kriegstein AR. Neurogenic radial glia in the outer subventricular zone of human neocortex. Nature. 2010;464:554–6. 7. Ortega JA, Radonjić NV, Zecevic N.  Sonic hedgehog promotes generation and maintenance of human forebrain Olig2 progenitors. Front Cell Neurosci. 2013;7:254. 8. Takashima S, Becker LE. Developmental changes of glial fibrillary acidic protein in cerebral white matter. Arch Neurol. 1983;40:14–8. 9. Del Bigio MR.  Cell proliferation in human ganglionic eminence and suppression after prematurity-associated haemorrhage. Brain. 2011;134:1344–61. 10. Freide RL.  Developmental neuropathology. 2nd ed. Berlin/ Heidelberg: Springer Verlag; 1989. 11. Hasegawa M, Houdou S, Mito T, Takashima S, Asanuma K, Ohno T. Development of myelination in the human fetal and infant cerebrum: a myelin basic protein immunohistochemical study. Brain Dev. 1992;14:1–6.

P. Shannon 12. Yakovlev PL, Lecours AR.  The myelogenetic cycles of regional maturation of the brain. In: Minkowski A, editor. Regional development of the brain in early life. Oxford: Blackwell; 1967. p. 3–70. 13. Brody BA, Kinney HC, Kloman AS, Gilles FH. Sequence of central nervous system myelination in human infancy. I. An autopsy study of myelination. J Neuropathol Exp Neurol. 1987;46:283–301. 14. Nadarajah B, Parnavelas JG. Modes of neuronal migration in the developing cerebral cortex. Nat Rev Neurosci. 2002;3:423–32. 15. Tabata H, Nakajima K.  Multipolar migration: the third mode of radial neuronal migration in the developing cerebral cortex. J Neurosci. 2003;23:9996–10001. 16. Bystron I, Blakemore C, Rakic P.  Development of the human cerebral cortex: Boulder Committee revisited. Nat Rev Neurosci. 2008;9:110–22. 17. Rakic P, Sidman RL.  Histogenesis of cortical layers in human cerebellum, particularly the lamina dissecans. J Comp Neurol. 1970;139:473–500. 18. Saito T, Hanai S, Takashima S, Nakagawa E, Okazaki S, Inoue T, et al. Neocortical layer formation of human developing brains and lissencephalies: consideration of layer-specific marker expression. Cereb Cortex. 2011;21:588–96. 19. Altman J, Bayer SA. Regional differences in the stratified transitional field and the honeycomb matrix of the developing human cerebral cortex. J Neurocytol. 2002;31:613–32. 20. Streeter GL. The cortex of the brain in the human embryo during the fourth month with special reference to the so-called “Papillae of Retzius”. Am J Anat. 1907;7(2):337–44. 21. Gelpi E, Budka H, Preusser M.  External granular cell layer bobbling: a distinct histomorphological feature of the developing human cerebellum. Clin Neuropathol. 2013;32:42–50.

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Eye Paul J. Bryar, David Gu, Samantha Agron, and Sarah E. Eichinger

Introduction A fully developed eye has three layers: the outer layer is composed of the cornea and sclera; the middle layer is the uvea, which comprises the iris, ciliary body, and choroid; and the inner layer is the retina. The lens is suspended from the ciliary body by numerous zonules. Aqueous is a liquid substance that fills the anterior part of the eye, and the vitreous is a gelatinous substance that fills the posterior part of the eye. Conjunctiva is a membranous structure that covers the anterior sclera and posterior surface of the eyelids. Six extraocular muscles attach to each eye, and the optic nerve (along with its vasculature) exits through an opening in the posterior eye wall. The first signs of development of the eye are apparent by thickening of neural folds of the future diencephalon, which then indent to form a pair of optic sulci. By about 32 days, the sulci expand and form the optic vesicles. The vesicles invaginate to form the optic cup, which is attached to the forebrain by an optic stalk. The inner wall of the optic cup develops into the neural retina, while the outer wall becomes the retinal pigment epithelium. The various cells of the neural retina will develop from week six through the eighth month. The surface ectoderm overlying the optic cup will thicken and form a lens placode. This area will invaginate and by day 33 forms the lens vesicle. Primary lens fibers form, and by the seventh week, the lens body is formed, and secondary lens fibers begin to develop. During weeks 6 and 7, the mesenchymal covering of the optic cup will differentiate into an P. J. Bryar (*) Departments of Ophthalmology and Pathology, Northwestern University Feinberg School of Medicine, Chicago, IL, USA e-mail: [email protected] D. Gu · S. Agron · S. E. Eichinger Department of Ophthalmology, Northwestern University Feinberg School of Medicine, Chicago, IL, USA e-mail: [email protected]; [email protected]; [email protected]

inner and outer layer. The inner layer will differentiate into the choroid, and the outer fibrous layer will become the sclera. Development of the cornea involves the surface ectoderm and mesenchyme from the neural crest. The corneal epithelium arises from the surface ectoderm, which is anterior to the lens. The outer fibrous layer of the mesenchyme of the neural crest will develop into the corneal stroma and endothelium. The three components of the middle uveal layer of the eye are the iris, ciliary body, and choroid. The choroid develops during the sixth and seventh weeks; it is derived from the inner vascular layer of the mesenchyme of the neural crest. The ciliary body has two developmental branches. The ciliary epithelium arises from the neuroectoderm, followed by the ciliary muscles, which develop from the neural crest-­ derived mesenchyme. The iris will develop from the neuroectoderm and periocular mesenchyme. The optic nerve and the vascular supply of the eye develop from the optic stalk, which is a narrow area between the optic cups and the forebrain. The axons from the retinal ganglion cells extend into the optic stalk to form the optic nerves. Myelination of the axons as they exit the eye begins during fetal development and is complete by the tenth postnatal week. Formation of the eyelids begins during the sixth week, when folds in the ectoderm above and below the eye appear. The epithelial surface of the lids fuses together. This will form the conjunctival sac, from which the conjunctiva will form and eventually cover the anterior sclera and posterior surface of the eyelids. During this time, the extraocular muscles develop from the paraxial mesoderm. By the end of the sixth month, the lids separate; they are capable of blinking by the eighth month. Development of each part of the eye may involve one or several different embryologic layers; development of various parts often occurs simultaneously. The histology of e­ mbryologic development of the eye is best understood by looking at the individual structures of the eye separately [1–3].

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Cornea and Sclera Embryology Development first begins with the formation of the corneal epithelium. The corneal epithelium is distinct from the other layers of the cornea in that it is derived from the surface ectoderm. Every other layer originates from mesenchymal cells derived from the neural crest. When the surface ectoderm first begins secreting collagen, it recruits mesenchymal cells to form the foundational stroma. The primitive epithelium is first distinguishable at approximately week 5. Junctional complexes between epithelial cells begin developing by week 6. When the eyelids open during week 24, the primitive epithelium begins to transition into a fully mature epithelium through cell layer stratification, reaching up to seven layers thick. The neural crest-derived mesenchyme migrates to the space between the corneal epithelium and the lens vesicle during week 7. By week 8, keratocytes begin building collagen lamellae to form the corneal stroma and Bowman’s layer [4]. The corneal stroma at birth is much less cellular than developing stroma. Development of Descemet’s membrane begins during week 8, when corneal endothelial cells secrete collagen lamellae that consist primarily of laminin and collagen types IV and VIII. The first layer formed during this period is the anterior banded layer and a thin, nonbanded layer adjacent to the stroma. The posterior nonbanded zone is only secreted by endothelial cells after birth and continues to thicken with age. At birth, Descemet’s membrane is approximately 3  μm thick; it increases to up to 10  μm by adulthood, mostly owing to the creation of the posterior nonbanded layer [5]. The corneal endothelium at birth has a monolayer of numerous endothelial cells. This cell density decreases over time through corneal growth and loss of individual cells. In response, each remaining endothelial cell grows in size to occupy the newly created space left behind by each lost cell [6, 7]. Scleral formation begins during the fifth week of development, as fibroblasts embedded in the connective tissue matrix continue to form the fibrous coat. By week 6, the sclera differentiates from the neural crest cells that surround the optic cup and divides into an anterior and posterior portion, each with differing cell types. Anterior cells contain more developed endoplasmic reticulum and Golgi complexes, whereas posterior cells contain more ribosomes [8]. Elastin fibers begin proliferation by approximately the ninth week. At this time, the posterior and anterior cells reconverge to resemble nearly identical cells, the sole difference being increased collagen in the anterior portion. By week 12, collagen growth reaches the posterior pole and the optic nerve. At week 16, the fetal sclera still is highly cellular when compared with the dense collagen bundles of adult sclera. During this period, until approximately week 20, the posterior portion of

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the sclera separates into an inner and outer layer. The outer layer joins with the optic nerve sheath, and the inner layer begins the formation of a primitive lamina cribrosa. The posterior scleral foramen fully matures by month 7. Fibrous tissue from the choroid and optic nerve contributes to the formation of the lamina cribrosa shortly before birth. The collagen makeup of the sclera continues to change throughout fetal development and even after birth. As a result, the once malleable and expansile sclera found in infants becomes more rigid by adulthood [8, 9].

Histology Together, the cornea and sclera form the outermost coat of the eye. The fully mature adult cornea can be divided into five distinct structures: the corneal epithelium, Bowman’s layer, corneal stroma, Descemet’s membrane, and corneal endothelium (Fig.  30.1). The corneal epithelium contains three distinct epithelial cells: superficial squamous cells, wing cells, and columnar basal cells (Fig. 30.2). Bowman’s layer is an acellular layer approximately 15 μm thick, found beneath the epithelial basement membrane. It is the anteriormost part of the corneal stroma and is visible on routine H&E staining because of its organization of collagenous lamellae. Bowman’s layer consists mainly of type I collagen fibrils, which serve as protection for the stroma. Beneath Bowman’s layer is the corneal stroma, which comprises more than 90% of the corneal thickness. The stroma is cellular during development, but it becomes much less cellular by adulthood (Fig.  30.3). Routine histologic processing of the cornea causes artifactual clefting of the collagenous stroma. Absence of clefting can be the result of corneal edema or scarring. Descemet’s membrane is the basement

Fig. 30.1  Cornea at term. The developed cornea has five distinct layers, from anterior to posterior: nonkeratinized squamous corneal epithelium (Ep); Bowman’s layer (BL); corneal stroma (St), which constitutes over 90% of the cornea; Descemet’s membrane (DM), which is the basement membrane of the corneal endothelium; and the corneal endothelium (En) (H&E, 10×)

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Fig. 30.2  Corneal epithelium 2 weeks postnatal. The mature corneal epithelium contains three distinct cell layers: the outermost squamous cell layer, the intermediate wing cells, and the deeper basal cells. Beneath the basal cells is the epithelial basement membrane. Posterior to the epithelium is the acellular Bowman’s layer (BL), the anteriormost part of the stroma (St). Though it is sometimes called Bowman’s membrane, Bowman’s is not a true membrane (unlike Descemet’s membrane); it is a part of the stroma with collagen arranged differently than the remainder of the stroma, allowing it to be readily identified on H&E stain (H&E, 40×)

Fig. 30.4  Posterior cornea at term. The stroma (St) has elongated keratocyte nuclei. The clefts are an artifact seen in all normal corneas that have undergone routine histology processing. Corneal endothelium (En) at this time consists of numerous round nuclei (H&E, 40×). DM Descemet’s membrane

membrane of the corneal endothelium and is posterior to the corneal stroma. It consists primarily of an anterior banded layer and a posterior nonbanded layer. The corneal endothelium makes up the posterior aspect of the cornea. It is comprised of a single cell layer that provides essential metabolic function to maintain the unique transparent characteristics of the cornea (Fig. 30.4). The sclera is the fibrous, protective outer layer of the eye, which consists primarily of collagen and elastic fibers. Although the cornea and sclera both contain extensive collagen layering, the cornea retains its transparent qualities because of the uniform diameter and structured positioning of the collagen, allowing for the passage and refraction of light [10, 11]. The sclera, by contrast, has wider collagen fibrils with a more interwoven structural makeup that result

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Fig. 30.3  Developing cornea at 21 weeks gestation. The corneal epithelium (Ep) at 21  weeks still has ill-defined cell layers. A primitive Bowman’s layer (BL) and Descemet’s membrane (DM) are forming by this period in development. Most notable is the corneal stroma (St). Much less cellular at birth, the stroma at 21 weeks contains numerous keratocytes that continue to produce collagen (H&E, 10×). En endothelium

Fig. 30.5  Sclera at term. The sclera (Sc) is highly collagenous, similar to the stroma of the cornea. However, the collagen structure of the sclera is interwoven, preventing the passage of light. The cornea, by contrast, has a highly organized collagen structure that forms a transparent layer (H&E, 20×). Ch choroid, RPE retinal pigment epithelium

in reduced passage of light as the infant continues to grow (Fig. 30.5). The lamina cribrosa is the sieve-like structure on the posterior of the eye that permits the passage of optic nerve fibers and associated vasculature.

Uvea Embryology The iris originates from the neuroectoderm and periocular mesenchyme and receives signaling from the lens. The lens

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has been shown to be essential to iris development, but the specific signaling molecules are not completely understood. Development of the iris and ciliary body begins with the formation of the optic cup, which arises from the mesenchyme upon interaction with the surface ectoderm. The non-neuronal, peripheral areas on the rim of the optic cup differentiate from the central, neuronal areas. At approximately day 45, the ciliary portion of the optic cup is still undeveloped and persists at the equator of the primitive lens [12]. The ciliary body at day 45 has two developmental branches, with the development of the ciliary epithelium from the neuroectoderm preceding the development of the ciliary muscle and stroma from the mesenchyme. Like the iridial muscles, the ciliary epithelium develops from the periphery of the optic cup. The inner, nonpigmented epithelium and the outer, pigmented layer of the ciliary epithelium are established when the optic vesicle folds inward to form the double-layered optic cup [13]. As the ciliary body matures, it begins folding onto itself to form the ciliary processes. The ciliary muscle and the stroma develop from neural crest-derived mesenchyme anterior to the ciliary epithelium. The first buddings of the ciliary muscle and stroma appear during the seventh week. The longitudinal, radial, and circular ciliary muscles begin developing simultaneously, with myofilaments visible by week 12 [14]. The first appearance of the zonular fibers can be found starting at week 13. They originate from the ciliary processes and will eventually attach to the periphery of the lens to secure the position of the lens inside the eye [15]. The inner layer of the optic cup gives rise to the posterior iris-pigmented epithelium (IPE) and the neural retina, while the outer layer gives rise to the anterior IPE and the retinal pigment epithelium (RPE). These optic cup margins extend to produce molecular markers specific to smooth muscle development, triggering the development of the two iridial muscles. The iris sphincter muscle begins developing at about 11 weeks. Iris dilator muscles develop later. Both are almost completely formed by approximately 32  weeks’ ­gestation. The iris dilator and sphincter muscles are the only muscles in the body to originate from the neuroectoderm [16–18]. After birth, the iris color darkens in some people as melanin gradually accumulates in the iris stroma. The development of the iris begins with the forward extension of the optic cup when invagination of the optic vesicle occurs. The first definitive features of the iris can be found during the fourth week with the early beginnings of vasculature that will eventually comprise the iris stroma. The iris stroma arises from the neural crest-derived mesenchyme, which migrates between the lens vesicle and the primitive corneal epithelium during the seventh week. The neural crest-derived mesenchyme also gives rise to the pupillary membrane during week 8. Fusion of the primitive iris occurs by approximately day 33. Failure to fuse the optic fissure

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Fig. 30.6  Anterior chamber at 21 weeks gestation. The iris is not fully developed, with myofilaments beginning to form (tip of arrow labeled iris stroma (IS)). At this point, the ciliary muscles have not yet developed. AC anterior chamber, CB ciliary body, Co cornea, CP ciliary process, Le lens, PIA primitive iridocorneal angle (H&E, 10×)

may result in an iris coloboma. At week 12, the first buddings of the ectodermal iris can be seen. By week 16, the primitive future iris and a primitive iridocorneal angle can be identified (Fig. 30.6) [18]. Complete development of the iris is driven by regulators such as PAX6. The formation of a definitive anterior chamber begins during week 7. The anterior chamber angle is formed by the junction of the cornea and iris and is the site of aqueous humor drainage. The trabecular meshwork originates from neural crest cells that migrate to the iridocorneal angle. During the fourth month of fetal development, the angle of the anterior chamber contains densely packed mesenchymal cells. Within the next month, these cells begin to separate, creating small columns of space known as trabecular beams. During this time, blood vessels begin extending from the sclera toward the chamber angle that eventually forms Schlemm’s canal. The migration of the iris root posteriorly creates the necessary space to facilitate drainage. The trabecular meshwork cells extend to the scleral spur (Fig. 30.7) [19–21]. The choroid begins to develop during the sixth and seventh weeks. The mesenchymal capsule, derived primarily from neural crest cells, differentiates into two layers: the inner vascular layer, which later becomes the choroid, and the outer fibrous layer, which develops into the sclera and the cornea. The choriocapillaris supplying the RPE and ­photoreceptors develops distinguishable arteries and veins by week 23. Eventually, these arteries will supply the choroid and the outer two-thirds of the retina. Formation of Bruch’s membrane begins during week 7 and continues growth with collagen expression from both the adjacent RPE and choriocapillaris (Fig. 30.8) [22, 23].

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Fig. 30.7  Anterior chamber 2 weeks postnatal. The developed anterior chamber (AC) is formed by the cornea (Co) and iris (Ir). The conventional outflow pathway of aqueous humor runs through the trabecular meshwork (TM) to Schlemm’s canal (SC). CB ciliary body, CP ciliary process, Sc sclera, SL Schwalbe’s line, SS scleral spur. (H&E, 10×)

Fig. 30.9  Iris 2 weeks postnatal, showing the mature iris with the notable structures. AC anterior chamber, IDM iris dilator muscle, ISM iris sphincter muscle, Le lens, PC posterior chamber, PE iris pigment epithelium, St stroma (H&E, 10×)

Histology The middle layer of the eye is called the uvea and consists of the iris, ciliary body, and choroid. The iris is the circular muscle layer between the cornea and the lens, which serves as the eye’s aperture, regulating the amount of light passing through the lens as it travels to the retina. The anterior stroma contains fibroblasts and melanocytes. The posterior iris contains the iris muscles: the sphincter pupillae (more centrally located) and the dilator pupillae (more peripherally located). These are the only muscles in the body that originate from the neuroectoderm [17]. The posterior surface of the iris is

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Fig. 30.8  Retina (Re), choroid (Ch), and sclera (Sc) at 21 weeks gestation. Note the numerous capillaries, in contrast to the collagen-heavy sclera. BM Bruch’s membrane, RPE retinal pigment epithelium (H&E, 20×)

lined by pigmented epithelium (closer to the iris stroma) and nonpigmented epithelium (closer to center of the eye). The IPE makes up the posterior border of the iris (Fig. 30.9). With the maturation of the iris and cornea comes the eventual creation of the iridocorneal angle (anterior chamber angle), where aqueous humor drains from the eye. The conventional drainage route involves movement of fluid through the trabecular meshwork by trabecular beams to Schlemm’s canal. From anterior to posterior, the structures of the anterior chamber angle are Schwalbe’s line, trabecular meshwork, scleral spur, and ciliary body. Schwalbe’s line is formed by the terminus of Descemet’s membrane of the cornea. The trabecular meshwork resides between Schwalbe’s line and the scleral spur, a small outcropping of the sclera. Just posterior to the trabecular meshwork lies the ciliary body (see Fig. 30.7). The anterior chamber is the space between the corneal endothelium and the anterior surface of the iris. The posterior chamber is the space that is bordered by the posterior surface of the iris, the peripheral lens, and the zonules (see Fig. 30.9). The ciliary body includes the ciliary muscle and the ciliary processes (Fig. 30.10). The ciliary muscles consist of an outer ring of longitudinal muscle, an intermediate layer of radial muscle, and an inner layer of circular muscle. Extending from the ciliary body toward the middle of the eye, at a section of the ciliary body called the pars plicata, are ciliary processes, which are long and fingerlike at birth but later become shorter and develop hyalinized cores by adulthood. The ciliary body and processes are lined by a dual layer of epithelium: an outer pigmented layer (closest to the ciliary body) and inner nonpigmented layer (closest to the vitreous cavity) that secretes aqueous humor into the posterior chamber. At the posteriormost section of the ciliary body, called the pars plana, the outer pigmented layer is contiguous with the retinal pigment epithelium, and the inner, nonpigmented epithelium is contiguous with the neurosensory retina.

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Retina and Optic Nerve Embryology The development of the retina begins around day 22 of embryogenesis with the appearance of two lateral grooves in the anterior neuroectoderm, called the optic sulci. The expression of eye-field transcription factors (EFTFs), such as the homeobox gene Rx, induces the formation of the optic sulci and the eventual differentiation of this portion of neuroepithelium into the various cell types seen in the mature neurosensory retina [21]. As the neural tube closes, the optic sulci expand laterally and evaginate, giving rise to the optic vesicles by day 25. The optic vesicles are then surrounded by mesenchymal cells that originate from the neural crest, Fig. 30.10  Ciliary processes at age 1 year. The ciliary processes (CP) while their apices come into contact with the surface ectoextend from the ciliary body (CB) toward the lens. They consist of an derm [22, 23]. outer nonpigmented epithelium and an inner pigmented epithelium. The distal portion of the optic vesicles will differentiate Ciliary processes at this age are long and fingerlike, which is one way into the neurosensory retina; the proximal optic vesicles will to histologically differentiate a globe from a young child versus one develop into the retinal pigment epithelium (RPE) and optic from an adult. Co cornea, Ir iris (H&E, 4×; inset, 40×) nerves. A gradient of Sonic Hedgehog (Shh) signaling is critical to the process of optic vesicle formation and eventual development of the eyes [21]. The development of the central and peripheral portions of the neurosensory retina proceed along distinct paths, with the central neural retina developing from the posterior optic vesicle and the peripheral retina arising from the distal tip of the optic vesicle. The RPE develops from the dorsal optic vesicle. By day 28 after fertilization, the RPE produces melanin, at which point the cilia of the RPE also recede and the cells begin to change in shape from columnar to cuboidal. Proliferation of the RPE, which forms a monolayer at the outermost retina, is completed before birth. The RPE expands during the postnatal period by hypertrophy of existing cells [22]. A basement membrane lines the outside of the optic vesicle, with the anterior or distal portion eventually forming the retinal internal limiting membrane. As the distal optic vesiFig. 30.11  Sclera (Sc), choroid (Ch), and retinal pigment epithelium cles invaginate, the two layers of the optic cups that will (RPE) at term. Bruch’s membrane (BM) lies beneath the RPE. It is a become the neural retina appear. The cavity between these multilayered structure that consists of collagen and elastin (H&E, 40×) two layers forms an intraretinal space that will disappear as the retina grows and becomes differentiated. As the optic cup The choroid is a pigmented vascular layer located between forms, the embryonic (choroid) fissure of the optic stalk, the retina and sclera (Fig. 30.11). The innermost structure of where the hyaloid artery runs, also appears. During the first the choroid is Bruch’s membrane, a multilayered membrane 6 weeks of development, retinal cells proliferate but remain consisting of collagen surrounding an inner layer of elastin. undifferentiated retinoblasts [22, 24]. The proximal optic vesicles narrow to become the optic Closest to the Bruch’s membrane are fenestrated capillaries known as the choriocapillaris, which supply the RPE and stalks between the optic cups and the forebrain, forming the photoreceptors. Eventually, the choriocapillaris will supply precursor to the optic nerves. The optic stalks secrete fibrothe rest of the choroid and the outer aspects of the retina. The blast growth factor (FGF), which induces the expression of second layer, known as Sattler’s layer, consists of medium-­ the transcription factor atoh7, a key regulator of retinal gansized vessels. The final blood vessel layer, Haller’s layer, glion cell (RGC) differentiation [25]. The axons of ganglion contains the largest vessels and is located closest to the cells grow into the optic stalks, forming the optic nerves by week 6 of development. The glial cells within the optic nerve sclera.

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Fig. 30.12  Optic nerve at 20 weeks gestation. The axons of the retinal ganglion cells begin to grow within the optic stalk by week 6 of development, forming the optic nerve. In the intrascleral portion of the optic nerve, these axons pass through the mesh-like lamina cribrosa (LC). Myelination of the extrascleral portion of the optic nerve is completed during the tenth postnatal week. Ch choroid, NFL nerve fiber layer, Sc sclera (H&E, 10×)

derive from the inner lining of the optic stalk; the outer epithelium of the outer stalk forms the peripheral glial mantle separating the nerve fibers from the adjacent mesenchyme. The optic nerves are encased in three layers of tissue, which are continuous with the meninges (dura, arachnoid, and pia mater) and terminate at the sclera. The meninges are derived from mesectodermal cells that migrate from the neural crest during early development. Myelination of the optic nerves begins with the central nerve fibers and proceeds outward toward the eye up to the lamina cribrosa of the sclera (Figs.  30.12 and 30.13). The nerve fibers within the retina and the proximal optic nerve remain unmyelinated. Although myelination of the axons of the optic nerves begins during fetal development, the process is not complete until the tenth postnatal week [22]. Around the time that the embryonic fissure closes during week 6, retinal cells begin to differentiate. Differentiation of progenitor cells into the various retinal cell types is regulated by a complex array of transcription factors and extracellular signals acting in various temporal and spatial patterns that are still being elucidated. Rx and other homeobox genes influence the differentiation of photoreceptors into rods and cones. The ratio and distribution of rods and cones is related to expression gradients of various extracellular signaling molecules, including Shh, Wingless-related integration site (Wnt), FGFs, basic helix-loop-helix (bHLH), and RA [22]. During the fourth month, the macula begins to develop, as ganglion cells proliferate near the center of the retina, resulting in a layer of RGCs that is eight to nine cells deep by the

Fig. 30.13  Optic nerve at 17 weeks gestation. A portion of the hyaloid artery (asterisk) is seen above the optic nerve head. At this stage in development, the retina (not shown) is hypercellular compared with its appearance at birth. RPE retinal pigment epithelium, Sc sclera (H&E, 10×)

sixth month. Cones predominate in the thickened outer nuclear layer (ONL) in this region. The foveal depression forms during the seventh month as ganglion cells begin to be displaced from this region. By the eighth month, the RGC layer in this region is two cells thick. In addition, the shape of the cone cells changes. The outer segments lengthen and the inner segments narrow, allowing for a higher density of photoreceptor cells. The high resolution of the fovea arises from the roughly 1:1 ratio of synaptic connections between cones and RGCs in this region. As the peripheral retina grows, it forms a junction with the ciliary body, called the ora serrata, by the sixth month [22, 24]. The early retina is divided into an outer nuclear zone, adjacent to the RPE, and an acellular marginal zone, in contact with the inner limiting membrane. The mitotically active outer nuclear zone is derived from the proliferative neuroepithelium of the neural tube. As cells from the outer nuclear zone migrate into the marginal zone during week 7 of development, an inner neuroblastic layer (INBL) is formed (the outer nuclear zone is then termed the outer neuroblastic layer, ONBL). Ganglion, amacrine, and Müller cells p­ opulate

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Fig. 30.14  Retina and choroid at 21  weeks gestation. The choroid forms from the mesenchyme surrounding the optic cup during weeks 6–7 of development. The layers of the retina appear distinct at this point in development, when photoreceptor outer segments and synapses are forming (H&E, 20×)

the INBL, which gives rise to the ganglion cell layer (GCL) and inner nuclear layer (INL) of the mature retina. Although RGCs are among the first retinal cells to differentiate, their development is not complete until the eighth month. The ONBL gives rise to the photoreceptors, which will form the ONL, as well as the bipolar and horizontal cells, which migrate to the INL.  During weeks 10–12 of development, microglia enter the retina from the vasculature and peripheral subretinal space [22, 26]. As retinal precursor cells become differentiated, the synaptic connections between the various cell types in this network begin to take shape. Synaptogenesis begins in rods during the fourth month of development, and in cones during the fifth month; development of the outer segments of both types of photoreceptors also begins during the fifth month (Fig. 30.14). The transient layer of Chievitz, which separates the ONBL and INBL, is replaced during the third month by the inner plexiform layer (IPL), which contains the synapses between RGCs and cells of the INL.  Horizontal cells, ­interneurons of the INL that synapse with multiple photoreceptors, differentiate during the fifth month [22]. The development of the retinal vasculature begins with the hyaloid artery, a branch of the ophthalmic artery that supplies the optic stalk, growing between the lens and the marginal zone of the early retina. Angiogenesis from the hyaloid artery gives rise to the central retinal artery and the vessels that supply the peripheral retina, extending to the ora serrata by the eighth month. The outer retinal plexus arises during the ninth month from vascular development that extends through the retinal layers to the ONL. The central retinal artery eventually supplies the inner one third of the mature retina, while the choroidal vasculature (choriocapillaris) supplies the outer two thirds (Fig.  30.15). The hyaloid arteries regress as the retinal circulation is estab-

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Fig. 30.15  Retina and choroid at 20 weeks gestation. RPE retinal pigment epithelium (H&E, 40×)

lished, occasionally leaving Bergmeister’s papillae on the optic nerve head or Mittendorf’s dots on the posterior lens capsule as remnants. Bergmeister’s papillae are outcroppings of fibrous tissue at the center of the optic discs; Mittendorf’s dots are circular opacities on the posterior lens. Regression of the hyaloid arteries during the fifth to seventh weeks is also accompanied by the closure of the choroid fissure. One cause of colobomas is the persistence of the choroid fissure [26].

Histology The mature retina is a multilayered sensory structure of the central nervous system located in the posterior segment of the eye that transduces light stimuli (Fig. 30.16). Closest to the vitreous is the inner limiting membrane, beneath which lies the nerve fiber layer (NFL) containing the axons of RGCs, which form the optic nerves. Once the nerve exits the globe, it becomes myelinated and is encased in the meninges (dura, arachnoid, and pia mater), which can be appreciated on histologic section (Fig. 30.17). The diameter of the optic nerve doubles as it travels through the sclera and becomes myelinated. It is covered externally by the dura and arachnoid. Pial septae are seen within the optic nerve. The RGC axons pass through the mesh-like lamina cribrosa of the sclera. The next retinal layer contains the cell bodies of the RGCs. The cell bodies of the bipolar cells, along with the amacrine and horizontal cells, are contained in the INL. The bipolar cells synapse with the photoreceptors in the OPL. The cell bodies of the photoreceptors (rods and cones) are located in the ONL. The ratio of rods to cones varies according to location in the retina, with the fovea containing mostly cones. Rods are more numerous in the periphery of the ret-

30 Eye

Fig. 30.16  Retina at term. Retinal cells begin to differentiate by the sixth week of development and the process continues until birth, forming the mature, multilayered retina: RPE retinal pigment epithelium, PR photoreceptors, ONL outer nuclear layer, OPL outer plexiform layer, INL inner nuclear layer, IPL inner plexiform layer, RGC retinal ganglion cells, NFL nerve fiber layer. Visible in the photoreceptor layer are cone cells (asterisk) and rod cells (arrows) (H&E, 40×)

319

Fig. 30.17  Optic nerve cross section at term. The optic nerve is encased in the meninges (dura, arachnoid, and pia), which terminate at the sclera. The arachnoid (Ar) and dura are on the outside of the optic nerve, while the pia forms septae (PS) within the optic nerve (H&E, 40×)

Lens and Vitreous Embryology

Fig. 30.18  Retinal pigment epithelium (RPE), choroid (Ch), and sclera (Sc) at term. Both the RPE and the choroidal melanocytes (ChM) contain melanin granules (H&E, 40×)

ina. Closest to the choroid lies the cuboidal retinal p­ igment epithelium (RPE) (Fig. 30.18). During embryonic and fetal development, the retina primordia undergoes a complex process of differentiation, resulting in the formation of these distinct layers of neurons, glial cells, and pigment cells. During routine histologic processing and fixation, the neurosensory retina above the ora serrata may become detached. This artifact, which is seen only in infants and young children, is called Lange’s fold; it does not represent a true retinal detachment (Fig. 30.19).

The lens placode, a thickening of surface ectoderm overlying the anterior wall of the nascent optic vesicle, forms during the third week of development and is regulated by the expression of SOX2 and Pou2f1 transcription factors [27]. By approximately day 22–23, cells from the lens placode begin to form a depression called the “lens pit.” As the lens placode further invaginates, it detaches from the surface ectoderm (day 33), forming a lens vesicle that lies in close apposition to the optic vesicle/cup [28]. At this stage, the lens vesicle is an epithelial monolayer bound by a basal lamina. The spatial relationship between the lens vesicle and the optic vesicle/cup has been shown to play a significant role in lens d­ evelopment [29]. Crystallins, the most abundant proteins in the lens, are first produced by the lens placode, but their transcriptional levels increase dramatically with the differentiation of the primary lens fibers. These proteins have a variety of roles, not only contributing to the transparency and refractive power of the lens, but also acting as molecular chaperones and mitigating the formation of cataracts. While β- and γ-crystallins in the lens are largely structural proteins, α-crystallins have diverse functions including the regulation of apoptosis and chaperone-mediated autophagy. Mutations in genes encoding crystallins are one of the causes of congenital cataracts [30, 31]. During week 5, the primary vitreous occupies the retrolental space, which also contains the hyaloid vasculature. The secondary (definitive) vitreous is then deposited behind the primary vitreous until week 12. At this time, fibers

320 Fig. 30.19 (a) Lange’s fold. The gross photograph of an eye from a 1-year-old shows an artifactual circumferential fold (Lange’s fold) in the peripheral retina (black arrow). The posterior surface of the lens has a concave dimple artifact commonly seen in enucleated infant globes (arrowhead). There is an unrelated peripheral retinal hemorrhage on the left. (b) H&E stain shows the artifactual Lange’s fold (H&E 10×)

P. J. Bryar et al.

a

b

c­ onnecting the secondary vitreous to the inner limiting membrane of the retina appear. Fibers called tertiary vitreous form between the optic cup and the lens equator during the third month. These fibers adhere to the inner limiting membrane of the retina. During the fourth month of development, the primary vitreous and the surrounding hyaloid vasculature atrophy as the retinal vasculature forms. Hyaluronan and type II collagen are largely added to the vitreous after birth. When the primary vitreous fails to regress, the result is either anterior or posterior persistent hyperplastic primary vitreous (PHPV), conditions that affect vision. Anterior PHPV results in a white retrolental mass that may be mistaken for retinoblastoma, whereas posterior PHPV, which damages the retina, is associated with alterations in Wnt signaling [32].

Histology Three histologically relevant structures comprise the mature lens: the capsule, the epithelium, and the lens substance (cortex and nucleus) (Figs. 30.20 and 30.21). The lens substance consists of the lens fibers, which grow throughout life. The lens lacks nerves and blood vessels. Nutrients and various molecular signals reach the lens by diffusing through the vitreous and anterior chamber fluid. After the lens vesicle forms, lens cells begin to differentiate under the regulation of FGF and other factors. The cells of the anterior lens vesicle remain a monolayer of cuboidal epithelium (Fig.  30.22). Appositional growth of the basal lamina toward the surface ectoderm gives rise to the lens capsule. The cells of the posterior lens vesicle elongate into

Fig. 30.20  Lens at 21 weeks gestation. At this stage in development, the lens is spherical, but at birth it will be biconvex in shape. The lens has an anterior capsule with a layer of cuboidal epithelium (AC-Ep) and a thinner acellular posterior capsule (PC). The older lens fibers lie in the inner lens nucleus (Nu), while new fibers are added to the outer cortex. Eq equator (H&E, 2×)

the lumen and differentiate into the primary lens fibers. By the end of the fourth week, the primary lens fibers will grow to completely fill the lumen, forming the solid primary embryonic lens nucleus. The secondary lens fibers arise from cells of the anterior lens epithelium that migrate equatorially, divide, and differentiate into lens fibers that encircle the primary lens fibers. This forms the fetal lens nucleus. During the process of lens fiber maturation, the nuclei and other organelles of these cells disappear. New lens fibers are continually generated throughout life in this manner (Fig. 30.23).

30 Eye

Fig. 30.21  Lens and zonules during postnatal month 15. The mature lens has a biconvex shape and is suspended by the zonules (Zo), which primarily emanate from the pars plicata. The zonules attach to the lens at the lens equator and anterior and posterior to it. AC anterior chamber, CB ciliary body, Le Co lens cortex, Le Nu lens nucleus, PC posterior chamber, PP/CP pars plicata/ciliary processes, Sc sclera (H&E, 2×)

Fig. 30.23  Lens fibers at 20  weeks gestation. As the elongated lens fibers mature, they lose their organelles. This lens was displaced during processing, so the ciliary processes are artifactually on the anterior aspect of the lens (H&E, 20×)

As the secondary lens fibers grow around the apices of the primary lens fibers during the second month and the lens loses its spherical shape, the lens sutures are formed. The early primary lens fibers span the lens from the anterior to the posterior pole, resulting in the spherical structure of the lens. However, the later secondary fibers elongate unequally toward the anterior and posterior poles from their equatorial origin. The meeting of lens fibers originating from opposite

321

Fig. 30.22  Anterior lens capsule at term. The anterior lens capsule (AC) has a layer of cuboidal epithelium (Ep). Le Fib lens fibers (H&E, 20×)

sides of the lens results in the formation of lens sutures, which are held together by ground substance. The initial sutures are Y-shaped: one upright in the anterior lens and one upside down in the posterior lens. However, continued lens fiber growth results in the appearance of more intricate suture patterns as development continues. This pattern of fiber growth contributes to the flattened, biconvex shape of the lens. By 9  weeks, the lens is surrounded by a vascular ­structure called the tunica vasculosa lentis, which receives its blood supply from the hyaloid artery [3, 33, 34]. In utero, the lens is more spherical in shape, but as the fetus develops, equatorial growth occurs, so that by birth, it has its biconvex shape. The equatorial diameter of the lens at birth is approximately 6.5  mm; it grows to approximately 9  mm. The anterior-to-posterior thickness of the lens is approximately 3.5  mm at birth; it eventually enlarges to about 5 mm. It is surrounded by a capsule, composed mostly of type IV collagen, and is suspended in the eye by zonules that extend primarily from the pars plicata and attach to the lens equator [24] (see Fig. 30.21). Anteriorly, just beneath the lens capsule is a monolayer of cuboidal lens epithelial cells (see Fig. 30.22). The mitotically active lens epithelial cells are in the periphery of the anterior lens capsule, called the germinative zone (Fig.  30.24). Posteriorly, the lens is covered by an acellular lens capsule (Fig. 30.25). The lens capsule is thickest in the post-­equatorial regions on the anterior and posterior surfaces of the lens. At birth, the central anterior capsule is thicker than the central posterior capsule (the thinnest region of the lens capsule). On histology, the anterior vitreous base can be appreciated just posterior to the posterior lens capsule (see Fig. 30.25).

322

Fig. 30.24  Posterior lens during postnatal month 15. The lens epithelium (Ep) does not extend to the posterior lens capsule (PLC). Le Nuc lens nucleus (H&E, 10×)

References 1. Sowden JC.  Development of the eye. In: Standring S, editor. Grays anatomy: the anatomical basis of clinical practice. 41st ed. Amsterdam: Elsevier Limited; 2016. p. 661–5. 2. Ovalle WK, Nahirney PC.  Eye and adnexa. In: Netters essential histology. Philadelphia: Elsevier/Saunders; 2013. p. 431–51. 3. Schoenwolf GC, Bleyl SB, Brauer PR, Francis-West PH. Larsen’s human embryology. 4th ed. Philadelphia: Elsevier/Churchill Livingstone; 2008. p. 488–500. 4. Secker GA, Daniels JT.  Limbal epithelial stem cells of the cornea. 2009 Jun 30. In: StemBook [Internet]. Cambridge, MA: Harvard Stem Cell Institute; 2008. Available from: https://www. ncbi.nlm.nih.gov/books/NBK27054/. https://doi.org/10.3824/ stembook.1.48.1. 5. Johnson DH, Bourne WM, Campbell RJ.  The ultrastructure of Descemet’s membrane. I.  Changes with age in normal corneas. Arch Ophthalmol. 1982;100:1942–7. 6. Bahn CF, Glassman RM, MacCallum DK, Lillie JH, Meyer RF, Robinson BJ, Rich NM. Postnatal development of corneal endothelium. Invest Ophthalmol Vis Sci. 1986;27:44–51. 7. Wörner CH, Olguín A, Ruíz-García JL, Garzón-Jiménez N.  Cell pattern in adult human corneal endothelium. PLoS One. 2011;6:e19483. 8. Sainz de la Maza M, Tauber J, Foster CS. The sclera. New York: Springer Science & Business Media; 2012. 9. Sellheyer K, Spitznas M.  Development of the human sclera. A morphological study. Graefes Arch Clin Exp Ophthalmol. 1988;226:89–100. 10. Bailey AJ.  Structure, function and ageing of the collagens of the eye. Eye (Lond). 1987;1:175–83. 11. Hassell JR, Birk DE. The molecular basis of corneal transparency. Exp Eye Res. 2010;91:326–35. 12. Davis-Silberman N, Ashery-Padan R.  Iris development in ver tebrates; genetic and molecular considerations. Brain Res. 2008;1192:17–28. 13. Beebe DC. Development of the ciliary body: a brief review. Trans Ophthalmol Soc U K. 1986;105:123–30. 14. Peces-Peña MD, de la Cuadra-Blanco C, Vicente A, Mérida-Velasco JR. Development of the ciliary body: morphological changes in the

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Fig. 30.25  Posterior lens at term. Posterior to the lens lies the vitreous. AVB anterior vitreous base, PLC posterior lens capsule (H&E, 20×)

distal portion of the optic cup in the human. Cells Tissues Organs. 2013;198:149–59. 15. Shi Y, Tu Y, De Maria A, Mecham RP, Bassnett S. Development, composition, and structural arrangements of the ciliary zonule of the mouse. Invest Ophthalmol Vis Sci. 2013;54:2504–15. 16. Qi Y, Li FM. The embryonic development of iridial muscles [Article in Chinese]. Zhonghua Yan Ke Za Zhi. 1991;27:295–8. 17. Link BA, Nishi R. Development of the avian iris and ciliary body: mechanisms of cellular differentiation during the smooth-to-­ striated muscle transition. Dev Biol. 1998;203:163–76. 18. Mann IC.  The development of the human iris. Br J Ophthalmol. 1925;9:495–512. 19. Reis LM, Semina EV. Genetics of anterior segment dysgenesis disorders. Curr Opin Ophthalmol. 2011;22:314–24. 20. Gage PJ, Zacharias AL. Signaling “cross-talk” is integrated by transcription factors in the development of the anterior segment in the eye. Dev Dyn. 2009;238:2149–62. 21. Stenkamp DL.  Development of the vertebrate eye and retina. In: Hejtmancik JF, Nickerson JM, editors. Progress in molecular biology and translational science. London: Elsevier/Academic Press; 2015. p. 397–414. 22. Forrester JV, Dick AD, McMenamin PG, Roberts F, Pearlman E. Embryology and early development of the eye and adnexa. In: The eye: basic sciences in practice. 4th ed. Edinburg: Elsevier; 2016. p. 103–29. 23. Fuhrmann S. Eye morphogenesis and patterning of the optic vesicle. Curr Top Dev Biol. 2010;93:61–84. 24. Jakobiec FA.  Ocular anatomy, embryology and teratology. Philadelphia: Harpercollins; 1982. 25. Vecino E, Rodriguez FD, Ruzafa N, Pereiro X, Sharma SC. Glia-­ neuron interactions in the mammalian retina. Prog Retin Eye Res. 2016;51:1–40. 26. Hendrickson AE, Yuodelis C. The morphological development of the human fovea. Ophthalmology. 1984;91:603–12. 27. Donner AL, Episkopou V, Maas RL.  Sox2 and Pou2f1 inter act to control lens and olfactory placode development. Dev Biol. 2007;303:784–99. 28. Apple DJ, Rabb MF.  Ocular pathology: clinical applications and self-assessment. 5th ed. St. Louis: Mosby; 1997. 29. Swindell EC, Liu C, Shah R, Smith AN, Lang RA, Jamrich M. Eye formation in the absence of retina. Dev Biol. 2008;322:56–64.

30 Eye 30. Cvekl A, McGreal R, Liu W. Lens development and crystallin gene expression. In: Hejtmancik JF, Nickerson JM, editors. Progress in molecular biology and translational science. London: Elsevier/ Academic Press; 2015. p. 129–67. 31. Andley UP.  Crystallins in the eye: function and pathology. Prog Retin Eye Res. 2007;26:78–98. 32. Shastry BS.  Persistent hyperplastic primary vitreous: congenital malformation of the eye. Clin Exp Ophthalmol. 2009;37:884–90.

323 33. Sellheyer K, Spitznas M.  Ultrastructure of the human posterior tunica vasculosa lentis during early gestation. Graefes Arch Clin Exp Ophthalmol. 1987;225:377–83. 34. Mitchell CA, Risau W, Drexler HC.  Regression of vessels in the tunica vasculosa lentis is initiated by coordinated endothelial apoptosis: a role for vascular endothelial growth factor as a survival factor for endothelium. Dev Dyn. 1998;213:322–33.

31

Ear Dale S. Huff

Introduction The ear is often involved in intrauterine and neonatal diseases such as congenital anomalies, chromosomal and gene defects, amniotic fluid infection sequence, transplacental infections, maternal-placental and fetal-placental perfusion defects, other causes of fetal hypoxia, congenital tumors, and other diseases. Therefore, histologic examination of the ear should be an important part of the perinatal autopsy. However, because the anatomy of the ear is complex, because the removal and plane of sectioning of the petrous bone are often haphazard, and because the specimen is often fragmented or incomplete, interpretation of routine microscopic sections of the ear is often exasperating and unhelpful. As a consequence, histologic examination of the ear is often neglected in perinatal autopsies. Understanding the anatomy of the ear, standardizing the method of removal and sectioning of the petrous bone, and making examination of the ear a routine part of the perinatal autopsy will make histologic examination of the ear more satisfying and fruitful. The purpose of this chapter is not only to demonstrate the histology of the ear but also persuade the reader that histologic examination of the ear is a worthwhile part of perinatal autopsy.

Anatomy I n Situ Surface Anatomy of the Petrous Bone in the Neonate The inner and middle ear and part of the external ear lie within the petrous bone. The surface anatomy of the neonatal petrous bone serves two functions for the perinatal patholoD. S. Huff (*) Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]

gist. First, understanding the surface anatomy allows the pathologist to develop a mental picture of the position and relationships of the components of the inner, middle, and external ears within the petrous bone. Second, the surface landmarks provide guideposts for the placement of resection lines for the removal of the petrous bone during the perinatal autopsy. This section describes those surface landmarks. The petrous bone is an oblong wedge obliquely oriented in the floor of the cranium. When viewed in situ from above in the opened cranium, it presents two surfaces. The superior horizontal surface is the roof of the petrous bone and forms much of the floor of the middle cranial fossa. The posterior vertical wall attaches to the posterior edge of the superior wall and drops vertically downward, forming a right angle with the superior wall. The posterior wall of the petrous bone forms the anterior wall of the posterior cranial fossa. The angle formed by the junction of the superior horizontal and posterior vertical walls is the ridge of the petrous bone. The petrous bone is oriented obliquely in an anteromedial to posterolateral direction. The anteromedial end, which is narrower, abuts against the lateral wall of the sella turcica. The posterolateral end, which is broader, abuts against the inside of the lateral wall of the cranium near the posterior end of the squamous portion of the temporal bone. The anteromedial end of the petrous ridge is sharp; the posterolateral end is more rounded. In the neonate, the boundaries of the cranial bones are sharply outlined by the wide-open sutures. The superior horizontal surface, the posterior vertical surface, the petrous ridge, and the open sutures are surface landmarks that aid in identification of the petrous bone at autopsy (Fig. 31.1) [1, 2]. The arcuate eminence and the subarcuate fossa are additional landmarks on the superior surface. The arcuate eminence is a curved ridge on the superior surface. It contains the superior semicircular canal and duct. The subarcuate fossa is a blind-ending depression that extends under the arcuate eminence. It is lined with dura, which transmits the subarcuate artery to the inner ear. The tegmen tympani,

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_31

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D. S. Huff Anterior clinoid process Pituitary fossa Posterior clinoid process

Cut edge of parietal bone Temporalis muscle Subcutaneous fat Pinna

Clivus Petrous ridge Internal auditory meatus Jugular fossa Ostium of cochlear aqueduct Foramen magnum

Tegmen tympani Arcuate eminence Subarcuate fossa Superior horizontal wall of petrous bone Squamous part of temporal bone Parietal bone Petrous ridge Posterior vertical wall Ostium of vestibular aqueduct

Fig. 31.1  In situ surface anatomy of the petrous bone of the neonate. This illustration views the right side of the cranium from above in the opened cranium. It focuses on the floor of the right middle fossa and the right petrous bone. The left edge is the midline. Dark green: petrous bone. Light green: squamous part of temporal bone. Orange: sphenoid

bone. Blue: occipital bone. Violet: parietal bone. Tan areas and lines between bones: open sutures and minor fontanelles. The ostia of the cochlear and vestibular aqueducts are tucked under the edge of the petrous bone and are not visible on the surface. The indicated positions are approximate

though not delineated by surface landmarks, is an important area to visualize. It is a very thin area of the superior horizontal wall that forms the roof of the middle ear. Because it is so thin, it can easily be removed to unroof the middle ear and expose the epitympanic recess and the other structures within those cavities (Fig. 31.1). The internal acoustic meatus, jugular fossa, ostium of the cochlear aqueduct, and the orifice of the vestibular aqueduct are additional landmarks located from anteromedial to posterolateral along the posterior wall. The internal acoustic meatus is the most anterior landmark. It is the orifice of the internal acoustic canal through which the eighth cranial nerve passes from the posterior fossa to the inner ear. The base of the cochlea lies adjacent to the internal end of the canal. The jugular fossa is important because it marks the site of the ostium of the cochlear aqueduct, which is tucked under the inferior edge of the petrous bone at the apex of the jugular fossa and is therefore not visible on the surface of the petrous bone. It contains the periotic duct. The orifice of the vestibular aqueduct is hidden in a depression in the posterior wall posterior to the jugular fossa. It contains the endolymphatic sac. The position of the cochlea can be estimated from the location of the internal acoustic meatus. The position of the saccular dilatation of the vestibule can be estimated from the

location of the ostium of the cochlear aqueduct. The position of the utricular dilatation of the vestibule can be estimated from the location of the ostium of the vestibular aqueduct. The arcuate eminence marks the exact position of the superior semicircular canal and duct (Figs. 31.1 and 31.2).

 rrangement of the Inner, Middle, and External A Ears Within the Petrous Bone The spatial relationships between the petrous bone, inner ear, and middle ear are intricate (Figs. 31.2 and 31.3). They are parallel to each other, with the middle ear lateral to the inner ear. They are obliquely oriented from anteromedial to posterolateral. They extend from the lateral wall of the sella turcica anteromedially to the posterior end of the squamous portion of the temporal bone posterolaterally. The inner ear consists of an outer osseous labyrinth, which is also called the otic capsule, and the membranous labyrinth, which lies within the otic capsule (Figs. 31.3, 31.4, 31.5, 31.6, 31.7, and 31.8). The auditory and vestibular sensory end organs lie within the membranous labyrinth (Figs.  31.8, 31.9, 31.10, 31.11, 31.12, and 31.13). In addition, the otic capsule, membranous labyrinth, and middle ears are divided into three seg-

31 Ear

327

Eustachian tube

Anterior part Tympanic membrane forming the lateral wall of the isthmus

External auditory canal

Isthmus or middle part Epitympanic recess Mastoid additus Mastoid antrum

Fig. 31.2  Arrangement of inner, middle, and external ears. This is the same as Fig. 31.1 but with the inner ear, middle ear, and external auditory canal projected into the petrous bone. The osseous labyrinth is depicted in blue, the middle ear in red, and the external auditory canal in green. See Fig. 31.1 for the color code and labels of the bones and surface landmarks, and see the following figures (especially Figs. 31.4 and 31.6) for identification of the various components of the osseous labyrinth of the inner ear. From anteromedial to posterolateral, these components are the cochlea, the vestibule (with saccular and utricular dilatations), and the semicircular canals. The various components of the middle ear from anterolateral to posterior in the figure are as fol-

lows: Eustachian tube, anterior part, isthmus or middle part, tympanic membrane, forming the lateral wall of the isthmus. Note that the tympanic membrane is composed of an inner red layer (mucosa of middle ear) and an outer green layer (mucosa of the external auditory canal). Note the aditus, the small window at the waist between the epitympanic recess and the mastoid antrum. Note the unlabeled cochlear aqueduct and vestibular aqueduct. Finally, note the alignment of the anterior part of the middle ear with the cochlea, the isthmus with the vestibule, and the epitympanic recess and mastoid antrum with the semicircular canals

ments from anterior to posterior. The three segments of the otic capsule (osseous labyrinth) from anterior to posterior are the cochlea, vestibule, and semicircular canals. The vestibule is further divided from anterior to posterior into the saccular dilatation and the utricular dilatation (see Figs. 31.4 and 31.6). The analogous segments of the membranous labyrinth from anterior to posterior are the cochlear duct within the cochlea, the saccule within the saccular dilatation of the vestibule, the utricle within the utricular dilatation of the vestibule, and the semicircular ducts within the semicircular canals (see Figs. 31.5 and 31.7). The analogous segments of the middle ear are the anterior part, which is anterior to the tympanic membrane, lateral to the cochlea, and gives rise to the Eustachian tube from its anterior end; the middle part or isthmus, which is lateral to the vestibule and whose entire lateral wall is the tympanic membrane; and the posterior part, which is posterior and superior to the tympanic membrane, lies lateral to the semicircular canals, and is further

subdivided from anterior to posterior into the epitympanic recess, the mastoid antrum, and the mastoid air cells. The inner and middle ears are so intimately pressed together that the basal spiral of the cochlear duct bulges into the tympanic isthmus anteriorly, forming the promontory on its medial wall. The isthmus of the tympanic cavity and the vestibule share a common boney wall. The lateral semicircular canal bulges into the epitympanic recess posteriorly (see Figs. 31.2 and 31.3). The surface anatomy of the petrous bone of the neonate is different from that of the adult. As the bones mature, the surface landmarks change. For example, the sutures close, making the borders of the petrous bone less clear. The petrous ridge becomes sharper throughout its entire length. The arcuate eminence and subarcuate fossa are smoothed over and become barely visible. The orifice of the vestibular aqueduct becomes hidden in a linear crevice that is higher on the posterior wall in the adult than in the neonate. The changing

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D. S. Huff

Epitympanic recess

1 Lateral semicircular duct and canal

Temporalis muscle Superior semicircular duct and canal

Tegmen tympani

Arcuate eminence Aditus to mastoid antrum

Internal auditory meatus and canal

External auditory canal

Utricle Saccule

Tympanic membrane

Helicotrema Vestibule

Round window Cochlear duct

Cecum of cochlear duct

1

Scala vestibuli

Nasopharynx

Scala tympani Eustacian tube

Fig. 31.3  Frontal section through the external, middle, and inner ears viewed from anterior to posterior, looking into the epitympanic recess and the aditus to the mastoid antrum. The tympanic membrane is tilted so that its inferior border is attached to the tympanic ring more medially than its superior border. As a result, the inferior wall of the external auditory canal is longer than its superior wall. Nearly the entire lateral wall of the tympanic cavity is formed by the tympanic membrane. The roof of the epitympanic recess is formed by the thin plate of bone called the tegmen tympani, which is an important surface landmark useful in the correct positioning of the incision lines for the removal of the petrous bone at autopsy. The aditus is a window in the posterior wall of the epitympanic recess, which opens into the mastoid antrum. The common boney wall between the tympanic cavity and the vestibule is perforated by two windows. The more superior is the oval window, which is closed by the footplate of the stapes and the radial ligament. The more inferior is the round window, which is closed by the secondary tympanic membrane. The scala vestibuli begins at the oval window, and the scala tympani ends blindly at the round window. The boney promontory, which is formed by the wall of the basal turn of the cochlea, lies between the oval and round windows. The opening of the Eustachian tube in the tympanic Fig. 31.4  Right osseous labyrinth (otic capsule) viewed from behind and medial, similar to the orientation in Fig. 31.2. The saccular and utricular dilatations of the vestibule contain the membranous saccule and utricle, respectively. The dilated ends of the three semicircular canals where they join the vestibules are the ampules of the semicircular canals. The sites of the ostia of the cochlear and vestibular aqueducts are approximate

cavity is inferior and anterior to the round window. The Eustachian tube passes beneath the cochlea, and its course to the nasopharynx is nearly horizontal. The superior semicircular canal protrudes upward to form a raised linear prominence, the arcuate eminence, on the floor of the middle fossa. The lateral semicircular canal protrudes laterally to form a boney prominence on the medial wall of the epitympanic recess. The blind end, or cecum, of the cochlear duct abuts against the vestibular side of the promontory so that the scala tympani ends blindly at the round window. The duct connecting the cochlear duct to the saccule is the ductus reuniens. The arrows indicate that the pressure waves created by the stapes in the perilymphatic fluid travel through the scala vestibuli toward the cupola (apex) of the cochlea, through the helicotrema, into the scala tympani, back through the cochlea, and to the secondary tympanic membrane in the round window. Cranial nerves VII and VIII pass through the internal auditory canal and meatus in the posterior vertical wall of the petrous bone below the petrous ridge. Line “1” (the curved vertical line through the temporalis muscle and external auditory canal) is the frontal view of the first incision in removal of the petrous bone (see Fig. 31.64). Color code of the segments of the ear: green, external auditory canal; red, middle ear; blue, membranous labyrinth of the inner ear [4] Vestibule

Cupola of cochlea

Saccular dilatation

Vestibular dilatation

Superior semicircular canal

Ostium of vestibular aqueduct

Lateral semicircular canal

Ostium of cochlear aqueduct

Common osseous limb

Round window

Posterior semicircular canal

31 Ear Fig. 31.5  Right membranous labyrinth viewed from behind and medial, similar to the orientation in Figs. 31.2 and 31.4. The membranous labyrinth fits within the osseous labyrinth. The vestibular and facial nerves have been omitted

329 Utricle Cochlear duct

Superior semicircular duct Common membranous limb

Cranial nerve VIII Saccule

Lateral semicircular duct

Utriculosaccular duct

Posterior semicircular duct

Endolymphatic duct and sac

Fig. 31.6  Right otic capsule viewed from in front and laterally. The bony wall of the vestibule in this view is also the medial wall of the tympanic cavity, containing the oval and round windows separated by the promontory

Ampula of superior semicircular canal

Superior semicircular canal

Utricular dilatation

Common osseous limb Posterior semicircular canal

Saccular dilatation

Lateral semicircular canal Oval window

Vestibule

Cupola of cochlea

Promontory Round window

Fig. 31.7  Right membranous labyrinth from in front and laterally, similar to the orientation of the osseous labyrinth seen in Fig. 31.6. Note the anterior recess of the utricle overlying the saccule

Superior semicircular duct Endolymphatic sac Endolymphatic duct Common membranous limb Lateral semicircular duct Posterior semicircular duct

Utricle Anterior recess of utricle Saccule

Blind end of cochlear duct Cecum of cochlear duct Ductus reuniens Cochlear duct

anatomy of the petrous bone from the embryo to the adult has been described previously [2]. Compare the petrous bone of the neonate in Fig. 31.1 with that of the adult in any atlas of adult anatomy [3]. Knowledge of the changing anatomy of the fetal and neonatal petrous bone will aid the perinatal pathologist in its identification and removal at autopsy.

therefore composed of bone (see Fig. 31.3) [3, 4]. The bony tympanic ring surrounds the inner end of the bony canal. The tympanic membrane is attached to a groove in the tympanic ring.

External Auditory Canal

The tympanic ring and the attachment of the tympanic membrane to it are tilted so that the posterior and inferior attachments are much deeper than the anterior and superior attachments. The tilt is so marked that the tympanic membrane is nearly parallel to the posterior-inferior wall of the

The wall of the external two thirds of the external auditory canal is composed of soft tissue and cartilage. The wall of the inner one third passes through the temporal bone and is

Tympanic Membrane

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Boney cochlea

Scala vestibuli

Modiolus

Reisener’s membrane

Stria vascularis

Spiral limbus Boney spiral lamina Spiral ganglion

A

C

Tectorial membrane Crest of spiral ligament Outer hair cells, three rows

Cochlear nerve fibers Pillar cells Scala tympani To cochlear nerve

B

Scala media (cochlear duct)

Inner hair cells, one row

A. Inner spiral sulcus B. Tunnel of corti C. Outer spiral sulcus

Fig. 31.8  Cross section of the bony cochlea, the membranous cochlear duct, and the organ of Corti. The bony cochlea encloses the membranous triangular cochlear duct (also called scala media) in the middle, the scala vestibuli above, and the scala tympani below. The bony central core of the cochlea, the modiolus, is at the left. The bilaminar bony spiral lamina extends from the area of the modiolus toward the center of the bony cochlear canal. The lateral wall of the cochlear duct is formed by a cushion of connective tissue, the spiral ligament, which lines the inner lateral surface of the bony cochlea. A sharp crest originates from the lower part of the spiral ligament and extends toward the center of the bony canal opposite to the bony spiral lamina. The basal membrane of the cochlear duct is attached to the bony spiral lamina medially and to the crest of the spiral ligament laterally, to form the floor or base of

Common membranous limb

Lateral semicircular duct

the cochlear duct. The roof of the cochlear duct is Reissner’s membrane, which attaches to the spiral limbus medially and to the upper end of the spiral ligament laterally. Therefore, the lateral wall of the cochlear duct lies between the attachment of Reissner’s membrane to the spiral ligament superiorly and the attachment of the basal membrane to the crest of the spiral ligament inferiorly. The organ of Corti lies on the basal membrane. The stria vascularis lies on the surface of the spiral ligament between Reissner’s membrane and the basal membrane. The fine peripheral fibers of the cochlear nerves, which innervate the organ of Corti, pass through the middle of the bony spiral lamina and join with the spiral cochlear ganglion located in the area of the modiolus medially. Fibers from the spiral cochlear ganglion pass into the hollow center of the modiolus to form the cochlear nerve

Superior semicircular duct Ampulla (Window cut in free wall) Cut edge of wall of ampulla Lumen of ampulla

Posterior semicircular duct

Utricle (window cut in anterior recess) Lumen of utricle

Cut edge of crista

Utricular otolithic macule(horizontal)

Crista Horizontal floor of anterior recess of utricle

Fig. 31.9  The vestibular portion of the membranous labyrinth looking from anterior to posterior. Windows have been cut in the saccule, utricle, and each of the ampules of the three semicircular ducts to show the sensory organs of the vestibular system. The saccular otolithic macula is oriented vertically on a vertical wall of the saccule. This is an en face view of the saccular macula. The utricular otolithic macula is oriented

Lumen of saccule Sacullar otolithic macule(vertical) Saccule (window cut in wall)

horizontally on the horizontal floor of the anterior recess of the utricle, above the saccular otolithic macula. The two macules are therefore perpendicular to each other. This view of the utricular macula is slightly on edge. The three ampullary cristae are partially circumferential ridges on the inner surfaces of the ampullae, which partially occlude the ampullary lumens

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external auditory canal. The tympanic membrane is the medial wall of the external auditory canal and the lateral wall of the tympanic cavity. It therefore is a common wall between the external auditory canal and the tympanic cavity. Because it is attached to the inner end of the bony external auditory canal, attempts to remove the middle and inner ears by an incision through the petrous portion of the temporal bone flush with the inner surface of the temporal bone often result in destruction of the tympanic ring, tympanic membrane, and the middle ear.

Middle Ear The tympanic cavity is a long, narrow, rectangular cavity. It lies parallel to the full length of the lateral surface of the inner ear (see Figs. 31.2 and 31.3) [5]. The anterior end is anterior to the tympanic membrane. Part extends below the lower border of the tympanic membrane, especially anteriorly, and is called the hypotympanum. The anterior end is adjacent to the cochlea of the inner ear. The opening of the Eustachian tube is in the anterior-inferior aspect of the medial wall adjacent to the cochlea. The middle part is adjacent to the vestibule of the inner ear. It is the main chamber

Otolith

of the middle ear and is sometimes referred to as the isthmus because it is narrower than the anterior and posterior ends. Its lateral wall is the tympanic membrane. The tympanic membrane is therefore a common wall between the external auditory canal and the tympanic cavity. The isthmus contains the handle of the malleus attached to the tympanic membrane, the long process of the incus, and the stapes. The medial bony wall of the isthmus is the lateral wall of the vestibule. The lateral wall of the vestibule is therefore a common wall between the tympanic cavity and the vestibule. There are two windows in the medial wall: the round window, located anterior-inferiorly, and the oval window, located posterior-superiorly. The round window lies immediately behind the orifice of the Eustachian tube and is closed by a delicate membrane called the secondary tympanic membrane, which separates the tympanic cavity from the scala

Semicircular duct cut end

Crista Cut edge of Crista

Cut edge of cupola

Cupola

Cut edge of ampulla Edge utricular otolithic macula Gelatinous membrane

Fig. 31.10  En face view of the utricular macula. The red otoliths are embedded in the free surface of the gelatinous membrane that covers the top of the macula. The cilia of the hair cells are embedded in the deep surface of the membrane

Otolith membrane

Tall epethelial cells Connective tissue with nerve fibers

Fig. 31.11  Cross section of an otolithic macula. Cross sections of the saccular and utricular otolithic maculae are identical. They cannot be differentiated microscopically. The red cells are neurosensory hair cells. The cilia are embedded in the base of the gelatinous membrane that covers the top of the macula. The green cells are supportive cells.

Junction of ampulla with utricle

Fig. 31.12  A close-up view of the ampulla of a semicircular duct. A window has been cut in the ampulla to show the details of the crista inside. The cupola is a tongue-shaped, gelatinous membrane attached to the free edge of the crista. It extends partway across the lumen of the ampulla and does not contain otoliths

Otoliths Hairs of hair cells Hair cells Support cells

The blue cells are tall epithelial cells adjacent to the macula. They secrete the gelatinous membrane. The otoliths are embedded in the free surface of the gelatinous membrane. A thick pad of connective tissue containing peripheral fibers of the vestibular nerve lies beneath the epithelial macula

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Gelatinous cupola

Hairs and cilia of hair cells Hairs cells (red) Support cells (green) Tall epithelial cells Connective tissue with nerve fibers

the middle fossa called the tegmen tympani (see Figs. 31.1, 31.2, and 31.3). The tympanic cavity can easily be unroofed by removing the thin tegmen tympani. The wall of the lateral semicircular canal bulges into the epitympanic recess. An aperture in the posterior wall of the epitympanic recess, the mastoid aditus, leads posteriorly into the mastoid antrum and from there to the mastoid air cells, which are part of the middle ear. In most fetuses and neonates, the mastoid antrum is the most posterior part of the middle ear because the mastoid air cells, which are the most posterior part in adults, usually do not develop until late fetal or postnatal life (see Fig. 31.2). The Eustachian tube exits from the anterior-inferior aspect of the medial wall of the tympanic cavity anterior to and below the round window, passes under the cochlea, and opens into the upper nasopharynx. In the fetus and infant, the tube is oriented horizontally (see Fig.  31.3). The tympanic cavity and mastoid air cells are in direct continuity with the outside environment through the Eustachian tube. Like the lung, before birth the middle ear contains amniotic fluid; after birth it contains air. The horizontal orientation of the Eustachian tube in the fetus and infant makes entry of amniotic fluid or nasopharyngeal fluid into the middle ear very easy. If those fluids are infected, intrauterine or neonatal otitis media and sepsis will likely follow. The spectrum of aspiration of amniotic fluid and the amniotic fluid infection sequence affects not only the lung but also the middle ear.

Inner Ear Fig. 31.13  Cross section of a crista. The red cells at the tip of the crista are neurosensory hair cells. The hairs and cilia of the hair cells are embedded in the base of the gelatinous cupola. Notice the absence of otoliths. The green cells at the tip are supportive epithelial cells. The blue cells along the edge of the stalk of the crista secrete the gelatinous cupola. The connective tissue core contains tiny peripheral branches of the vestibular nerve

tympani on the vestibular side of the window. The oval window is located posterior and superior to the round window. The foot plate of the stapes fills the oval window. The radial ligament attaches the edges of the foot plate to the edges of the oval window. The oval window separates the tympanic cavity from the scala vestibuli on the vestibular side. The base of the cochlea pushes the bony wall toward the tympanic cavity, producing a bulge in the wall between the round and oval windows, which is called the promontory. The posterior part is a recess in the posterior roof of the tympanic cavity, which forms an attic-like space superior and posterior to the upper rim of the tympanic membrane (see Figs. 31.2 and 31.3). This space is the epitympanic recess. It lies adjacent to the semicircular canals and contains the head of the malleus and the body and short process of the incus. The roof of the epitympanic recess is a thin region of the bony floor of

Osseous Labyrinth The inner ear is an irregular, oblong cavity in the petrous portion of the temporal bone. It is surrounded by an osseous capsule called the osseous labyrinth, bony labyrinth, or otic capsule (see Figs. 31.4 and 31.6) [6]. These three terms are used interchangeably. The osseous labyrinth has three interconnected parts (from anteromedial to posterolateral): the cochlea, which is a snail-shaped spiral tube joined to the anterior aspect of the vestibule; the vestibule, which is the central chamber; and the three semicircular canals, which are joined to the posterior aspect of the vestibule. The common cavity of the osseous labyrinth is the perilymphatic space, which is filled with fluid, the perilymph. The base of the snail-shaped cochlea faces posteriorly and slightly medially and lies at the inner opening of the internal auditory meatus. The apex or cupola of the “snail” points anteriorly and slightly laterally. The cochlea has a bony central core, the modiolus, which has been likened to a screw with the head of the screw at the base of the cochlea and its tip at the apex. The spiral lamina of the cochlea is analogous to the thread of the screw, and it projects like a shelf into the lumen of the cochlear canal along its equator. Its tip reaches the center of the canal. The spiral lamina forms part of the floor or base of

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the triangular cochlear duct. The vestibule presents two dilatations: the saccular dilatation anteriorly at the junction of the vestibule with the cochlea and the utricular dilatation posteriorly at the junction of the vestibule with the semicircular canals (see Figs. 31.4 and 31.6). The anatomy of the lateral wall of the vestibule, round window, promontory, and oval window is described above in the description of the middle ear. Three openings in the posterior wall of the vestibule lead to canals through the posterior wall of the petrous bone. The three canals are the internal auditory canal (also called the internal acoustic canal), the cochlear aqueduct (also called the cochlear canaliculus), and the vestibular aqueduct (also called the vestibular canaliculus) (see Figs. 31.1, 31.2, and 31.4). The most anterior is the internal auditory canal. Its opening in the osseous labyrinth is at the base of the spiral cochlea. Cranial nerves VII (facial) and VIII (vestibuloacoustic) pass through the canal and exit through its foramen, the internal auditory meatus (also called the internal acoustic meatus), in the posterior wall of the petrous bone. The opening of the cochlear aqueduct in the osseous labyrinth is in the wall of the scala tympani at the junction of the cochlea with the vestibule, and it exits the posterior inferior wall of the petrous bone into the subarachnoid space at the apex of the jugular foramen, creating a direct communication between the perilymphatic and subarachnoid spaces. It contains the periotic duct and a fluid that may be perilymph or subarachnoid fluid. It has no connection with the membranous labyrinth [7]. The most posterior is the vestibular aqueduct. Its opening in the osseous labyrinth is in the utricular dilatation of the vestibule, near its junction with the semicircular canals. It exits through a slit-­ shaped foramen in the posterior wall of the petrous portion of the temporal bone posterolateral to the jugular foramen (see Figs. 31.1 and 31.2). It contains the endolymphatic duct, which is a tube of membranous labyrinth that arises from the utriculosaccular duct and ends blindly in the dilated endolymphatic sac in the epidural space (see Figs.  31.1, 31.2, 31.5, and 31.7). It contains endolymph. The posterior segment of the osseous labyrinth is composed of the three semicircular canals: anterior (also called superior), posterior, and lateral. Both ends of each canal join the utricle. The three anterior ends are dilated; they are called the ampullae at their sites of union with the posterior-superior aspect of the vestibule. The posterior limbs of the anterior and posterior canals join to form the common crus. Therefore, there are only two openings into the posterior aspect of the utricle for the posterior limbs: one for the common crus and one for the lateral canal (see Figs. 31.4 and 31.6).

Membranous Labyrinth The membranous labyrinth is a closed system of epithelial ducts and sacs derived from the ectodermal otic vesicle. It lies within the cavity of the osseous labyrinth (see Figs.  31.3,

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31.5, and 31.7). It is composed of three major parts (from anteromedial to posterolateral): the cochlear duct within the bony cochlea; the saccule and utricle, which are two sacs within the saccular and utricular dilatations of the vestibule; and the three semicircular ducts within the bony semicircular canals. The cochlear duct begins as a blind end in the vestibule adjacent to the promontory and below the saccule. It follows the osseous cochlear spiral to its apex, the cupola, where it ends blindly. The cochlear duct therefore has two blind ends. One abuts the bony promontory in the vestibule below the saccule. The other lies at the apex of the spiral cochlea (see Figs. 31.3, 31.5, 31.7, and 31.8). The saccule is located anteriorly within the saccular dilatation of the vestibule, above the blind end of the cochlear duct and below and in front of the utricle. It is oval in shape and smaller than the utricle. The utricle is located posteriorly in the utricular dilatation of the vestibule, above and behind the saccule. A blindended recess of the utricle, the anterior recess, arises near the anterior opening of the lateral semicircular duct. This recess extends horizontally over the roof of the saccule (see Fig.  31.7) [8]. The three semicircular ducts empty into the posterior-superior aspect of the utricle. One end of each forms a dilated ampule at its site of insertion into the utricle. The three components of the membranous labyrinth are interconnected by small ducts. The ductus reuniens connects the cochlear duct to the saccule (see Figs.  31.3 and 31.7). The orifice of the ductus reuniens in the cochlear duct is some distance from its blind end. The segment of the cochlear duct between its blind end and the orifice of the ductus reuniens is called the cecum of the cochlea (see Figs. 31.3 and 31.7). The utriculosaccular duct connects the saccule with the utricle. The endolymphatic duct arises from the utriculosaccular duct (see Figs. 31.5 and 31.7). The posterior segment of the membranous labyrinth is composed of the three semicircular ducts inside the semicircular canals. Each has an ampulla, and each necessarily follows the course of its corresponding canal. Unlike the canals that join with the vestibule, the semicircular ducts connect to the utricle. The various parts of the entire membranous labyrinth are interconnected, from the cochlear duct anteromedially to the semicircular canals posterolaterally (see Figs. 31.3, 31.5, and 31.7).

Perilymphatic Space The perilymphatic space completely surrounds the membranous labyrinth and separates the osseous labyrinth from the membranous labyrinth. Like the osseous labyrinth, membranous labyrinth, and middle ear, it can be divided into three parts, from anteromedial to posterolateral. The anterior part, in the cochlea, is composed of two parts: the scala vestibuli and the scala tympani. The membranous cochlear duct divides the spiral bony cochlear tube into three tubes: the triangular cochlear duct (scala media) in the middle; the scala vestibuli, above the cochlear duct adjacent to Reissner’s

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membrane; and the scala tympani, below the cochlear duct adjacent to the basal membrane (see Figs.  31.3 and 31.8). The lumens of the scala vestibuli and scala tympani become continuous with each other at the apex of the spiral, through an aperture called the helicotrema. The scala tympani ends blindly at the round window. The scala vestibuli joins the perilymphatic space of the vestibule at the oval window. The perilymphatic space of the vestibule is the cisterna vestibuli. It surrounds the saccule and utricle and is continuous with those of the semicircular canals (see Fig. 31.3).

Periotic Duct The periotic duct is a direct communication between the scala tympani part of the perilymphatic space and the subarachnoid space of the posterior fossa. The duct lies within the cochlear aqueduct, a bony canaliculus in the otic capsule and posterior wall of the petrous bone (see Figs. 31.1, 31.2, and 31.4). (The otic capsule is described above and in the Embryology and Histology sections below).  uditory and Vestibular Sensory Organs A Two systems of sensory end organs are present in the inner ear: the cochlear system, which detects sound, and the vestibular system, which detects motion and gravity. The cochlear auditory sensory end organ is the organ of Corti, which is a strip of neurosensory epithelium located in the basal membrane (the floor) of the entire length of the spiral cochlear duct (see Fig. 31.8). The vestibular system is composed of two groups of neurosensory epithelial end organs. The first group is composed of two otolithic maculae, the saccular macula and the utricular macula, which detect linear motion. The two otolithic maculae are an interrelated pair. The saccular otolithic macula is a thickened ellipsoid plaque of neurosensory epithelium in the epithelial lining of the vertical wall of the saccule,

D. S. Huff

which lies below the anterior recess of the utricle. Its flat surface is oriented vertically, and its long axis is oriented anteroposteriorly (Fig. 31.9) [5, 9]. It detects vertical linear motion and gravity. The utricular otolithic macula is a thickened ellipsoid plaque of neurosensory epithelium in the epithelium of the horizontal floor of the anterior recess of the utricle. Its flat surface lies horizontally, and its long axis is oriented anteroposteriorly (Figs. 31.9, 31.10, and 31.11). It lies immediately above and perpendicular to the vertically oriented saccular macula [5, 10]. It detects horizontal linear motion. The second group is the three ampullary cristae or ridges of sensory epithelium, one in each of the three ampullae of the semicircular ducts. The three ampullary cristae, an interrelated trio, are linear ridges of thickened neurosensory epithelium oriented transversely to the long axis of the ducts. They extend only partially around the inner circumference of the ampullae (see Figs. 31.9, 31.12, and 31.13). Each is oriented in a plane different from the others. The cristae detect angular motion.

Embryology Both the epithelial and connective tissues of the ear are derived from two closely related embryologic primordia, both of which arise within the ectodermal ring [11, 12]. The inner ear is derived from the otic placode, which is an ectodermal thickening within the rostral segment of the ectodermal ring. The external and middle ears are derived from the first and second pharyngeal complexes, which arise from the pharyngeal segment of the ectodermal ring (Table 31.1). Embryonic development of the ear begins early in the sixth postmenstrual week with the appearance first of the otic placode (the future membranous labyrinth of the inner ear), followed by the first and second pharyngeal arches (the future auricle), the first pharyngeal cleft (the future external auditory

Table 31.1  The embryological origins of the external, middle, and inner ears First pharyngeal arch  Anterior part of auricle (from auricular hillocks 1, 2, and 3)  The head of malleus, body and short process of incus (from posterior end of Meckel’s cartilage and mesoderm of first pharyngeal arch) Second pharyngeal arch  Posterior part of auricle (from auricular hillocks 4, 5, and 6)  Handle of malleus, long process of incus, head and limbs of stapes (from Reichert’s cartilage and mesoderm of second pharyngeal arch). Note: the foot plate of the stapes is from the otic capsule First pharyngeal cleft  External auditory canal  External epidermal surface of tympanic membrane First pharyngeal pouch  Eustachian tube  Tympanic cavity  Internal endodermal surface of tympanic membrane Otic placode, pit, and vesicle  Membranous labyrinth: cochlear duct, ductus reuniens, saccule, utriculosaccular duct, endolymphatic duct and sac, utricle, and semicircular ducts Periotic mesoderm  Osseous labyrinth (otic capsule): cochlea, vestibule, and semicircular canals  Foot plate of stapes

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canal), and the first pharyngeal pouch, also referred to as the tubotympanic recess (the future tympanic cavity and Eustachian tube). All appear early in the sixth week. During the rest of the sixth week, the first pharyngeal cleft and first pharyngeal pouch grow toward each other, but their distal ends remain separated by a layer of mesenchyme from the arches. The distal ends of the cleft and pouch and the intervening mesenchyme are the first pharyngeal membrane, which will become the future tympanic membrane. The primordia of the ossicles condense within the mesenchyme of the first pharyngeal membrane and the mesenchyme adjacent to the first pouch: first the malleus, in the mesenchyme of the first pharyngeal arch, located in first pharyngeal membrane; followed by the incus, in the mesenchyme of the first arch, adjacent to the first pharyngeal pouch; then the stapes, in the mesenchyme of the second arch, adjacent to the first pouch. Meanwhile, the otic placode develops into the otic pit. A layer of mesenchyme condenses around the epithelial pit. The mouth of the pit becomes narrow, and the pit becomes the otic vesicle. The epithelium of the otic pit becomes the membranous labyrinth of the inner ear. The condensed mesenchyme surrounding the epithelial pit becomes the osseous labyrinth of the inner ear. These primordia of the auricle, external auditory canal, tympanic membrane, ossicles, tympanic cavity, Eustachian tube, and membranous and osseous labyrinths are established by the end of the sixth week. From the beginning of the seventh week to the end of the tenth week (the end of the embryonic period), the final form of all of the organs develops from these primordia. The following events are occurring simultaneously during the last 4 weeks of the embryonic period. The auricle is completely formed by the middle of the tenth week. The first pharyngeal cleft becomes the external auditory canal. In the tenth week, the lining epithelium of the canal proliferates and forms a plug, the meatal plug, which completely occludes the lumen of the distal third of the canal [5, 13]. The tympanic end of the tubotympanic recess enlarges to become the tympanic cavity, and its pharyngeal end becomes the Eustachian tube. The head of the malleus forms in the mesenchyme of the future tympanic membrane in association with the posterior end of the cartilage of the first arch, Meckel’s cartilage. The body and short process of the incus form adjacent to the wall of the tympanic cavity, also in association with the cartilage of the first arch, Meckel’s cartilage. The head of the malleus and the body and short process of the incus come to lie in the epitympanic recess. The handle of the malleus arises from the mesenchyme of the second arch in association with posterior end of the cartilage of the second arch, Reichert’s cartilage, and remains attached to the tympanic membrane. The long process of the incus and the head and limbs of the stapes also form in association with the cartilage of the second arch, Reichert’s cartilage, and come to lie within the main part of the tympanic cavity, the isthmus. The footplate of the stapes arises from the otic capsule and fills the oval window. The parts of the ossicles derived from the first arch are located in the epitympanic recess, whereas the parts derived from the second arch

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are located in the isthmus of the tympanic cavity. By the end of the tenth week, the mesenchymal ossicles have nearly attained their adult morphology and have begun to convert to cartilage. The diarthrodial synovial incudomalleolar and incudostapedial joints are formed, and the stapediovestibular joint is formed by the radial ligament. The enlarging tympanic cavity envelops the ossicles and the mesenchyme that surrounds them, so that they lie within the tympanic cavity, and the tympanic cavity is filled with the primitive mesenchyme. The development of the inner ear is even more complex [11, 12]. The otic vesicle detaches from the surface ectoderm and comes to lie in its final position, medial to the developing tympanic cavity. The epithelium of the otic vesicle is thick, multilayered, and composed of small, oval, basophilic, undifferentiated cells with the appearance of pluripotent placodal cells. This histology remains unchanged until the epithelium begins to differentiate during the tenth week. During the seventh week, the round otic vesicle elongates in the ventral and dorsal directions. The ventral elongation will become the cochlear duct; the dorsal elongation, the endolymphatic duct. During the eighth week, the semicircular ducts appear as three hollow plates at the base of the endolymphatic duct. The epithelium in the center of the plates fuses, leaving the patent semicircular ducts at the periphery of the plates. By the end of the eighth week, the fused epithelium in the center of the plates disappears, and one end of each duct dilates at its point of insertion into the utricle, forming the ampullae of the semicircular ducts. The otic vesicle between the developing semicircular ducts and the cochlear duct forms two dilatations. One, the saccule, is located anteriorly, above the developing cochlear duct. The other, the utricle, is located above and behind the saccule. Both ends of each semicircular duct enter the utricle. The end of the endolymphatic duct enlarges into the endolymphatic sac. The elongating cochlear duct begins to spiral. The connection between the saccule and the cochlear duct becomes a narrow tube called the ductus reuniens. Its origin from the cochlear duct is some distance from the blind end of the cochlear duct, which is called the cecum of the cochlear duct. The connection between the utricle and saccule becomes a narrow tube called the utriculosaccular duct. The endolymphatic duct becomes a branch of the utriculosaccular duct. The cochlea elongates and spirals. During the tenth week, the epithelium of the membranous labyrinth begins to differentiate [12, 14]. The primitive, multilayered epithelium becomes a single layer of flat epithelium, except in the sites of the future auditory and vestibular sensory end organs. In these sites, the ingrowing peripheral branches of the eighth cranial nerve induce the epithelium to differentiate into plaques of tall, ciliated neurosensory epithelium that are the rudiments of the neurosensory sensory end organs. The auditory organ is the organ of Corti, located in the basal membrane of the cochlear duct. It first appears in the vestibular end of the cochlear duct and then extends along the full length of the duct. The vestibular system of balance consists of the vertically oriented macula of the saccule, the horizontally oriented macula of the utricle, and the

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three radially oriented cristae of the ampullae of the semicircular ducts. Each of these six organs begins with the differentiation of tall, ciliated, neuroepithelial sensory cells. The epithelium of the inner ear continues to induce the condensation of adjacent mesenchyme around itself to become an increasingly dense envelope of connective tissue, which abuts directly against the membranous labyrinth [12]. This is the precursor of the osseous labyrinth, also called the bony labyrinth or otic capsule. There is no space between the mesenchymal otic capsule and the membranous labyrinth. By the end of the tenth week, the elongation of the cochlear duct has reached nearly two and one-half spirals, the mesenchymal otic capsule has begun to convert into cartilage, and the auditory and vestibular sensory end organs are rapidly differentiating. At the end of the embryonic period (tenth week), most of the major components of the ear are well-developed. However, a few important processes in the histologic maturation of the ear remain for the fetal period: • In the external auditory canal, the canalization of the meatal plug and the concomitant final maturation of the tympanic membrane • In the middle ear, the resorption of the embryonic mesenchyme, the establishment of the tympanic cavity, and the maturation of the secondary tympanic membrane covering the round window • In the inner ear, the final elongation of the cochlear duct to two and three-quarter turns, the formation of the perilymphatic space, the final maturation of the sensory end organs, and the ossification of the ossicles and otic capsule

Histology External Auditory Canal The entire external auditory canal is lined by keratinizing, stratified squamous epithelium, which is continuous with the skin of the pinna. The development of the embryonic and fetal histology of the skin of the external auditory canal is similar to that of the skin elsewhere in the body. The skin of the outer cartilaginous segment of the canal differs from that of the inner osseous segment. Recanalization of the meatal plug begins during the 13th week and is complete by the 18th week [13]. In the outer cartilaginous segment, early keratinization and primitive hair follicles appear in the third month. Sebaceous glands, ceruminous glands, and extensive keratinization appear in the fifth or sixth month in utero. By the time of birth, the lining epithelium of the outer cartilaginous portion of the canal includes numerous hair follicles, sebaceous glands, and ceruminous glands, but no sweat glands (Fig. 31.14). The ceruminous glands are simple, coiled, tubular glands that are thought to be modified apocrine glands

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Fig. 31.14  Epidermal lining of the outer part of the external auditory canal at 6 months postnatal age. The surface is covered by keratinizing, stratified squamous epithelium. Hair follicles and sebaceous glands are confined to the upper half of the lining. Ceruminous glands occupy the lower half of the lining below the hair follicles and sebaceous glands. Lining with these histologic features is characteristic of the outer cartilaginous part of the canal but may extend a short distance into the bony inner part of the canal, as it does in this case. Sweat glands are not present in any part of the external auditory canal (H&E, 2×)

similar to those in the eyelids, axilla, areola, groin, and elsewhere. Their coiled secretory portions form large lobules deep to the hair follicles and sebaceous glands. Their ducts empty into the hair follicles above the mouths of the sebaceous glands and onto the epidermal surface. The epithelium of the ceruminous glands is composed of a luminal layer of large secretory cells with granular, acidophilic cytoplasm and apical secretory granules and an outer layer of myoepithelial cells (Fig.  31.15). The secretory cells may contain yellow-brown pigment. The ceruminous glands continue to mature into childhood and do not become fully functional until puberty or adolescence. Therefore, ceruminous waxy plugs in the external auditory canal are unusual before late childhood, although vernix caseosa may fill the canal in late fetuses and neonates. The lining epithelium of the inner bony portion of the external auditory canal is different from that of the outer ­cartilaginous portion, in that it is thin and lacks dermal papillae and adnexa (Fig. 31.16) [15]. Adjacent to the tympanic membrane, its deep surface is undulating. Early in the fetal period, the stratified squamous epithelium is thin and not yet keratinized (Figs.  31.16 and 31.17). By mid-gestation it is keratinized and may be pigmented (Figs. 31.18, 31.19, 31.20, and 31.21). Before birth, the external auditory canal is filled with amniotic fluid. After birth, it is filled with air. However, routine microscopic sections of the middle and inner ears often do not include the external auditory canal, so these features are usually not appreciated.

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Fig. 31.16  Epidermal lining of the inner part of the external auditory canal near its junction with the tympanic membrane at 15 weeks gestation. The bilayered membrane between the lumen of the external auditory canal (EAC) and the tympanic cavity of the middle ear (TC) is the tympanic membrane. The epithelial membrane beneath the lumen of the external auditory canal is the epidermal lining of the inner part of the external auditory canal. It is a thin, nonkeratinizing, stratified squamous epithelium that lacks hair follicles, sebaceous glands, ceruminous glands, and rete ridges (H&E, 20×)

Fig. 31.15  Higher-power view of the ceruminous glands seen in Fig. 31.14. The ceruminous glands are located deep to the sebaceous glands. They are simple, coiled tubular glands with a single layer of tall, bright pink secretory cells and a single basal layer of myoepithelial cells. The three arrows point to the ceruminous duct, which empties either into the hair follicle above the ostium of the sebaceous gland or into the surface epithelium (H&E, 20×)

Tympanic Membrane The mature histology of the tympanic membrane cannot be appreciated until the meatal plug is removed at 13–18 weeks gestational age. The mature tympanic membrane is composed of three layers, except in the pars flaccida [15]. The outer layer is extremely thin, stratified keratinizing squamous epithelium lacking dermal ridges and adnexa. It is continuous with the lining of the inner bony segment of the external auditory canal, which derives from the ectoderm of the first pharyngeal cleft. In the early fetal period, it is extremely thin and nonkeratinized (Fig.  31.22). By mid-­ gestation, it is thicker and well-keratinized (Fig. 31.23). By 6  months postnatal age, it remains heavily keratinized (Fig.  31.24). Through a process of “auditory epithelial migration,” the keratin and outer layers of the epidermis migrate from the middle to the periphery of the tympanic membrane and then onto the surface of the inner bony segment of the external auditory canal, from which they are ultimately expelled. This process ensures that the epidermal

Fig. 31.17  Higher-power view of the lining of the inner part of the external auditory canal seen in Fig.  31.16. The thin, nonkeratinizing, stratified squamous epidermis lacks a granular layer (H&E, 60×). EAC the lumen of the external auditory canal

surface of the tympanic membrane remains thin, devoid of keratin, and pliable [15]. The middle layer of the tympanic membrane is a connective tissue from the first pharyngeal cleft and pouch; it is composed of two layers, an outer layer of radial fibers and an inner layer of circular fibers, which are well-developed by 15  weeks gestation (Fig.  31.22). The inner surface of the tympanic membrane is continuous with the ­lining of the tympanic cavity. It is derived from the endoderm from the first pharyngeal pouch and consists of squamous epithelium (Figs. 31.23 and 31.24).

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Fig. 31.18  Epidermal lining of the inner part of the external auditory canal near its junction with the tympanic membrane at 23 weeks gestation. The membrane between the lumen of the external auditory canal (EAC) and the tympanic cavity (TC) is the tympanic membrane. The wall of the tympanic cavity is to the right, and the epidermal lining of the external auditory canal is to the left. Unlike the epidermal lining at 15 weeks gestation (seen in Figs. 31.16 and 31.17), this epidermis is thicker, has granular and keratin layers, and has an undulating rather than a flat deep surface. Like the epidermis at 15  weeks gestation, it lacks hair follicles, sebaceous glands, and ceruminous glands. Sweat glands are not present (H&E, 10×)

Fig. 31.19  Higher-power view of the keratinized epidermal lining of the external auditory canal seen in Fig. 31.18. The epidermis may be heavily pigmented, as in this case (H&E, 40×)

Fig. 31.20  Epidermal lining of the inner part of the external auditory canal at 6  months postnatal age, from the same patient depicted in Figs. 31.14 and 31.15. In contrast to the epidermal lining of the outer part of the external auditory canal, the epidermal lining of the inner part of the external auditory canal lacks hair follicles, sebaceous glands, ceruminous glands, and rete ridges. The right edge of the section demonstrates the abrupt change from the ceruminous-bearing epidermis of the outer part of the canal to the nonceruminous-bearing epidermis of the inner part of the canal (H&E, 40×)

Fig. 31.21  Epidermal lining of the inner part of the external auditory canal at its junction with the tympanic membrane at 6 months postnatal age, from the same patient depicted in Figs. 31.14, 31.15, and 31.20. The tympanic membrane separates the lumen of the external auditory canal (EAC) from the tympanic cavity of the middle ear (TC). The deep surface of the epidermal lining near the tympanic membrane changes from flat to undulating. As the epidermis of the canal is reflected onto the inner surface of the tympanic membrane, it becomes thin, and its deep surface again becomes flat. The lumen of the external auditory canal may contain much keratin debris despite the auditory epithelial migration described in the text (H&E, 10×)

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Fig. 31.22  Tympanic membrane at 15 weeks gestation (higher-power view of Fig. 31.16). The tympanic membrane is the upside-down “U” in the lower half of the field. The space below the membrane is the lumen of the external auditory canal (EAC). The space above the membrane is the tympanic cavity of the middle ear (TC). The tympanic membrane is composed of two layers of connective tissue: an outer layer of radial fibers (the side facing the EAC) and an inner layer of circular fibers (facing the TC). The extremely thin epidermis has fallen off the outer surface, and the single layer of endoderm has fallen off the inner surface (H&E, 40×)

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Fig. 31.23  Tympanic membrane at 23 weeks gestation (higher magnification of Fig.  31.18). In contrast to the tympanic membrane at 15 weeks gestation in Figs. 31.16 and 31.22, the epidermis of the tympanic membrane at 23 weeks gestation is thicker and is heavily keratinized. The inner surface is covered by a single layer of flat epithelium, which is an extension of the lining of the tympanic cavity. The two layers of connective tissue forming the middle layer of the membrane are not as distinct as those at 15  weeks gestation or as those seen at 6 months postnatal age, as in Fig. 31.24 (H&E, 40×). EAC lumen of external auditory canal, TC tympanic cavity of middle ear

is complete by 24 weeks [2], and the ossicles are adult size by term. Beginning with the embryonic period and extending into the early fetal period, the tympanic cavity is almost completely filled with embryonic mesenchyme, which surrounds the ossicles and leaves very little lumen (Fig. 31.25). In the early fetal period, the epithelium lining the small cavity consists of simple or slightly stratified flat endoderm. It is thinnest over the inner surface of the tympanic membrane and the convex surfaces of the mounds of mesenchyme that fill the lumen. In these thin areas, it is a single layer of flat cells that lack cilia (Figs. 31.25 and 31.26). It is thickest in the corners and crevices of the cavity, where it becomes irregularly stratified and cuboidal to columnar, with patches of cilia (Figs. 31.25 and 31.27). With increasing gestational age, the subepithelial mesenchyme becomes thicker and more edematous. At Fig. 31.24  Tympanic membrane at 6  months postnatal age (higher-­ mid-gestation, it continues to surround the ossicles and other power view of Fig. 31.21). The epidermis of the outer surface is thinner structures that traverse the tympanic cavity, such as the tenthan that seen at 23 weeks. The two layers of connective tissue are dis- don of the tensor tympani muscle (Figs. 31.28 and 31.29). The tinct. The epithelial lining of the inner surface is similar to that seen at mesenchyme contains variable amounts of extramedullary 23 weeks gestation (H&E, 40×). EAC lumen of external auditory canal, hematopoietic tissue (Figs. 31.30 and 31.31). The epithelium TC tympanic cavity of middle ear remains thin and inconspicuous over the inner surface of the tympanic membrane, where it is a single layer of flat endothelium (see Fig. 31.23), and over the mounds of mesenchyme, Middle Ear where it is a single layer of flat or cuboidal endothelium (see The histology of the middle ear changes significantly during Fig. 31.30). In the corners and crevices, it becomes irregularly the fetal period. Ossification of the ossicles begins at 16 weeks, ­pseudostratified, columnar, and cuboidal, with many patches first in the incus and followed sequentially by the malleus at of cilia (see Fig. 31.31). As pregnancy approaches term, the 16–17 weeks and finally the stapes at 18 weeks. Ossification subepithelial embryonic mesenchyme is reabsorbed, and

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Fig. 31.25  Tympanic cavity at 15  weeks gestation, from the same patient as depicted in Figs. 31.16, 31.17, and 31.22. The tympanic cavity (TC) is a narrow slit behind the tympanic membrane (to the left) and becomes slightly wider at its corner (to the right). The asterisk indicates the root of the tympanic membrane; the double asterisks mark the tympanic ring. The extremely thick layer of edematous and embryonic-­ appearing mesenchyme between the epithelial lining and the cartilage fills the space that would be tympanic cavity if not for the presence of the mesenchyme. The epithelium lining the thick mound of mesenchyme is thin; the epithelium lining the corner is thicker (H&E, 4×)

Fig. 31.26  Higher magnification of Fig. 31.25, depicting the thin epithelial lining of the tympanic cavity at 15 weeks gestation. The thin epithelial lining over the thick mesenchyme is a single layer of squamous cells (H&E, 20×)

Fig. 31.27  Higher magnification of Fig.  31.25 depicting the thicker epithelial lining of the corner of the tympanic cavity (TC) at 15 weeks gestation. The epithelium is stratified cuboidal epithelium with patches of cilia (H&E, 60×)

Fig. 31.28  Tympanic cavity at 23  weeks, from the same patient as depicted in Figs. 31.18, 31.19, and 31.23. Shown from top to bottom are an unidentified ossicle (O), stapes (S), edge of oval window (OW), tendon of the tensor tympani muscle (TT), lumen of the external auditory canal (EAC), tympanic cavity (TC), and tympanic ring (TR). The tympanic cavity remains a narrow slit behind the tympanic membrane but becomes slightly wider at the corner of the cavity. The unidentified ossicle, the tendon of the tensor tympani muscle, and the stapes are buried in the mass of mesenchyme that fills most of the tympanic cavity. The space to the right of the edge of the oval window is the periotic space (the cistern) of the vestibule. The connective tissue of the tympanic membrane inserts into the groove of the tympanic ring (H&E, 4×)

the  lumen becomes larger. Less mesenchyme surrounds the ­ossicles and other structures (Figs. 31.32 and 31.33). The mesenchyme disappears from the tympanic side of the s­ econdary tympanic membrane in the round window. The secondary tympanic membrane attains its adult appearance of a thin, trilayered membrane: a single layer of squamous epithelium derived from that of the tympanic cavity on the tympanic side, a middle layer composed of a thin sheet of connective tissue, and an inconspicuous single layer of periotic reticular squamous cells on the vestibular side (Figs. 31.32 and 31.34). Extramedullary

hematopoiesis gradually diminishes from the embryonic mesenchyme. The epithelium continues to become thinner and less conspicuous (see Figs. 31.32 and 31.33). By the last week of pregnancy or the neonatal period, the mesenchyme com-

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Fig. 31.29  Thick mesenchyme filling the tympanic cavity (TC) at 24 weeks gestation. Shown are the tendon of the tensor tympani muscle (TT), the foot plate of the stapes (S) in the oval window, and the radial ligament (arrows). The space to the right of the oval window is the periotic space (the cistern) of the vestibule (H&E, 4×)

Fig. 31.30  Higher magnification of Fig. 31.28, depicting the thin epithelial lining of the tympanic cavity at 23  weeks gestation. The thin epithelial lining over the thick mesenchyme is simple cuboidal epithelium, which often falls off owing to mild autolysis, as seen in this section. Mild extramedullary hematopoiesis is present (H&E, 60×)

Fig. 31.31  Higher magnification of Fig. 31.28, depicting the thicker epithelial lining of the corner of the tympanic cavity at 23  weeks. Pseudostratified, ciliated columnar epithelium lines the corners and crevices of the tympanic cavity. Mild extramedullary hematopoiesis is present (H&E, 40×)

Fig. 31.32  Tympanic cavity at 38 weeks gestation. The membrane separating the lumen of the external auditory canal (EAC) from the tympanic cavity (TC) is the tympanic membrane. The handle of the malleus is attached to its inner surface. The irregular common bony wall between the tympanic cavity and the cavity of the vestibule is depicted. Toward the end of pregnancy, the masses of edematous mesenchyme that filled the tympanic cavity earlier in pregnancy have mostly disappeared, leaving a wideopen tympanic cavity. Only thin remnants of the mesenchyme remain covering the bony wall in the upper right and between the round and oval windows. The radial ligament at the bottom (arrows) attaches the edges of the head of the stapes to the inner edge of the oval window. Only a thin layer of mesenchyme surrounds the handle of the malleus, in contradistinction to the masses of mesenchyme in which the ossicles were buried earlier in pregnancy (H&E, 4×). RW round window enclosed by the secondary tympanic membrane, ST scala tympani, SV scala vestibuli

pletely disappears, including that surrounding the ossicles and the tendon of the tensor tympani muscle. The lining epithelium lies directly upon the underlying bone and the ossicles, and the mucosa becomes a mucoperiosteum (Figs. 31.35 and 31.36). The ossicles become fully mobile for the first time, allowing auditory signals, which previously could reach the inner ear only by bone conduction, to now reach the inner ear through the tympanic membrane and ossicles. The tall, pseudostratified, ciliated columnar endothelium in the corners and crevices may increase in thickness (Fig.  31.37). During the late fetal and neonatal periods, the mucosa around the opening of the Eustachian tube may form complex rugae. The normal epithelium in this area may contain a few mucus-producing

cells (Fig. 31.38), but the presence of serous or mucous glands or large numbers of goblet cells anywhere in the mucosa of the tympanic cavity is abnormal and is probably a metaplastic response to excessive amniotic fluid or infection. Before birth, the tympanic cavity contains amniotic fluid and debris. After birth, it contains air.

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Fig. 31.33  Another view of the tympanic cavity (TC) at 38 weeks gestation, from the same patient depicted in Fig.  31.32. The membrane separating the lumen of the external auditory canal (EAC) from the tympanic cavity is the tympanic membrane. The cavity is wide open. A thin layer of edematous mesenchyme remains covering the bony wall only at the upper end of the cavity. The articular head and limbs of the stapes (S) are visible, along with the radial ligament (arrow), which anchors the foot plate of the stapes in the oval window. The mesenchyme covering the stapes is very thin. The incudostapedial diarthrodial joint is also visible (H&E, 2×)

Fig. 31.34  The secondary tympanic membrane at 38 weeks gestation. The secondary tympanic membrane encloses the round window. The space above and to the left of the round window is the tympanic cavity (TC). The space below and to the right of the window is the scala tympani (ST), which ends blindly at the round window. The secondary tympanic membrane is a delicate, trilayered sandwich composed of a thin middle layer of connective tissue covered on its tympanic surface by a single layer of squamous epithelial cells (a continuation of the epithelial lining of the tympanic cavity) and covered on its vestibular side by a single layer of squamous periotic reticular cells (a continuation of the reticular lining of the scala tympani). Earlier in pregnancy, the tympanic surface of the secondary tympanic membrane is buried under a mass of edematous mesenchyme (H&E, 10×)

Fig. 31.35  The tympanic cavity (TC) at 6 months postnatal age. The lumen of the external auditory canal is in the lower left. The membrane in the lower left that separates the lumen of the external auditory canal from the tympanic cavity is the tympanic membrane. In the lower right, the structure separating the scala vestibuli (SV) from the scala tympani (ST) is the basal turn of the cochlear duct. The cavity is wide open. The edematous mesenchyme is completely absent. The stapes (S, with limbs tangentially cut) and the tendon of the tensor tympani muscle (TT) are devoid of a mesenchymal covering. The radial ligament (arrows) attaches the foot plate of the stapes in the oval window. Note the common bony wall between the tympanic cavity and the perilymphatic spaces of the inner ear (H&E, 2×)

Fig. 31.36  The mucoperiosteum of the tympanic cavity at 6 months postnatal age. This is a higher magnification of Fig. 31.35, depicting the epithelial lining of the tympanic cavity. A single layer of squamous cells and a very thin layer of subepithelial connective tissue cover the bony wall of the tympanic cavity. This epithelial lining is called the mucoperiosteum (H&E, 20×)

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Fig. 31.37  The thicker epithelial lining of the corners and crevices of the tympanic cavity at 6 months postnatal age. This is a higher magnification of Fig. 31.35, showing the pseudostratified, ciliated cuboidal and columnar epithelium, which covers the recesses of the tympanic cavity (H&E, 40×)

Eustachian Tube The short segment of the tympanic end of the Eustachian tube is supported by a bony wall. The remainder of the tube is supported by a wall of elastic cartilage. The entire length is lined by pseudostratified, ciliated columnar epithelium. Its pharyngeal cartilaginous end includes seromucous glands and lymphoid organs referred to as Gerlach’s tubal tonsils. Like all lymphoid tissues, these do not develop germinal centers or mature plasma cells until 2–6 weeks postnatal age.

Inner Ear Otic Capsule Ossification of the otic capsule begins only after the cartilaginous capsule reaches adult size at 16  weeks. Multiple sites of ossification appear during the 16th week. Ossification of the otic capsule is complete by the 24th week [2]. The otic capsule is seen in many sections of the cochlea, vestibule, and semicircular canals. It is a thin rim of very dense, often basophilic, bone, which is distinct from the petrous bone in which it is embedded. Perilymphatic Space The formation of the perilymphatic space of the inner ear occurs during fetal life [11, 12, 14]. As early as the tenth week, the mesenchyme of the inner surface of the cartilaginous osseous labyrinth becomes vacuolated and is trans-

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Fig. 31.38  The epithelial lining of the tympanic cavity near the opening of the Eustachian tube at 6 months postnatal age from the same patient as the previous three figures. The mucosal lining of the cavity near the opening of the Eustachian tube is rugated, and the pseudostratified, ciliated columnar epithelium may contain occasional mucus-­ producing cells. Mucusproducing cells should not be seen in other parts of the cavity (H&E, 60×)

formed into a reticulum called the perilymphatic reticulum. The reticulum is gradually resorbed, which results in the formation of a space, the perilymphatic space, between the osseous labyrinth and the membranous labyrinth. This space contains the perilymphatic fluid. This process begins in the scala tympani, then extends into the scala vestibuli, and finally spreads throughout the vestibule and into the semicircular canals. The scala tympani and scala vestibuli are completely devoid of reticulum by the end of the 12th postfertilization week (Figs. 31.39 and 31.40). The process may be completed in the vestibule and then in the ­semicircular canals at any gestational age from 20 weeks to term, but the perilymphatic reticulum may never be completely resorbed from parts of the vestibule and semicircular canals in some individuals. The periotic reticulum forms the perichondrium of the cartilaginous otic capsule and presumably the periosteum of the osseous otic capsule. Several of the figures of the vestibule and semicircular canal demonstrate various degrees of resorption of the perilymphatic reticulum.

 ochlear Aqueduct and Periotic Duct C The development of the periotic duct and cochlear aqueduct is related to the development of the perilymphatic space and the otic capsule and is completed during the fetal period. The primitive periotic duct develops in conjunction with the inferior cochlear vein and its bony canal and Hyrtl’s fissure

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Fig. 31.39  Cross section of the cochlea at 27 weeks gestation. Three sections of the spiral bony cochlear canal are shown around the central bony core of the cochlea, the modiolus (M), like the leaves of a three-­leaf clover. The two on either side of the modiolus in this image are nearly perfect cross sections, which are mirror images of each other. They are near the base of the cochlea. The one at the top of the modiolus is an oblique section. It is toward the apex (cupola) of the cochlea. The bony spiral laminae (arrows) extend like arms from the edge of the modiolus into the lumen of the cochlear canal. The modiolus is traversed by multiple channels to accommodate fibers of the cochlear nerve (CN). Lacunae at the edge of the modiolus contain the spiral cochlear ganglia (SG). At the bottom of the modiolus, the multiple channels and spaces empty into a large canal within the bone, the internal auditory canal. Small branches of the cochlear nerve stream through these fenestrae into the canal and join together to form larger branches of the cochlear nerve, one of which is seen in the canal (CN). The space at the bottom right edge is the saccule of the membranous labyrinth within the vestibule. Small fibers of the vestibular nerve (VN) stream from the saccular otolithic macula through fenestra in the bony wall of the vestibule to join together to form larger branches of the vestibular nerve within the internal auditory canal, one of which is seen in the canal (VN). Each of the three sections of the cochlear canal is surrounded by a thin rim of very dense bone, the otic capsule of the cochlea. Note its distinction from the surrounding petrous bone. The medial ends of the bony walls end in the modiolus. At the bottom of the modiolus, they end in the bony wall of the external auditory canal. The saccule is surrounded by a similar thin bony wall, the otic capsule of the vestibule. The bony cochlear canal is divided into three channels, which are clearly depicted in the two perfect cross sections. The central triangular channel is the cochlear duct (also called the scala media, SM) of the membranous labyrinth. It contains endolymph and ends blindly at both ends, one at the apex of the cochlea and the other in the vestibular cistern below the saccule. The channel above the cochlear duct is the scala vestibuli (SV), which ends in the vestibular cistern near the oval window. The channel below the cochlear duct is the scala tympani (ST), which ends blindly at the round window. The details of the cross section of the canal to the right of the modiolus are seen in the following four figures (H&E, 2×)

(the tympanomeningeal fissure). The periotic duct forms a direct communication between the scala tympani component of the perilymphatic space and the subarachnoid space. Hyrtl’s fissure forms a direct continuation between the tympanic cavity and the subarachnoid space and is separate from the cochlear aqueduct and periotic duct. In early fetuses, subarachnoid hemorrhage in the posterior fossa commonly

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Fig. 31.40  Higher magnification from Fig.  31.39 depicting the bony cochlear canal and the membranous cochlear duct at 27 weeks gestation. The modiolus is to the left. The bony wall of the canal is the distinct, thin, blue rim of dense bone around the lateral wall of the canal. It curves over the top of the canal and then medially to join the modiolus. It curves around the bottom of the canal and off the bottom edge of the picture. In Fig. 31.39, it can be seen to fuse with the junction of the modiolus and the wall of the internal auditory canal. The bony spiral lamina (arrow) arises from the modiolus and extends like an arm halfway across the equator of the canal, bisecting its lumen. The periosteum along the inner surface of the lateral wall of the bony canal is thickened to form a pyramidal cushion of stroma. This cushion is the spiral ligament (SL). Its apex, the crest of the spiral ligament, extends into the lumen of the canal along its equatorial plane, opposite the spiral lamina. A membrane, the basal membrane, extends from the tip of the spiral lamina to the crest of the spiral ligament. This is the base or floor of the cochlear duct. A delicate membrane, Reissner’s membrane, extends from the upper surface of the tip of the spiral lamina diagonally upward and inserts into the spiral ligament near its upper end. The triangular duct formed by the basal membrane, Reissner’s membrane, and the spiral ligament is the cochlear duct (H&E, 4×). ST scala tympani, SV scala vestibuli

spreads into the perilymphatic space of the inner ear through the periotic duct and into the tympanic cavity through Hyrtl’s fissure. By 24 weeks, Hyrtl’s fissure is closed, and the direct communication between the subarachnoid space and the tympanic cavity is lost. The direct communication between the subarachnoid space and the perilymphatic space through the periotic duct becomes permanent. Therefore, after 24  weeks, subarachnoid hemorrhage in the posterior fossa may spread through the periotic duct into the inner ear but not into the tympanic cavity [2].

Cochlear Duct During the 11th week, the cochlea and cochlear duct attain their final length, and their spirals consist of two and three-­ quarter turns [11, 14]. Growth and maturation of the organ of Corti begins at the base of the cochlear duct as a thickening in the epithelium in one side of the round cochlear duct. This side of the wall becomes flat and forms the basal membrane or floor of the mature cochlear duct. The tall, columnar epithelium covering the floor of the developing organ of Corti

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Fig. 31.41  Higher magnification of Fig.  31.40 depicting the cochlear duct (also called the scala media, SM) at 27 weeks gestation. The two thin lamina of bone, upper and lower, that constitute the spiral lamina are apparent at this magnification. The basal membrane stretches from the tip of the spiral lamina to the crest of the spiral ligament (SL). The stria vascularis (StV) forms the surface of the spiral ligament between the insertion of Reissner’s membrane (RM) above and the spiral prominence, the bump immediately above the crest of the spiral ligament below. Note the thin, dense otic capsule (H&E, 10×). ST scala tympani, SV scala vestibuli

continues to become thicker and develops two longitudinal ridges separated by a valley—a larger inner ridge nearer the center of the spiral and a smaller outer ridge nearer the lateral wall of the spiral. The tall epithelium of the larger, inner ridge secretes the gelatinous tectorial membrane that covers the top of the epithelium. The outer, smaller ridge will become the organ of Corti. By 14 weeks, the round cochlear duct has attained its mature triangular shape in cross section. The floor of the duct is the basal membrane containing the organ of Corti. The wall opposite the basal membrane is the vestibular or Reissner’s membrane, which is the roof of the cochlear duct. Reissner’s membrane and the basal membrane join to form an acute angle, the apex of the triangle, toward the center of the cochlear spiral. The lateral wall of the triangle is formed by the stria vascularis and the spiral ligament. The space above Reissner’s membrane is the scala vestibuli. The space below the floor is the scala tympani. By 14 weeks, the inner and outer hair cells and the pillar cells are present. The inner margin of the tectorial membrane is attached to the spiral limbus, and the free outer edge is embedded in the cilia of the inner and outer hair cells. By 16 weeks, the epithelial cells of the inner ridge degenerate and disappear, forming the inner spiral sulcus. The tectorial membrane stretches across the inner spiral sulcus. Cells between the inner and outer hair cells degenerate and disappear, forming the tunnel of Corti. The development of the organ of Corti approaches completion as early as 20 weeks gestation [14], when primitive fetal response to auditory stimuli received through bone conduction is first detectable. The development of the organ of Corti may not be complete

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Fig. 31.42  Higher magnification of Fig. 31.41 depicting the organ of Corti (OC) at 27 weeks. In routine sections, the organ of Corti is typically severely fragmented. This is one of the best-preserved examples found among the many cases in our material. The tectorial membrane (TM) often survives. The two bony lamina of the spiral lamina are clearly depicted. Tiny nerves pass from the neurosensory cells of the organ of Corti through the spiral lamina between its two laminae to enter the spiral cochlear ganglia in the modiolus. See Fig. 31.8 for more details (H&E, 20×). ISS inner spiral sulcus, OSS outer spiral sulcus, SM scala media, ST scala tympani, SV scala vestibuli

until 25  weeks or later in some fetuses. This discrepancy may be explained in part by the fact that maturation begins at the base of the cochlear duct and progresses toward the apex of the spiral, so that the time of complete maturation will depend upon where along the spiral the section is obtained (see Figs. 31.4 and 31.40). The sophistication with which the fetus responds to auditory stimuli increases simultaneously with the increasing maturation of the organ of Corti as gestational age advances. The organ of Corti is often autolyzed in routine sections even in otherwise well-preserved specimens (Figs. 31.41 and 31.42). Reissner’s membrane is delicate and trilayered. Its inner cochlear surface is covered by simple squamous epithelium derived from the ectoderm of the otic vesicle. The inconspicuous middle layer is a wisp of connective tissue. The outer vestibular surface is covered by simple squamous epithelium derived from the periotic reticulum. The histology of all free walls of the membranous labyrinth in the saccule, utricle, and semicircular ducts is similar to that of Reissner’s membrane (see Figs.  31.40, 31.41, and 31.42). The spiral ligament is a thick, curved, pyramidal pad of stroma on the inner surface of the lateral bony wall of the cochlea. It forms the lateral wall of the cochlear duct. It is thickened periosteum and may be a specialized remnant of the periotic reticulum (see Figs. 31.40 and 31.41). The stria vascularis is a unique vascularized, stratified epithelial membrane on the surface of the spiral ligament. Its surface is a single layer of dark, eosinophilic, cuboidal cells referred to as dark cells or marginal cells. Deep to the dark cells is a thick layer of haphazardly arranged, more lightly stained

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Fig. 31.43  The stria vascularis at 27 weeks gestation, from the same specimen shown in Figs. 31.39, 31.40, 31.41, and 31.42. The stria vascularis is a layer of vascularized, stratified epithelium. The surface is covered by a single layer of dark, eosinophilic cuboidal cells referred to as dark cells or marginal cells. Deep to the dark cells is a thick layer of haphazardly arranged, more lightly stained polygonal cells referred to as light cells or basal cells. There are numerous capillaries in the layer of light cells. The stroma of the spiral ligament lies beneath the stria vascularis (H&E, 60×)

polygonal cells referred to as light cells or basal cells. There are numerous capillaries in the layer of light cells. Epithelial membranes are characteristically not vascularized. The stria vascularis may be the only vascularized epithelial membrane in the human body and is thought to contribute to the maintenance of the volume and ionic makeup of the endolymph (Figs.  31.41 and 31.43). The nerves from the sensory hair cells pass through channels in the center of the spiral lamina (see Fig. 31.42) and join the spiral ganglion in the periphery of the modiolus. Nerves from the spiral ganglion pass through bony channels and join to form larger nerves in the modiolus (Fig. 31.44). Branches of the cochlear and vestibular nerve join to form the eighth cranial nerve in the internal auditory canal and exit the posterior wall of the petrous bone through the internal auditory meatus (Fig. 31.45).

Vestibular System The delicate membranes of the saccule and utricle are usually destroyed in routine sections of even well-preserved specimens of the middle and inner ears (Fig.  31.46). Occasionally these structures are reasonably intact in routine sections (Figs. 31.47 and 31.48). The periotic reticulum of the saccule, utricle, and semicircular ducts may be incompletely resorbed in some fetuses of mid to late gestational ages (Figs. 31.47, 31.48, and 31.49). The progression of maturation of the utricular and saccular maculae and the ampullary cristae is similar [11, 14]. They first appear at 10 weeks, and their maturation is complete by 14  weeks. They differ in that the maculae are

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Fig. 31.44  The spiral cochlear ganglion at 27  weeks gestation. The ganglion is in the left lower corner. Nerve fibers pass through numerous fenestrae in the bone to join a branch of the cochlear nerve in the upper left corner (H&E, 10×)

Fig. 31.45  The internal auditory meatus at 27 weeks, from the same specimen as Fig.  31.44. The vestibular nerve (VN) and the cochlear nerve (CN) join together in the internal auditory canal to form the eighth cranial nerve (VIII N.), which passes through the canal and exits through the internal auditory meatus in the posterior wall of the petrous bone (H&E, 2×)

ellipsoid plaques covered by gelatinous otolithic membranes (see Figs. 31.9, 31.10, 31.11, 31.50, 31.51, 31.52, 31.53, 31.54, and 31.55), whereas the cristae are narrow transverse ridges capped by gelatinous tongue-like membranes called the cupola, without otoliths (see Figs. 31.9, 31.12, 31.13, 31.53, and 31.56). At 10 weeks, the maculae and cristae are ­composed of a thick layer of two types of epithelial cells: ciliated neurosensory cells and supportive cells. The tall epithelial cells surrounding the sensory organs secrete a gelatinous membrane in which the cilia of neurosensory cells are embedded. In the maculae, this

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Fig. 31.46  The vestibule (V) and its junction with the cochlea at 24 weeks gestation. SV in the mid vestibule indicates the scala vestibuli adjacent to the oval window; SV in the lower end of the vestibule is the scala vestibuli at the junction of the basal turn of the cochlea with the vestibule, also the site of the scala tympani (ST). At the middle left is the footplate of the stapes, which is anchored in the oval window by the radial ligament (short arrows). The delicate membranes of the saccule and utricle of the membranous labyrinth, which should be seen in the vestibule, are not seen because they were destroyed during removal and sectioning of the specimen as almost always occurs. The membranous labyrinth is more well-preserved in the following figures. The thin bony otic capsule of the vestibule is distinct from the petrous bone. The junction of the cochlea with the vestibule is at the bottom of the vestibule, where the long arrow points to the spiral lamina and basal membrane of the cochlear duct. Reissner’s membrane is almost completely collapsed. The crest of the spiral ligament is clearly seen (H&E, 2×)

Fig. 31.48  Vestibule at 27 weeks gestation. A partial cross section of the cochlear canal and duct are seen at the lower right. The otic capsule of the cochlea and vestibule is clearly visible. The saccule (S) is distinguished from the utricle (U) by its location closest to the cochlea. The otolithic macula of the utricle is the thickened plaque in its wall closest to the saccule. The otolithic macula of the saccule is the thickened area in its wall closest to the utricle. Nerve fibers from the two otolithic maculae pass through fenestra in the bone and join together to form the branch of the vestibular nerve (VN) seen here in a canal within the otic capsule of the vestibule (H&E, 2×). ST scala tympani, StV stria vascularis, SV scala vestibuli

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Fig. 31.47  The vestibule and its junction with the lateral semicircular canal at 23  weeks gestation. In contrast to the previous figure, this membranous labyrinth is unusually well preserved. The otic capsule is distinct from the surrounding petrous bone. The periotic reticulum of the ampulla of the lateral semicircular duct (A) has not been completely resorbed. The bidirectional arrow marks the opening of the ampulla into the utricle (U). The walls of the utricle and the saccule (S) are partially disrupted. A fragmented membrane of unknown origin lies between the utricle and the saccule (H&E, 2×). C crista of the ampulla

membrane is flat and covers the entire surface of the ovoid organ. In the cristae, it is a tongue-shaped membrane called the cupola, which arises from the apex of the full length of the crista and extends like a valve flap into the lumen. A thick layer of stromal cells and nerves beneath the epithelium adds to the thickness of the organs. By the 14th week, the free surfaces of the membranes of the otolithic saccular and utricular maculae contain basophilic crystals of calcium carbonate and protein called otoliths, so that the maculae are referred to as otolithic membranes (see Figs.  31.11, 31.52, 31.54, and 31.55). The microscopic appearance of the otoliths varies. In some sections, they are nearly invisible (Fig.  31.52). In others, they are small and round, or in some others, they may be irregular and oblong (Fig.  31.54). In still others, the otolithic ­membranes may be fragmented and lie in the lumen of the labyrinth, in which case they are easily mistaken for infectious organisms (Fig.  31.55). In contrast, the membranes of the cristae, the cupola, are devoid of otoliths and lengthen to stretch across almost the entire lumen of the ampullae (see Figs. 31.12, 31.13, and 31.56). The semicircular canals may be seen in perfect cross sections (Figs. 31.57 and 31.58) or in oblique sections of various degrees (not illustrated). The bony wall of the canal, the otic capsule, is distinct from the surrounding petrous bone. The duct is attached to the inner surface of the bony canal by perilymphatic reticulum. The canal is lined by the simple squamous epithelium that is derived from the perilymphatic reticulum and serves as peri-

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Fig. 31.49  Higher magnification of the left lower wall of the saccule in Fig. 31.48, depicting the periotic reticulum of the saccule at 27 weeks gestation. The lumen of the saccule is in the upper right. The otic capsule is in the lower left. The lining of the saccule is a single layer of flat epithelium common to the entire membranous labyrinth. The periotic reticulum fills the space between the epithelial lining of the saccule and the otic capsule (H&E, 20×)

Fig. 31.50  An unusually well-preserved saccule (S) at 27 weeks gestation. The basal turn of the cochlear canal and duct appear in the upper right corner. The otic capsule of the vestibule and cochlea is unusually well delineated. The delicate free wall of the saccule, like Reissner’s membrane, is trilayered: The middle layer is connective tissue. The inner surface is covered by simple squamous epithelium characteristic of the membranous labyrinth. The outer surface is covered by the simple squamous reticular epithelium of the periotic reticulum. The opposite wall is secured to the inner surface of the otic capsule by the thick saccular otolithic membrane. Fine peripheral nerve filaments from the neurosensory hair cells of the macula pass through the subepithelial stroma of the macula and then through multiple fenestra in the otic capsule to join together in the internal auditory canal to form the branch of the vestibular nerve (VN) seen here (H&E, 4×)

Fig. 31.51  Saccular otolithic macula at 27 weeks gestation. This is a higher-magnification view of the same specimen seen in Fig.  31.50. Nerve fibers from the macula pass through fenestra in the bony otic capsule (H&E, 10×)

Fig. 31.52  Higher magnification of Fig. 31.51, depicting the saccular otolithic macula at 27 weeks gestation. A gelatinous membrane (short arrow) covers the surface of the epithelial layer of the macula, which consists of several layers of vertically oriented cells (long arrow). Even in this relatively well-preserved specimen, the epithelial cells are mildly autolyzed. Some of the hair cells can be recognized; their cilia are embedded in the deep surface of the gelatinous membrane. A basement membrane lies beneath the epithelium. The cells of the subepithelial stroma are predominantly horizontally oriented, and the stroma contains numerous tiny nerve fibers and capillaries. Coarse basophilic crystals, called otoliths or otoconia, are usually seen on the surface of the gelatinous membrane but are not visible here. See also Fig. 31.11 for details and compare with Fig. 31.54 (H&E, 60×)

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Fig. 31.53  Higher magnification of Fig. 31.47, depicting the junction of the ampulla (A) of the lateral semicircular duct with the utricle (U) at 27 weeks gestation. The arrow marks the opening between the ampulla and the utricle. The utricular otolithic macula (M) is the thick blue lining of the utricle in the right lower corner (H&E, 4×). C crista of the ampulla

Fig. 31.54  Higher magnification of Fig. 31.53 depicting the utricular otolithic macula at 27 weeks gestation. The arrow on the left indicates the gelatinous membrane covering the surface of the epithelium. The arrow on the right indicates the sensory epithelial membrane. Numerous coarse basophilic crystals, the otoliths, are embedded in the surface of the gelatinous membrane, unlike the otolithic macula seen in Fig. 31.52. See also Fig. 31.11 for details and the text for a discussion of variability in the appearance of the otoliths (H&E, 40×)

Fig. 31.55  Fragments of an otolithic membrane lying free in the vestibular cistern at 27 weeks gestation. The deeply basophilic otoliths in this section vary from coarse and elongate in the upper layer to small and round in the lower layer. These are easily mistaken for infectious organisms. Compare with the otoliths in Fig. 31.52, which are nearly invisible, and those in Fig. 31.54, which are smaller and rounder than those in this section. See Fig. 31.11 for more details (H&E, 40×)

Fig. 31.56  Higher magnification of Fig. 31.53, depicting a cross section of the crista of the lateral semicircular ampulla at 27 weeks gestation. The stromal core of the crista, in the upper right corner, contains numerous small nerve fibers and capillaries. The strip of sensory epithelium forms a cap over the tip of the crista. It is mildly autolyzed. The gelatinous cupola is the eosinophilic, tongue-shaped flap that is attached to the sensory epithelium and extends toward the lower left. For details, see Fig. 31.13. Compare with the otolithic macula of the saccule and utricle in Figs. 31.11, 31.13, 31.52, and 31.54 (H&E, 20×)

osteum. The duct is lined by simple squamous epithelium derived from the ectodermal membranous labyrinth. Various amounts of perilymphatic reticulum may remain in the canal.

Endolymphatic Duct The endolymphatic duct arises from the utriculosaccular duct and passes through the vestibular aqueduct of the vestibular otic capsule and into the dura. Its distal end is a large, dilated

sac in the dura of the posterior wall of the petrous bone (see Figs. 31.1, 31.2, 31.4, 31.5, and 31.7). The narrow duct is lined by simple epithelium, which varies from squamous to cuboidal (Fig. 31.59). The large sac has a very irregular wall. The epithelial surface is thrown into numerous rugae and fjord-like inlets (Fig.  31.60). The epithelium ranges from simple low cuboidal to irregularly stratified cuboidal (Fig. 31.61).

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Fig. 31.57  Semicircular canal (SC) and semicircular duct (SD) at 27 weeks. The thin, bony otic capsule of the canal is distinct from the petrous bone in which it is embedded. Some periotic reticulum remains around the duct and the opposite side of the canal. The canal is lined by simple squamous periotic reticulum (H&E, 4×)

Fig. 31.58  Higher magnification of Fig. 31.57, depicting the semicircular duct at 27 weeks gestation. The duct is anchored to the wall of the canal by periotic reticulum. It is lined by simple squamous epithelium (H&E, 10×)

Fig. 31.59  Endolymphatic duct at 27 weeks. The duct in this section is located in the dura. It is lined by simple squamous and cuboidal epithelium. The irregular epithelial structure adjacent to the duct at the bottom right is the beginning of the endolymphatic sac (H&E, 40×)

Fig. 31.60  Endolymphatic sac at 27 weeks gestation. The sac is a dilatation of the distal end of the duct. Its wall is extremely irregular, with numerous rugae and fjord-like inlets (H&E, 10×)

Special Considerations Examination of the Ear in the Perinatal Period Why should perinatal pathologists include the examination of the ears in perinatal autopsies? This question is frequently asked. At a time when perinatal autopsy pathology is forced to compete for resources with the increasing demands of pediatric surgical pathology, the tendency is to spend fewer resources on perinatal pathology, not more. Several factors challenge that tendency. First, most of us humans die before we are born. This is a difficult concept, but many studies show that between 60% and 80% of pregnancies end in embryonic or fetal death.

Second, the ear is often involved in the major categories of causes of intrauterine diseases and deaths, such as congenital anomalies with or without chromosomal or gene defects, infections (either amniotic fluid infection sequence or transplacental infections), perfusion abnormalities (either maternal placental or fetal placental), and congenital tumors [16]. The intrauterine abnormalities of the ear often cause neonatal hearing loss in those who survive. Third, the rapidly advancing fields of fetal medicine, imaging, and surgery impart an increasing clinical relevance to perinatal pathology. If perinatal pathologists are to keep abreast of or lead our clinical colleagues in the study of perinatal diseases, we need to perform more perinatal autopsies, not fewer and, in more detail, not less—including the examination of the ears.

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Fig. 31.61  Higher magnification of Fig. 31.60, depicting the epithelium of the endolymphatic sac at 27 weeks gestation. The sac is lined by simple and irregularly stratified cuboidal epithelium (H&E, 60×)

Examples of abnormalities of the ear that accompany perinatal death or threaten the hearing of those who survive are numerous [16–19]. The histologic continuum of normal aspiration of amniotic fluid, excessive aspiration of amniotic fluid, and amniotic fluid infection sequence is one of the most common findings in sections of the ear. Because the tympanic cavity is in direct continuity with the amniotic cavity through the Eustachian tube, and because the course of the Eustachian tube in fetuses and neonates is nearly horizontal, a small amount of amniotic debris is often seen in sections of the middle ear of fetuses and neonates. Normally, this small amount of amniotic debris is removed, as it is from the lung, but fetal distress may result in an excessive amount of amniotic debris in the tympanic cavity, similar to that occurring in the lung. Therefore, an excessive amount of amniotic debris in the tympanic cavity should suggest significant intrauterine fetal distress. In extreme situations, the amniotic debris may persist and incite a foreign-body inflammatory reaction resulting in polypoid nodules attached to the ossicles or the wall of the tympanic cavity. Glandular and squamous metaplasia of the lining epithelium may accompany retention of amniotic debris. This is more likely in neonates than in fetuses. In cases of amniotic fluid infection sequence, acute inflammatory debris and infectious agents— including cocci, bacilli, fungi, and spirochetes—may be found in the tympanic cavity. Acute inflammation of the embryonic mesenchyme of the tympanic cavity, the mucosa, or the subjacent bone indicates invasive otitis media. A normal amount of extramedullary hematopoiesis should not be mistaken for otitis media. Blood-borne transplacental infections may affect the inner and middle ears of fetuses and newborns. Cytomegalovirus commonly involves the stria vascularis of the cochlear duct (Figs.  31.62 and 31.63) and the sensory organs of the saccule and utricle. Hearing loss is one of the most frequent clinical findings in congenital cytomegalovi-

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Fig. 31.62  Congenital cytomegalovirus infection of the cochlear duct at 37  days postnatal age in a baby born at 33  weeks gestation. The infected cells are in the stria vascularis overlying the stroma of the spiral ligament (H&E, 20×)

Fig. 31.63  Higher magnification of Fig.  31.62, depicting congenital cytomegalovirus infection of the cochlear duct at 37 days postnatal age. The infection has severely damaged the stria vascularis. Compare Fig. 31.43 (H&E, 60×)

rus infection. Rubella, Herpes simplex infection, toxoplasmosis, and Listeria also may infect the middle or inner ears. H. simplex and Listeria infections can be either ascending or transplacental. Fetal distress may cause acute hemorrhage in the middle and inner ears, as it does in other organs. Acute abruptio placenta with subependymal, intraventricular, and subarachnoid hemorrhage is especially likely to be associated with severe hemorrhage of the middle and inner ears. One cause for the hemorrhage in the ears could be hypoxic-ischemic damage to the tissues of the middle and inner ears. Another reason could be the direct communication between the subarachnoid space and the inner ear through the periotic duct throughout fetal and neonatal life and the direct communication between the subarachnoid space and the tympanic cavity

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through Hyrtl’s fissure prior to 24 weeks, when the fissure closes. These two communications allow subarachnoid hemorrhage in the posterior fossa to flow from the subarachnoid space into the inner and middle ears [7]. Rarely, congenital neuroblastoma, leukemia, teratoma, or Langerhans cell histiocytosis may involve the middle or inner ears. Many congenital anomalies affect the ears, but special care in removing and sectioning the temporal bone may be required to demonstrate many of these malformations.

Removing the Temporal Bone The reliable demonstration of normal and pathologic histology depends upon the use of effective techniques in removal and sectioning of the temporal bone. Haphazard removal is likely to yield unsatisfactory results due to fragmentation,

crushing, incomplete removal, and poor orientation. Several methods have been advocated, which vary from simple methods to complex methods that are labor-intensive and expensive. Each institution can choose a technique that best fits its goals and resources. Four incision lines are needed to remove the petrous bone (Fig. 31.64). The first passes through the temporalis muscle, separates the soft tissue from the outside of the temporal bone, and transects the external auditory canal (see Fig. 31.3). The second separates the anteromedial end of the petrous bone from the side of the sella turcica and is nearly parallel to the first incision. It should be placed as close to the wall of the sella turcica as possible. The third cuts through the roof of the petrous bone anterior to the area of the tegmen tympani, as close to the anterior vertical wall of the middle fossa as possible. It is extended into the underlying bone. It is perpendicular to the first two. The fourth is a cut along the infe-

2

3

Tegmen tympani

Petrous ridge

Internal auditory meatus

1

Arcuate eminence

2

Subarcuate fossa Superior horizontal wall of petrous bone

Jugular fossa

Petrous ridge

Ostium of cochlear aqueduct

Posterior vertical wall 4

1 Ostium of vestibular aqueduct

Fig. 31.64  Four incision lines for removal of the petrous bone. The first lateral incision does not pass through bone but rather through the soft tissue of the scalp. It is depicted by the black line through the temporalis muscle following the curve of the cranium and forms a flap of the scalp including the temporalis muscle, subcutaneous tissue, skin, and pinna, which can be folded laterally away from the cranium. The separation of this flap is extended downward and under the cranium anteriorly, where the external auditory canal is located and transected. See Fig. 31.3, which shows this first incision in the frontal plane, represented by the black line labeled “1,” which passes through the temporalis muscle, curves under the cranium, and transects the outer cartilaginous portion of the external auditory canal, which is located on the anterior surface of the removed petrous bone near its lateral end. The second medial incision passes through bone and separates the anteromedial end of the petrous bone from the side of the sella turcica and clivus. It should be made flush against the side of the sella and cli-

vus. The third anterior incision is through the boney floor of the middle cranial fossa as far anterior as possible. It should include all of the tegmen tympani. It extends through the lateral wall of the cranium, which was exposed by incision number “1.” Incision number “3” connects incisions “1” and “2.” The fourth posterior boney incision is along the inferior margin of the posterior vertical wall of the petrous bone. Laterally, incision number “4” also extends through the lateral wall of the cranium. All incisions are extended until the petrous bone is free and can be removed. Compare this figure with Figs. 31.1 and 31.2. Note that the incision lines in this figure include all the surface landmarks in Fig. 31.1 and all the components of the inner, middle, and external ears indicated in Fig.  31.2. Color code: dark green, petrous bone; light green, squamous portion of temporal bone; orange, sphenoid bone; blue, occipital bone; violet, parietal bone; tan areas, open sutures and minor fontanelles

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rior edge of the posterior wall of the petrous bone to include the orifice of the cochlear aqueduct at the superior edge of the jugular fossa. This is extended anteriorly toward the third cut. The four cuts are extended until the bone is free. This method provides a complete petrous bone, including the ­distal external auditory canal, tympanic membrane, middle and inner ears, ostia of the external auditory meatus and cochlear and vestibular aqueducts, and a portion of the Eustachian tube. After removal, the specimen should be thoroughly fixed. After fixation, the amount of decalcification varies. Ossification begins in the third month and continues throughout fetal life and into infancy. Therefore, the amount of decalcification should be carefully monitored and tailored to each specimen. Specimens from first-trimester and early second-trimester fetuses may need no decalcification. After decalcification, the specimen is bisected by a horizontal section through the external and internal acoustic meatuses in a plane parallel to the plane of the superior surface of the petrous bone (Fig. 31.65). An alternative is a vertical section perpendicular to the superior surface of the petrous bone, parallel to the petrous ridge. Additional cuts parallel to either choice can be made, depending on the size of the specimen. The instruments used in removal and sectioning of the specimens are important. Unossified or minimally ossified specimens should be removed with a scalpel blade. Because of unossified sutures and minor fontanelles, specimens may crumble even with the use of scalpels unless special care is taken in their removal and sectioning. Partially or extensively ossified specimens require a saw for removal. Adult Stryker saws are usually too large for use in fetuses and newborns. Pediatric Stryker saws are more appropriate and are easily available. Small, sharp bone scissors can be used, but they may cause crushing and fragmentation of the specimen. The use of chisels should be avoided. Sectioning of the appropriately decalcified specimen can be accomplished with a s­ calpel blade or one of the several available longer, razor-like blades. More extensive and detailed examinations enter the realm of research and are beyond the scope of this chapter. For instance, if the entire length of the Eustachian tube is desired, a more invasive excision will be required. If the best preservation for histology and immunohistochemistry is desired, the unfixed specimen can be cut into thin serial slices by precision saws. These slices require minimal fixation; various fixatives can be used, and minimal decalcification is required. These slices can be serially microscopically sectioned and stained with a variety of stains and immunohistochemical techniques. The microscopic sections can be utilized for three-dimensional reconstruction to document the details of congenital malformations. All these histologic procedures are complicated by multiple artifacts. The fixed whole specimen can be studied by

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Petrous ridge Internal auditory meatus

Petrous ridge Superior horizontal wall

Fig. 31.65  Removed petrous bone showing the preferred plane of section. Often the removed petrous bone includes excess lateral wall of the cranium composed of the squamous part of the temporal bone and possibly the inferior parietal bone. Trim off this excess bone through the squamous part of the temporal bone, flush with the superior horizontal wall of the petrous bone. Locate the internal auditory meatus on the posterior vertical wall of the petrous bone and the cut end of the external auditory canal located on the underside of the anterior surface of the specimen (not shown in this figure). The cut passes through the internal auditory meatus and the external auditory canal. Start the cut on the posterior surface parallel to the petrous ridge and through the internal auditory meatus. Continue the cut forward in the horizontal plane parallel to the superior surface of the petrous bone and through the external auditory canal. The cut bisects the petrous bone in the horizontal plane. Embed the two halves with the cut surfaces down. The histologic sections will include the internal auditory canal cut longitudinally, a longitudinal cut of the cochlea, the vestibule, the middle ear, the tympanic membrane, the external auditory canal (cut longitudinally), and various cuts of the semicircular canals. Multiple microscopic sections of both halves will include all of the structures depicted in the previous photomicrographs. The black line is the horizontal plane of section. The red is the remnants of the temporalis muscle fibers attached to the cranium. Note the trimmed edge of the squamous part of the temporal bone

CT microscopy [20] or MR microscopy [21]. These techniques avoid the artifacts of histologic procedures and provide excellent anatomic and pathologic detail of the middle and inner ears.

References 1. Crelin ES. Anatomy of the newborn: an atlas. Philadelphia: Lea and Febiger; 1969. p. 56–66. 2. Scheuer L, Black S.  Developmental juvenile osteology. Oxford: Elsevier; 2007. p. 67–84. 3. Netter FH. Atlas of human anatomy. 4th ed. Philadelphia: Saunders; 2006. Plates 9–12 and 92–97. 4. Florin TA, Ludwig S, editors. Netter’s pediatrics. Philadelphia: Elsevier; 2011. p. 187–9. 5. Standring S, editor. Gray’s anatomy. 39th ed. Philadelphia: Elsevier; 2005. p. 649–80. 6. Skrzat J, Wrobel A, Walocha J.  A preliminary study of three-­ dimensional reconstruction of the human osseous labyrinth from microcomputed tomography scans. Folia Morphol (Warsz). 2013;72:17–21.

354 7. Spector GJ, Lee D, Carr C, Davis G, Schnnettgoecke V, Strauss M, Rauchbach E. Later stages of development of the periotic duct and its adjacent area in the human fetus. Laryngoscope. 1980;90:1–31. 8. Ash LM, Ibrahim M, Schipper MJ, Mukherji SK.  The utricular macula: qualitative and quantitative analysis using 3-T imaging. J Comput Assist Tomogr. 2010;34:93–7. 9. Naganuma H, Tokumasu K, Hashimoto S, Okamoto M, Yamashina S.  Three-dimensional analysis of morphological aspects of the human saccular macula. Ann Otol Rhinol Laryngol. 2001;110:1017–24. 10. Naganuma H, Tokumasu K, Hashimoto S, Okamoto M, Yamashina S.  Three-dimensional analysis of morphological aspects of the human utricular macula. Ann Otol Rhinol Laryngol. 2003;112:419–24. 11. O’Rahilly R, Müller F. Human embryology and teratology. 3rd ed. New York: Wiley-Liss; 2001. p. 471–86. 12. Streeter GL. The histogenesis and growth of the otic capsule and the contained periotic tissue spaces in the human embryo. Contrib Embryol. 1918;7:5–54. 13. Wright CG.  Development of the human external ear. J Am Acad Audiol. 1997;8:379–82.

D. S. Huff 14. Tanaka O.  Ear. In: Nishimura H, editor. Atlas of human prenatal histology. Tokyo: Igako-Shoin; 1983. p. 64–75. 15. Wenig BM, Michaels L.  The ear and temporal bone. In: Mills SE, editor. Histology for pathologists. Philadelphia: Lippincott Williams and Wilkins; 2012. p. 399–432. 16. deSa DJ, Gilbert-Barness E.  The ear. In: Gilbert-Barness E, editor. Potter’s pathology of the fetus, infant and child. 2nd ed. Philadelphia: Mosby Elsevier; 2007. p. 2181–206. 17. deSa DJ. Pathology of neonatal intensive care: an illustrated reference. London: Chapman and Hall; 1995. p. 132–6, 150. 18. deSa DJ. Mucosal metaplasia and chronic inflammation in the middle ear of infants receiving intensive care in the neonatal period. Arch Dis Child. 1983;58:24–8. 19. deSa DJ.  Infection and amniotic aspiration of middle ear in stillbirths and neonatal deaths. Arch Dis Child. 1973;48:872–80. 20. Lane JI, Witte RJ, Driscoll CLW, Camp J, Robb RA.  Imaging microscopy of the middle and inner ear. Part I: CT microscopy. Clin Anat. 2004;17:607–12. 21. Lane JI, Witte RJ, Henson OW, Driscoll CLW, Camp J, Robb RA. Imaging microscopy of the middle and inner ear. Part II: MR microscopy. Clin Anat. 2005;18:409–15.

Part VIII Musculoskeletal and Connective Tissues

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Bone Linda M. Ernst

Introduction The skeletal system is developing throughout fetal life and continues to develop well into postnatal life. The changes in the skeleton in embryonic and fetal life are dramatic, and the study of the normal histological changes in bone development can be important for understanding abnormalities encountered in such disease states as intrauterine growth restriction, skeletal dysplasia, and metabolic disturbances. This chapter reviews the normal development of the skeletal system and the histological components of the bone throughout gestation.

Embryology The bones of the skeletal system are derived from paraxial mesoderm, lateral plate mesoderm, and neural crest cells. After gastrulation, the mesodermal cells that migrated from the primitive streak begin to form three condensations lateral to the notochord: the paraxial mesoderm, the intermediate mesoderm, and the lateral plate mesoderm. The most medial, known as the paraxial mesoderm, is destined to contribute to the axial skeleton, the skeletal muscles of the trunk and limbs, and the dermis of the skin [1, 2]. In the vertebrate embryo, the paraxial mesoderm begins to further divide into adjacent, rounded somitomeres, which first appear in the cranial region and extend as paired structures along the length of the embryo [1]. It should be noted that the existence of somitomeres in human embryos is not universally accepted [3]. From the eighth somitomere to the caudal end of the embryo, the somitomeres continue to show further segmentation into somites, which are paired, block-like condensaL. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected]

tions of primitive mesenchymal cells clearly seen in human embryos. The first somites form about day 20 and continue to form in a cranial-to-caudal direction until approximately day 30. About 42–44 pairs of somites are formed, with eventual regression of the caudal-most somites [1]. Somites are extremely important in the developing embryo because they establish the initial segmentation of the body and provide the mesenchymal tissue for skeletal and muscular elements of most of the body (Table 32.1). The somitic mesoderm is further subdivided in the fourth week of gestation into the ventromedial portion known as the sclerotome, which will form vertebrae and ribs, and the more lateral segment known as the dermomyotome, which will give rise to the precursors of the skeletal muscles and the dermis of the skin. Within each sclerotome, the ventral cells migrate to surround the notochord and form the vertebral bodies. The dorsal cells surround the neural tube and become the vertebral arches and spinous processes. Cells in the lateral portion of the sclerotome will form the transverse processes and ribs. The formation of ribs begins on day 35, and the first endochondral ossification in the ribs is seen at about 6 weeks of development [2]. The sternum develops from the lateral plate mesoderm, specifically from structures known as the sternal bars, which are two paired, longitudinal mesenchymal condensations in the ventrolateral body wall. The sternal bars begin to fuse around the seventh week of development in a cranial-to-­caudal fashion. Ossification centers begin to form on day 60, with multiple ossification centers forming in a cranial-to-­caudal progression [2]. The skeletal elements of the limbs are derived from the parietal layer of the lateral plate mesoderm, which gives rise to the limb buds at the end of the fourth week. The mesenchyme of the limb bud is under the inductive influence of an ectodermal thickening at the tip of the bud known as the apical ectodermal ridge [1, 2], which arises from the ectodermal ring. The inductive influences of the apical ectodermal ridge promote proliferation and differen-

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Table 32.1  Skeletal and muscular derivatives of somites Somite pairs 1–4, occipital 5–12, cervical

Skeletal derivatives Occipital skull Bones of nose, eyes, inner ears Portion of occipital bone

Muscular derivatives Extrinsic ocular muscles Muscle of tongue

25–29, lumbar

Lumbar vertebrae

30–34, sacral 35–37, coccyx

Sacral vertebrae

Cervical paraspinal muscle Upper limb muscles Thoracic paraspinal muscle Intercostal muscles Abdominal wall musculature Upper limb muscles Lumbar paraspinal muscle Abdominal wall musculature Lower limb muscles Sacral paraspinal muscle

Coccyx

Paracoccygeal muscle

13–24, thoracic

Cervical vertebrae Thoracic vertebrae Ribs Sternum

Fig. 32.1  Metacarpal bone ossification in late first trimester gestation. Metacarpal bones of the hand show central primary ossification centers, which can be recognized by the proliferating cartilaginous tissue showing early ossification (arrow) (hematoxylin and eosin (H&E), 4×)

tiation of the limb bud. Bone formation within the long bones occurs in the center of the diaphysis, with an initial condensation of mesenchymal cells that differentiate into chondrocytes. Formation of a primary center of ossification (Figs.  32.1 and 32.2) begins when stem cells in the perichondrium produce a layer of osteoblasts that fabricate a thin layer of woven bone on the surface of the condensation of chondrocytes. These chondrocytes then become organized into progressive layers of resting, proliferating, prehypertrophic, and hypertrophic chondrocytes. Once the nutrient vessel of the long bone enters this cartilaginous model, near the end of the embryonic period, osteoblasts derived from perivascular stem cells enter the region and

Fig. 32.2  Metacarpal bone ossification in late first trimester gestation. Higher-power view of a primary ossification center in the central area of a metacarpal bone (arrow). Note the thin rim of woven bone over the surface of the cartilaginous condensation (H&E, 10×)

begin replacing the chondrocytes and forming the mineralized trabeculae of the bone centrally. The cartilaginous tissue is resorbed, and ossification by osteoblasts then spreads toward the ends of the bone. Most of the long bones of the extremities have primary ossification centers by the midtrimester, but they retain cartilaginous caps at their epiphyseal ends (Fig. 32.3). Ossified cortical bone is formed from the primary bone collar in all the long bones by birth, and ossification centers can be seen in the distal femoral epiphyses by approximately 35 weeks gestation (Fig. 32.4) and in the proximal tibial epiphyses at term (Fig.  32.5, Table 32.2) [4]. The presence or absence of primary ossification centers can be used as a reliable method to determine the gestational age or gestational age range of a fetus. Therefore, radiologic examination is recommended for fetal autopsy, not only to demonstrate any fetal skeletal abnormalities but also to help establish the gestational age. Accurate assessment of bone age can assist with interpretation of fetal growth abnormalities and estimation of timing of intrauterine fetal demise. The bones of the skull and face develop from neural crest cells, paraxial/somitic mesoderm, and the pharyngeal arches [1, 2]. The membranous neurocranium, composed of flat bones that surround the brain, is derived from both neural crest cells and somitic mesoderm. The bones ossify by membranous ossification and show bony spicules radiating from the central area to the periphery of the bone. Growth occurs by resorption of the central area and new bone formation at the periphery. The chondrocranium, the bones at the base of the skull, forms from cartilaginous precursors derived from the neural crest and paraxial mesoderm. Ossification of the bones of the chondrocranium occurs by endochondral ­ossification. Lastly, the viscerocranium, the bones of the face, develops from the first two pharyngeal arches and

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Fig. 32.3  Long-bone ossification at 20 weeks gestation. Low-power view of the long bone showing central ossification and formation of trabeculae in the medullary cavity. The end of the bone consists of a rounded, cartilaginous cap that allows for continued growth (H&E, 2×)

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Fig. 32.4  Radiograph of the lower limb of a 38-week fetus. Note ossification of the distal femoral epiphysis (arrow)

n­eural crest cells. These bones ossify by membranous ossification.

Histology Histological Types of the Bone The bone is a mixture of organic and inorganic materials. The main organic components are the bone’s cellular elements and the organic matrix, which consists predominantly of type I collagen. The inorganic component of the bone is a calcium-deficient variant of hydroxyapatite [5]. The mineralization of tissue occurs by distinct methods in each of the two histological types of the bone, woven bone and lamellar bone [5]. Woven bone predominates in fetal life. Lamellar bone is composed of parallel-oriented type I collagen fibers. It is typically produced more slowly than woven bone and is seen in mature bone. Using routine H&E stains, lamellar bone is eosinophilic and appears less cellular than woven bone. The lacunae are more evenly distributed along the lamellae, which consist of parallel-oriented collagen fibers that can be highlighted by examination under polarized light (Fig. 32.6).

Fig. 32.5  Radiograph of the lower limb of a 40-week fetus. Note ossification of the distal femoral epiphysis and proximal tibia epiphysis (arrows)

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Table 32.2  Ossification in the fetus Body region Head Mandible Occipital bone Maxilla Temporal bone Sphenoid Nasal bone Frontal bone Bony labyrinth Neck Hyoid bone Thorax Clavicle Scapula Ribs, 2nd–11th Ribs, 1st and 12th Vertebra Cervical, thoracic, lumbar, upper sacral arches Cervical, thoracic, lumbar, and upper sacral bodies Fourth sacral arches Fifth sacral body Coccyx Odontoid process of axis Pelvis Ilium Ischium Pubis Upper extremity Humerus Proximal humeral epiphysis Radius Ulna Metacarpals, second and third Metacarpals, first, fourth, and fifth Distal phalanges Proximal phalanges, second and third Proximal phalanges, first and fourth Proximal phalanx, fifth Middle phalanges, second, third, and fourth Middle phalanx, fifth Lower extremity Femur Distal femoral epiphysis Tibia Proximal tibial epiphysis Fibula Metatarsals, second and third Metatarsals, first, fourth, fifth Calcaneus Talus Cuboid Distal phalanx, first Distal phalanges, second, third, fourth Distal phalanx, fifth Proximal phalanges Middle phalanx, second Middle phalanx, third Middle phalanx, fourth Middle phalanx, fifth Data from Siebert [4]; with permission from Elsevier

Week 7 8–10 8 9 9–14 10 9–10 7–20 28–32 7 8–9 8–9 10 9–12 10–12 19–25 13–28 37–40 17–20 9 16–17 21–28 8 37–42 8 8 9 10–12 9 9 10 11–12 12 13–16 8–9 35–40 8–9 40 9 9 10–12 21–29 24–32 40 9 10–12 13–14 13–14 20–25 21–26 29–32 33–36

Woven bone is characterized by type I collagen fibers arranged in an irregular meshwork. In embryonic and fetal development, woven bone predominates because woven bone can be produced and resorbed more quickly than the more organized lamellar bone. In H&E-stained sections, woven bone appears hypercellular, with lacunae arranged in a haphazard pattern. Examination with polarized light reveals collagen fibers arranged in nonparallel, random directions (Fig. 32.7).

Structure of the Bone Periosteum  The periosteum is a thin connective tissue layer that covers the outer surface of the bone (Figs.  32.8 and 32.9). It is composed of a fibrocollagenous outer layer and a more cellular inner layer known as the cambium layer. The cells in the cambium layer are osteoprogenitor cells. Cortical Bone  The cortex of long bones of the fetus is composed predominantly of dense woven bone (see Figs. 32.8 and 32.9). In the first trimester, ossification centers are just beginning to form, and cortical bone is just beginning to be present by the end of the first trimester. In the midtrimester, cortical bone is seen and is thickest in the middle of the diaphysis. The cortex thickens over gestation but may be very thin to absent near the growth plate (Fig. 32.10). Medullary Canal  The bone of the medullary cavity is not composed of solid bone but is formed by trabeculae of bone, with bone marrow spaces interspersed between the bony trabeculae (Figs.  32.11 and 32.12). In developing bone, the trabeculae are composed of woven bone, but in mature bone, the bony trabeculae are comprised of lamellar bone. Vascular Supply  The blood supply of bone is derived from nutrient arteries, metaphyseal and epiphyseal arteries, and periosteal vessels [5]. The nutrient arteries consist of one or two large vessels per long bone, which enter at the diaphysis, course through cortical bone (Fig. 32.13), and provide blood supply to the medullary cavity and the inner two thirds of the cortical bone. Branches of the nutrient artery can be seen within the medullary cavity and ultimately give rise to the capillaries and venous vessels in the medullary cavity (Fig. 32.14). The metaphyseal and epiphyseal arteries enter the bone near the ends through small apertures [5] and provide the blood supply to those regions, including the important area of active endochondral ossification. Lastly, the outer cortex and periosteum are supplied by the periosteal vessels [5]. The venous drainage of the bone is via the medullary sinusoids, which drain to a central venous sinus and ultimately to the nutrient veins [5].

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a

a

b

b

Fig. 32.6  Lamellar bone. In the center of each field is a bony trabecula composed of lamellar bone from an adult specimen. (a) Note the organized, parallel-oriented collagen fibers and osteocytes with small, pyknotic nuclei. (b) The parallel-oriented collagen fibers are highlighted by examination under polarized light (a, b H&E, 20×)

Fig. 32.7  Woven bone. In the center of each field is cortical bone from a 20-week fetus. (a) Woven bone generally appears more disorganized than lamellar bone. Note the haphazard arrangement of collagen fibers and osteocytes. (b) The haphazardly oriented collagen fibers are highlighted by examination under polarized light (a, b H&E, 20×)

Cell Types in Bone

amphophilic to basophilic cytoplasm (Fig.  32.15). The nucleus is typically positioned eccentrically away from the surface of the matrix and shows prominent nucleoli. Perinuclear halos representing the active Golgi apparatus can be seen [5]. As an individual osteoblast produces matrix, it surrounds itself with matrix, inhabits a small lacunar space within the matrix, and becomes an osteocyte (see Fig.  32.15). In mature lamellar bone, osteocyte nuclei within the lacunae are generally smaller and more pyknotic than osteoblasts. In addition, the mature osteocyte is oriented in the direction of its surrounding lamellae. In the fetus, because most of the bone is woven bone, osteocytes are more plump and their orientation is more random (compare Figs. 32.6 and 32.7). Although not well-appreciated in H&E-stained sections, osteocytes have numerous long

Osteoprogenitor cells are committed mesenchymal cells that will form osteoblasts under the influence of proper growth and transcription factors [5]. In fetal bone, they are most easily recognized in the periosteum as spindle-shaped to stellate-­shaped cells in the inner/cambium layer (see Figs. 32.8 and 32.9). Osteoblasts are the main cell involved in bone production. Osteoblasts produce and arrange the organic matrix of the bone and are responsible for the regulation of the mineralization of the bone [5]. Osteoblasts are present on the surfaces of bone, and their appearance varies with their metabolic activity. When inactive, they have a benign spindle-cell appearance similar to fibroblasts. When active, osteoblasts become larger and adopt a polygonal shape with abundant

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Fig. 32.8 Periosteum and cortical bone at 20  weeks gestation. Medium-power view of the surface of a fetal long bone, showing the periosteum composed of two layers. The top layer is slightly less cellular than the bottom layer. Note also the woven bone of the cortical bone and deeper medullary cavity (H&E, 10×)

L. M. Ernst

Fig. 32.9  Periosteum and cortical bone at 36  weeks gestation. This image of the surface of a fetal long bone shows the periosteum, which appears slightly thicker than in the midtrimester. The top layer shows dense collagenous tissue, and the lower cambium layer is more cellular. Note also the woven bone of the cortical bone and the deeper medullary cavity (H&E, 10×)

cytoplasmic processes that extend throughout the canaliculi of the matrix. These cellular processes are important for intercellular communication between osteocytes and with osteoblasts and aid the osteocyte function of maintaining the bone. Osteoclasts are the cells responsible for resorption of bone by release of their lysosomal contents in the resorption pit [5]. Osteoclasts appear as multinucleated, large cells on the surface of the bone (Fig. 32.16). As the matrix is resorbed by the osteoclast, a small resorption pit is formed (known as Howship’s lacuna), within which the osteoclast is found.

Types of Bone Formation Endochondral Ossification Histological assessment of the growth plate is important to determine that bone formation and growth is occurring normally. Routine samples of the long bones may be difficult to acquire, but a sample of the rib that includes the ­costochondral junction typically demonstrates the growth plate nicely. The morphologic changes as chondrocytes mature at the growth plate are demonstrated by the five poorly delimited, recognizable histological zones (Figs.  32.17, 32.18, 32.19, 32.20, and 32.21) [5–7], listed here from the distal end of the bone toward the ossified central bone: 1. Zone of resting cartilage: Resting chondrocytes are present in lacunae in this zone. 2. Zone of proliferating cartilage: Chondrocytes multiply in this region and appear as aggregates of small cells in small longitudinal columns.

Fig. 32.10  Cortical bone near the growth plate at 20 weeks gestation. This image is a low-power view of the end of a long bone. The woven bone of the cortical bone (arrow) becomes extremely thin to nonexistent at the growth plate (H&E, 4×)

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Fig. 32.12  Medullary bone at 36 weeks gestation. Note the partially ossified trabeculae of woven bone with central areas composed of basophilic cartilaginous tissue. Osteocytes are present within the woven bone. Bone marrow spaces are present between the trabeculae and appear congested in this image (H&E, 10×) Fig. 32.11  Medullary bone at 20 weeks gestation. Note the partially ossified trabeculae of woven bone rimmed by osteoblasts. Osteocytes are present within the woven bone. Bone marrow spaces are present between the trabeculae (H&E, 20×)

Fig. 32.13 Nutrient vessel at diaphysis. This is cortical bone from a term fetus. Note the cross sections of a branch of the nutrient vessel (arrows) perforating the cortex. Nutrient vessels provide the blood supply of the inner bone marrow cavity on the right (H&E, 20×)

3. Zone of hypertrophic cartilage: Chondrocytes in this zone remain arranged in columns, but the cytoplasm expands and appears clear on H&E-stained sections. The cytoplasm of these chondrocytes is loaded with glycogen. 4. Zone of cartilage degeneration with cartilage matrix mineralization. In this zone, the chondrocytes lose their

Fig. 32.14  Vessel within the medullary cavity. This is a high-power image of the medullary cavity of the bone in a term fetus. Note the longitudinal section of a thin-walled capillary within the medullary cavity (arrows) (H&E, 60×)

nuclei and appear as “degenerated” cells, although the exact nature of these morphologic changes is not clear [6, 7]. The cartilage matrix also begins to undergo mineralization in this zone. 5. Zone of woven bone formation on mineralized cartilage matrix. In the zone closest to the metaphysis, woven bone is deposited on the mineralized cartilage matrix.

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Fig. 32.15  Woven bone with prominent osteoblasts. This is cortical bone from a 20-week fetus. The osteoblasts line the undersurface of the bone (arrowheads). Osteoblasts have abundant cytoplasm, and the nucleus is oriented away from the surface of the bone. Note the plump-­ appearing osteocytes within lacunae of the woven bone. An osteoclast is present on the opposite surface of the bone (arrow) (H&E, 60×)

Fig. 32.16  Woven bone with osteoclast. This is cortical bone from a 20-week fetus. Note the multinucleated osteoclast (arrow) within the resorption pit (H&E, 40×)

Fig. 32.17  Growth plate at 20  weeks gestation. Low-power view of the five zones of endochondral ossification: (1) zone of resting cartilage; (2) zone of proliferating cartilage; (3) zone of hypertrophic cartilage; (4) zone of cartilage degeneration with cartilage matrix mineralization; and (5) zone of woven bone formation on mineralized cartilage matrix (H&E, 4×)

Fig. 32.18  Growth plate at 20 weeks gestation. Higher-power view of the physeal area. At the top of the image, chondrocytes appear as aggregates of small cells in small, longitudinal columns. In the middle, the hypertrophic zone, the cytoplasm of the chondrocytes balloons and appears clear, with abundant glycogen. In the lower zone, the cartilage matrix appears basophilic and acellular, with thin, pink rims of ossification (H&E, 20×)

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Fig. 32.20  Growth plate at 36 weeks gestation. Low-power view of the five zones of endochondral ossification: (1) zone of resting cartilage; (2) zone of proliferating cartilage; (3) zone of hypertrophic cartilage; (4) zone of cartilage degeneration with cartilage matrix mineralization; and (5) zone of woven bone formation on mineralized cartilage matrix (H&E, 4×)

Fig. 32.19  Growth plate at 36  weeks gestation. Low-power view of the end of a long bone in the third trimester. The overall organization of the growth plate is unchanged (H&E, 2×)

Membranous Ossification As mentioned in the embryology section of this chapter, the bones of the skull and face ossify by membranous ossification, a process of bone formation along a fibrous lining membrane. In the fetus, the lining membrane is typically the periosteum, which contains osteoprogenitor cells that differentiate into osteoblasts. These osteoblasts directly deposit bone matrix. Membranous bone formation is also important in cortical bone formation of the long bones, so nearly all bones undergo membranous ossification [5]. Growth occurs by resorption of the central area and new bone formation at the periphery.

Fig. 32.21  Growth plate at 36 weeks gestation. Higher-power view of the physeal area. At the top of the image, chondrocytes appear as aggregates of small cells in small, longitudinal columns. In the middle, the hypertrophic zone, the cytoplasm of the chondrocytes balloons and appears clear, with abundant glycogen. In the lower zone, the cartilage matrix appears basophilic and acellular, with thin, pink rims of ossification (H&E, 10×)

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References 1. Sadler TW, Langman J. Langman’s medical embryology. 11th ed. Philadelphia: Wolters Kluwer Lippincott Williams & Wilkins; 2010. 2. Schoenwolf GC, Larsen WJ. Larsen’s human embryology. 4th ed. Philadelphia: Elsevier/Churchill Livingstone; 2009. 3. O’Rahilly R, Müller F.  The skeletal system and the limbs. In: Human embryology & teratology. 3rd ed. New  York: Wiley-Liss; 2001. p. 357–94. 4. Siebert J.  Perinatal, fetal and embryonic autopsy. In: Gilbert-­ Barness E, editor. Potter’s pathology of the fetus, infant, and child. 2nd ed. Philadelphia: Mosby Elsevier; 2007. p. 712.

L. M. Ernst 5. Rosenberg AE, Roth SI. Bone. In: Mills SE, editor. Histology for pathologists. 4th ed. Philadelphia: Lippincott Williams & Wilkins; 2012. p. 85–106. 6. Gilbert-Barness E. Osteochondrodysplasia—constitutional diseases of bone. In: Gilbert-Barness E, editor. Potter’s pathology of the fetus, infant, and child. 2nd ed. Philadelphia: Mosby Elsevier; 2007. p. 1836–97. 7. Yang SS.  The skeletal system. In: Wigglesworth JS, Singer DB, editors. Textbook of fetal and perinatal pathology. 2nd ed. Malden: Blackwell Science; 1998. p. 1039–82.

33

Skeletal Muscle Linda M. Ernst and Patrick Shannon

Introduction Fetal movements in utero are an important part of the developmental process, and normal fetal movements are a marker of fetal well-being. The muscular system of the fetus provides the contractile apparatus necessary to move the extremities, trunk, and head and produce fetal breathing movements. Primary abnormalities in the development of skeletal muscle or secondary conditions that reduce fetal movements can produce pathologic changes in skeletal muscle and lead to abnormalities in the development of other organ systems. Therefore, an understanding of the normal development and histological appearance of skeletal muscle is necessary to interpret fetal pathology. This chapter reviews the developmental changes in skeletal muscle from the embryo to the term fetus/ neonate.

Embryology Skeletal muscle is mesodermal in origin [1–3]. After gastrulation, the mesodermal cells that migrated from the primitive streak begin to form three condensations lateral to the notochord: the paraxial mesoderm, the intermediate mesoderm, and the lateral plate mesoderm. The most medial condensation, known as the paraxial mesoderm, is destined to contribute to the axial skeleton, the skeletal muscles of the trunk and

L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected] P. Shannon Department of Pathology and Laboratory Medicine, University of Toronto, Mount Sinai Hospital, Toronto, ON, Canada e-mail: [email protected]

limbs, and the dermis of the skin [2, 3]. In the vertebrate embryo, the paraxial mesoderm begins to divide further into adjacent, rounded somitomeres, which first appear in the cranial region and extend as paired structures along the ­ length of the embryo [2]. It should be noted that the existence of somitomeres in human embryos is not universally accepted [4]. By electron microscopy, the somitomeres consist of whorled mesenchymal cells without further differentiation [1]. The skeletal muscles of the cervical and cranial structures are derived from the first seven cranial pairs of somitomeres, which become the mesenchyme of the branchial arches [2]. From the eighth somitomere to the caudal end of the embryo, the somitomeres continue to show further segmentation into somites, which are paired, block-like condensations of primitive mesenchymal cells clearly seen in human embryos. The first somites form around day 20 and continue to form in a cranial-to-caudal direction until approximately day 30. Approximately 42–44 pairs of somites are formed, with eventual regression of several of the most caudal somites [2]. Somites are extremely important in the developing embryo because they establish the initial segmentation of the body and provide the mesenchymal tissue for skeletal and muscular elements of most of the body (see Fig. 32.1). The somitic mesoderm is further subdivided in the fourth week of gestation into the ventromedial portion known as the sclerotome, which will form vertebrae and ribs, and the more lateral segment known as the dermomyotome, which will give rise to the precursors of the skeletal muscles and dermis of the skin. Within the dermomyotome, two muscle-forming areas are identified, the ventrolateral compartment and the dorsomedial compartment. Cells in the ventrolateral compartment migrate into the parietal layer of the lateral plate mesoderm and form infrahyoid, abdominal wall, and limb muscles. Cells in the dorsomedial area form the muscles of the back, shoulder girdle, and intercostal spaces [2, 3].

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During early embryogenesis, some primitive mesenchymal cells differentiate into “pre-myoblasts,” so-named because these cells lack definitive histological features to distinguish them from the other mesenchymal cells, but they have begun differentiation toward myogenesis. Once these cells achieve some phenotypic features of muscle, such as the presence of desmin and myofilaments in the cytoplasm, they are known as myoblasts [1]. These are spindle-shaped, elongated cells with fine,

granular cytoplasm and ovoid nuclei with prominent nucleoli [5]. Muscle fibers are formed by the fusion of multiple muscle cells, and mature muscle fibers are recognized by their peripheral nuclei and organization of filaments into sarcomeres. The first step toward the formation of muscle fibers begins around 7 weeks gestation with the formation of primary myotubes, formed by the fusion of embryonic myoblasts. Primary myotubes have both immature-appearing cytoplasm and nuclei. They are characterized by chains of large central nuclei, each with prominent central nucleoli (Figs. 33.1, 33.2, 33.3, and 33.4), surrounded by cytoplasm containing myofibrils, glycogen, and mitochondria [1].

Fig. 33.1  Embryonic skeletal muscle. Note the formation of primary myotubes characterized by longitudinally arranged nuclei with open chromatin and nucleoli. The cytoplasm is eosinophilic, and cross-­ striations are not seen (hematoxylin and eosin (H&E), 20×)

Fig. 33.2  Embryonic skeletal muscle. A higher-power view of the primary myotubes with long chains of immature-appearing nuclei with prominent nucleoli. The cytoplasm appears clear to filamentous (H&E, 40×)

Fig. 33.3  Embryonic skeletal muscle. High-power view of longitudinally arranged primary myotubes. Note nuclear detail and cytoplasm of primary myotubes (H&E, 60×)

Fig. 33.4  Embryonic skeletal muscle. High-power view of a cross section of primary myotubes. The nuclei are placed centrally within the myotube, and central clearing is seen (H&E, 60×)

Early Histogenesis of Muscle

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Fig. 33.5  Fetal skeletal muscle at 17  weeks gestation. Low-power view of fascicular arrangement of muscle fibers. Note the interstitial connective tissue containing nerves and blood vessels. The muscle fibers shown are seen in cross section, and most of the fibers have peripherally placed nuclei (H&E, 10×)

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Fig. 33.6  Fetal skeletal muscle at 17 weeks gestation. Medium-power view of muscle fascicles. The muscle fibers have a fairly uniform polygonal shape, and most of the fibers have peripherally placed nuclei. Occasional central nuclei are seen at this gestational age (H&E, 20×)

Fetal Muscle Development In the early fetal period, secondary myotubes begin to form in a manner similar to primary myotubes. Secondary myotube development is characterized by the fusion of fetally derived myoblasts, which are larger in diameter than embryonic myoblasts and contain more myofilaments. Therefore, physiologically, secondary myotubes begin to show contractility [5]. Histologically, secondary myotubes also show central nuclei similar to primary myotubes. Secondary myotube formation continues throughout the early midtrimester but begins to decline at midgestation [5], although some myoblasts continue to divide and fuse until 30 weeks gestation [1]. Secondary myotubes mature into recognizable myofibers in the midtrimester (Figs.  33.5, 33.6, and 33.7). With further maturation, myofibers develop an increasing number of myofibrils with easily visible cross-striations, and more nuclei become peripherally located along the muscle fiber (Figs. 33.8, 33.9, and 33.10). By term, most of the myofibers have peripherally placed nuclei (Figs.  33.11, 33.12, and 33.13), but occasional central nuclei (up to 5%) are considered within the range of normal. Early muscle fibers are considered to be undifferentiated type 2c fibers. Further muscle fiber type differentiation occurs at approximately 18 or 19  weeks gestation (in the range of 16–20 weeks) [1, 6]. Type 1 fibers are the first to appear at this time. Later, type 2 fibers appear [1, 7], with type 2b fibers present at approximately 25  weeks gestation, and type 2a fibers recognizable at approximately 35 weeks gestation [6]. By term, nearly 80% of muscle fibers are identified as either type 1 or type 2 (Figs. 33.14 and 33.15) [5]. Type 2c fibers are normally absent in postnatal infants beyond 1–2 weeks [7].

Fig. 33.7  Fetal skeletal muscle at 17  weeks gestation. High-power view of fetal muscle fibers in cross section, highlighting the polygonal shape and mostly peripherally placed nuclei (H&E, 40×)

Postnatal Muscle Development Newborn muscle fibers have polygonal shapes, but they appear more rounded than the typical angulated polygonal fibers seen in older children and adults (Fig.  33.16) [7]. Usually muscle fibers are tightly packed together, but there is a variable amount of perimysial connective tissue consisting of collagen fibers with blood vessels and nerve tissue. Postnatal growth of muscle fibers occurs by increasing the length of sarcomeres and by adding new sarcomeres. There is also some evidence of a gradual rise in fiber number after birth [8]. Satellite cells are quiescent

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Fig. 33.8  Fetal skeletal muscle at 23  weeks gestation. Low-power view of cross section of skeletal muscle. Note the tight fascicular arrangement of muscle fibers. There is less interstitial connective tissue that was seen at 17 weeks gestation (as in Fig. 33.5) (H&E, 10×)

Fig. 33.9  Fetal skeletal muscle at 23  weeks gestation. High-power view of cross section of skeletal muscle showing the fairly uniform polygonal shape of muscle fibers. By this gestational age, nearly all the fibers have peripherally placed nuclei (H&E, 40×)

Fig. 33.10  Fetal skeletal muscle at 23  weeks gestation. High-power view of longitudinally arranged fibers of skeletal muscle. Longitudinal sections do not show the shape of the muscle fiber as well as cross sections, but myofilaments are seen more easily on longitudinal sections. By this gestational age, nearly all the fibers have peripherally placed nuclei (H&E, 60×)

Fig. 33.11  Skeletal muscle at term. Low-power view of cross section of muscle fibers showing fascicular arrangement of the fibers at term. Note the interstitial connective tissue containing nerves and blood vessels. The muscle fibers are larger than earlier in gestation and have rounded, polygonal shapes that are relatively uniform. The majority of the fibers have peripherally placed nuclei in this cross section (H&E, 10×)

myoblast cells with regenerative capacity, which are present beneath the b­ asement membrane of the myofiber [3, 6]. These cells can be called upon to regenerate myofibers if needed. Compared with postnatal muscle, normal fetal muscle contains very little neutral lipid on routine frozen sections

and very abundant glycogen (Fig. 33.17), which is normally washed out by routine tissue processing and paraffin embedding. Skeletal muscle sampled from premature neonates several weeks after birth can show stainable lipid in a normal pattern (Fig. 33.18).

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Fig. 33.12  Skeletal muscle at term. Medium-power view of cross section of muscle fascicles. The muscle fibers have a fairly uniform, rounded polygonal shape, and most of the fibers have peripherally placed nuclei. Occasional central nuclei can be seen at this gestational age (H&E, 20×)

Fig. 33.13  Skeletal muscle at term. High-power view of muscle fibers in cross section, highlighting the rounded, polygonal shape and mostly peripherally placed nuclei (H&E, 40×)

Fig. 33.14  Skeletal muscle fiber types in full-term neonate. This frozen section muscle biopsy examined after ATPase reaction at pH 9.4 demonstrates type 1 fibers (light brown) and type 2a and 2b fibers (dark brown). Note the presence of both type 1 and type 2 fibers by this method (frozen section, ATPase pH 9.4, 60×)

Fig. 33.15  Skeletal muscle fiber types in full-term neonate. This frozen section muscle biopsy examined after ATPase reaction at pH 4.6 demonstrates type 1 fibers (dark brown), type 2b fibers (light brown), and type 2a fibers (unstained). Note the presence of both type 2a and type 2b fibers at this gestational age (frozen section, ATPase pH 4.7, 60×)

Special Considerations

are each surrounded by a connective-tissue sheath called the perimysium. The perimysium contains the nerves and blood vessels to and from the muscle unit. Each individual fiber is invested, either fully or partially, by the endomysium, which is a connective-tissue layer composed of collagen, elastic fibers, and reticulin fibers [5]. The terminal blood supply to the muscle fiber is found within the endomysium (Fig. 33.19).

Supporting Framework of Muscle The epimysium is the collagenous tissue that surrounds groups of fascicles and forms the outer fascia of the entire muscle. At the muscle ends, the epimysium merges with the dense connective tissue of the tendon. The muscle fascicles

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Fig. 33.16  Newborn skeletal muscle. Note the rounded, polygonal fibers with peripheral nuclei (frozen section, H&E, 60×)

a

Fig. 33.18  Lipid in skeletal muscle. (a) Frozen section of skeletal muscle from a 4-week-old neonate with a corrected age of 30 weeks gestational age. Occasional large fibers, which are probably type 1 fibers, contain microvesicular lipid (arrowhead) in an intermyofibril-

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Fig. 33.17  Newborn skeletal muscle with abundant glycogen. This is a frozen section of muscle stained with periodic acid–Schiff (PAS) stain. Note the abundant glycogen in both a subsarcolemmal and sarcolemmal pattern (frozen section, PAS, 10×)

b

lary pattern. (b) Frozen section of skeletal muscle from a 30-week fetus showing very little neutral lipid with Oil Red O staining (frozen section, Oil Red O stain, 40×)

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Muscle Spindles

muscle spindle begin to differentiate from the extrafusal fibers in week 12 of gestation, and the formation of the capMuscle spindles are mechanoreceptors that sense length and sule also begins at about that time [9]. By the 15th week of tension in the muscle. Histologically, the muscle spindle gestation, it appears that most, if not all, muscle spindles appears as an encapsulated structure within the perimysium. have formed [9]. Over gestation, the size of the muscle spinThe capsule is composed of multiple thin layers of collagen-­ dles increases and the capsule thickens, as seen in containing flattened, specialized fibroblasts. Contained Figs. 33.20, 33.21, and 33.22. At 12–14 weeks, the capsule within the capsule are several intrafusal fibers, known as bag is only a single layer, and by 15 weeks it begins to have two fibers and chain fibers. The bag fibers are generally larger layers (see Fig.  33.20). The layers increase over gestation and longer than the chain fibers [5]. with up to a maximum of five noted at 31 weeks gestation The first histological evidence of the muscle spindle (see Fig. 33.21) [9]. The capsule of the mature, adult muscle appears at 11 weeks gestation as a network of nerve fibrils spindle generally has 10–15 layers [5]. The intrafusal fibers around a few young muscle cells. The intrafusal fibers of the enlarge over gestation, and the number of intrafusal fibers

Fig. 33.19  Perimysium and endomysium. This image of term muscle demonstrates the edge of a muscle fascicle with its surrounding perimysium (P). Note the wispy connective tissue, the endomysium, surrounding the individual muscle fibers and containing capillaries (arrow) (H&E, 60×)

Fig. 33.20  Muscle spindle at 16 weeks gestation. In the center of the image is an encapsulated muscle spindle (arrow) housing intrafusal fibers of varying sizes with central nuclei. At this gestational age, the capsule is only a few layers thick, and the muscle spindle is small in this high-power image (H&E, 60×)

Fig. 33.21  Muscle spindle at 34 weeks gestation. In the center of the image is a longitudinally oriented muscle spindle (arrows). The connective-­tissue capsule is slightly thicker, and the muscle spindle is larger than in Fig. 33.20. Intrafusal fibers of varying sizes with central and peripheral nuclei are noted (H&E, 20×)

Fig. 33.22  Muscle spindle in a newborn. In the center of this frozen section, muscle biopsy of a newborn is an encapsulated muscle spindle. Note the larger size of this muscle spindle and several concentric thin layers of collagen forming the capsule, with specialized fibroblasts within the capsule. Within the capsule are several intrafusal fibers cut in cross section (frozen section, H&E, 20×)

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within a muscle spindle increases by splitting or by incorporation of adjacent fibers up until approximately 31  weeks gestation [9]. Adult muscle spindle usually contains 3–15 intrafusal fibers [5].

References 1. Carpenter S, Karpati G.  Pathology of skeletal muscle. 2nd ed. New York: Oxford University Press; 2001. 2. Sadler TW, Langman J.  Langman’s medical embryology. 11th ed. Philadelphia: Wolters Kluwer Lippincott Williams & Wilkins; 2010. 3. Schoenwolf GC, Larsen WJ. Larsen’s human embryology. 4th ed. Philadelphia: Elsevier/Churchill Livingstone; 2009.

L. M. Ernst and P. Shannon 4. O’Rahilly R, Müller F.  The skeletal system and the limbs. In: Human embryology & teratology. 3rd ed. New  York: Wiley-Liss; 2001. p. 357–94. 5. Heffner RR, Balos LL.  Skeletal muscle. In: Mills SE, editor. Histology for pathologists. Philadelphia: Lippincott Williams & Wilkins; 2007. p. 195–8. 6. Romansky SG.  Neuromuscular diseases. In: Gilbert-Barness E, editor. Potter’s pathology of the fetus, infant, and child. 2nd ed. Philadelphia: Mosby Elsevier; 2007. p. 1889–957. 7. Lake BD, Barbet JP.  Skeletal musculature. In: Wigglesworth JS, Singer DB, editors. Textbook of fetal and perinatal pathology. 2nd ed. Malden: Blackwell Science; 1998. p. 1083–109. 8. Adams RD, DeReuck J. Metrics of muscle. In: Basic research in myology. Amsterdam: Excerpta Medica; 1972. 9. Cuajunco F. Development of the neuromuscular spindle in human fetuses. Contrib Embryol. 1940;28:95–128.

Part IX Integument

Skin

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Maria Cristina Isales, Timothy Tan, and Lily Marsden

Introduction

naling blocks the ability of these neuroectodermal cells to respond to fibroblast growth factors (FGFs), instead allowing In both the fetus and adult, the skin is the largest organ of the expression of bone morphogenic proteins (BMPs) and develbody. During fetal life, the skin undergoes a complex expan- opment into the epidermis [3]. Between weeks 4 and 6 of gession to form a multilayered structure that can re-epithelialize tation, the monolayer of neuroectoderm evolves into an after injury, prevent dehydration, and protect against infec- epithelial bilayer composed of an epitrichial layer known as tion. Not surprisingly, fetal and adult skin differ greatly both the periderm, overlying a basal or germinative layer [1]. The in structure and in epidermal and dermal protein component periderm is a transient embryologic structure that is thought to expression. Though not well understood, these unique have two functions: (1) protecting the fetus prior to keratinizaexpression profiles, especially in extracellular matrix mole- tion of the epidermis and (2) carbohydrate uptake from the cules, impart fetal skin with the ability to rapidly re-­ amniotic fluid [1]. Through the rest of the first and second triepithelialize and restore normal skin architecture without mesters, the epidermis undergoes stratification through develscar formation. By late gestation, this scarless wound healing opment of multiple intermediate cell layers beneath the has ceased, and protein expression and the cellular composi- periderm, which will become the spinous, granular, and cornition of the skin layers have evolved into those recognized in fied layers of the epidermis. By the end of the second trimesadults. This chapter highlights the structural evolution and ter, the periderm has been sloughed into the amniotic fluid, histological changes that occur in the skin during fetal life. and the epidermis has been covered by multiple cornified cell layers [1]. These provide the fetus with a water impermeability barrier derived from lipids secreted by cells of the stratum Embryology granulosum, which are then processed into hydrophobic species in the interstitium of the stratum corneum. Development of the skin occurs between week 3 and week 30 The epidermis is composed of four cell types: keratinoof gestation, through the juxtaposition of the prospective epi- cytes, melanocytes, Merkel cells, and Langerhans cells. dermis, which develops from ectoderm, and the prospective Keratinocytes comprise 95% of the epidermis and are the dermis, which develops predominantly from mesoderm [1, 2]. cells that form the stratum layers described above [1]. After gastrulation, the embryo is covered by the neuroecto- Melanocytes are derived from the neural crest [1]. They can derm, a single-cell layer that will differentiate into either the be detected in the epidermis by 7 weeks gestation, with dennervous system or skin epithelium [1, 3]. The deciding factor sities highest as the epidermis begins to stratify and appendlies in the Wnt signaling pathway. When turned on, Wnt sig- ages develop [4]. Melanin production begins around 3–4 months gestation, with transference of melanosomes to keratinocytes at about 5  months [4]. Merkel cells are denM. C. Isales · T. Tan dritic or globular neuroendocrine cells of the skin that funcDepartment of Pathology, Northwestern Memorial Hospital, tion as mechanoreceptors [5]. They are believed to be derived Northwestern University Feinberg School of Medicine, from epidermal stem cells in the stratum germinativum and Chicago, IL, USA are found in both the epidermis and dermis, with highest e-mail: [email protected]; [email protected] concentrations along the epidermal ridge of the developing epidermis and in the outer root sheath of developing hair folL. Marsden (*) Utah State Office of the Medical Examiner, Salt Lake City, UT, USA licles [5, 6]. These cells can be detected as early as 8 weeks e-mail: [email protected]

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_34

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gestation in the palmar epidermis and between 10 and 11 weeks in hairy skin [5]. Dermal Merkel cells are located mostly in the upper dermis, close to the dermoepidermal junction, and can be seen as early as 11–12 weeks gestation [5]. Lastly, Langerhans cells, which are thought to migrate to the epidermis from the bone marrow, can be detected through HLA-DR expression by 6–7  weeks gestation, with gain of OKT-6 (CD1) expression by 8–9 weeks gestation [7]. Their morphology varies from small and truncate early in gestation to larger and more dendritic as gestational age increases [7]. The derivation of the dermis is dependent upon anatomic location. The dorsal trunk dermis develops from the dermomyotome portion of the differentiated somite, the dermis of the limbs is derived from the lateral plate mesoderm, and the dermis of the head and neck is derived from neural crest cells [8]. The mesenchymal cells that populate the dermis give rise to dermal blood vessels and connective tissue. These cells also interact with the overlying epithelium through a complex interplay between Notch, Wnt, and sonic hedgehog (Shh) signaling with promoting FGFs and BMP-inhibitory factors to form hair placodes/follicles [1, 3]. The timing of hair development varies by site, with the earliest hair rudiments identifiable in regions of the eyebrow, upper lip, and chin by 9  weeks gestation but closer to 14 weeks gestation in the skin from the abdomen [1, 9]. The pregerm stage of hair development occurs as a response to promoting FGFs and BMP-inhibitory factors in the subjacent mesenchyme [1, 3]. Corresponding Shh signals from hair placodes then direct adjacent mesenchymal cells and fibroblasts to increase in number and condense to form the hair papilla. In the subsequent germ stage, Wnt signaling from the ectoderm instructs the basal cells to grow downward into the dermis [1, 3, 9]. On the scalp, hair canals are fully formed by 19–21 weeks, and hair is visible [10]. The hairs lengthen until 24–28 weeks gestation, at which point they progress from the anagen or growth phase to the telogen or resting phase [10]. Because overall embryonic maturation occurs from the head to the caudal pole, histology of the skin varies with development, appearing more advanced in skin from the neck than in skin from the abdomen or thigh [9]. By the 34th week of gestation, the skin has achieved an architecture similar to that seen in the adult [2], which consists of three components: (1) the epidermis, divided into stratum germinativum, stratum spinosum, stratum granulosum, and stratum corneum; (2) the dermis, which contains connective tissue, blood vessels, hair follicles, sebaceous glands, and sweat glands; and (3) the hypodermis, composed of fat and connective tissue (Fig. 34.1) [1].

Histology Epidermis The original neuroectoderm layer that covers the embryo during the third week of fetal life is composed of a multipo-

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Fig. 34.1  Skin at 37 weeks gestation. By term, fetal skin has achieved an architecture similar to that seen in adults, with well-defined epidermis, dermis, and hypodermis and well-developed hair follicles and adnexal glands (H&E, 10×)

Fig. 34.2  Skin at 10 weeks gestation. The epidermis consists of two epithelial layers and overlies a cellular dermis composed of fibroblastic-­ appearing mesenchymal cells in a hyaluronic acid-rich extracellular matrix. Collagen fibrils are beginning to combine into bundles (H&E, 20×)

tent monolayer of glycogen-filled cells [1]. These embryonic cells differ from basal cells at later stages of development in that they are more columnar and lack hemidesmosomes [10]. This layer is replaced by two epithelial layers between weeks 4 and 6 of gestation: the stratum germinativum, in contact with the dermis, and the overlying periderm (Fig. 34.2) [1]. The stratum germinativum is composed of cuboidal cells with glycogenated cytoplasm, whereas the periderm is composed of flattened squamous cells (Fig. 34.3). A third layer of cells, known as the stratum intermedium, develops from the stratum germinativum under the periderm at 8–11  weeks gestation, marking the beginning of epidermal stratification (Fig. 34.4) [1]. At this stage, the basal cell layer has a higher nuclear to cytoplasmic ratio, begins to express hemidesmo-

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Fig. 34.3  Skin at 10 weeks gestation. A high-power view of the epidermis shows the transient periderm layer of squamous cells overlying the basal layer of cuboidal cells with glycogenated cytoplasm (H&E, 60×)

Fig. 34.4  Skin at 13 weeks gestation. In the epidermis, multiple intermediate cell layers have developed between the periderm and basal cell layers (H&E, 60×)

Fig. 34.5  Skin at 30 weeks gestation. By the end of the second trimester, the periderm has been sloughed, and discrete layers of the epidermis can be distinguished: stratum germinativum, stratum spinosum, stratum granulosum, and stratum corneum. The dermis has a rich network of vessels parallel to the epidermis, with clustering of perivascular cells and increased collagen (H&E, 40×)

Fig. 34.6  Skin at 30  weeks gestation. The stratum germinativum is composed of a single layer of cuboidal cells with high nuclear to cytoplasmic ratios. Above that, the stratum spinosum consists of several layers of keratinocytes with the classic “spiny” appearance created by the desmosomal bridges that connect these cells to each other (H&E, 40×)

somal proteins, and secretes collagen types V and VII, which form the majority of the dermal anchoring fibrils [1]. Between 12 and 26 weeks gestation, the number of intermediate cell layers increases, with cells maturing toward the periderm. By the end of the second trimester, upon cornification of the superficial layers of the epidermis, the periderm is sloughed into the amniotic fluid, where it combines with sebum from sebaceous glands and shed hair (lanugo) to form the vernix caseosa. At this time, the intermediate cells have developed into histologically distinct spinosum, granulosum, and corneum layers through progressive maturation from the stratum germinativum toward the surface (Fig. 34.5) [1]. The

stratum spinosum consists of enlarged, epibasal keratinocytes, held together by desmosomes, and named for the “spiny” histologic appearance that occurs from shrinkage of the microfilaments between those desmosomes during slide preparation (Fig.  34.6) [1]. The stratum granulosum above the stratum spinosum is named for the keratohyaline granules that begin to appear around 21  weeks gestation (Fig.  34.7) [1]. The stratum corneum, the most superficial layer, is composed of multiple rows of tightly cross-linked keratinocytes (now referred to as corneocytes) that have lost their nuclei and cytoplasmic organelles and have become flattened (Fig. 34.8) [1].

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Fig. 34.7  Skin at 23 weeks gestation. High-power view shows the fine, keratohyaline granules that define the stratum granulosum (H&E, 60×)

Fig. 34.8  Skin at 37 weeks gestation. The stratum corneum consists of multiple rows of tightly cross-linked, flat keratinocytes that have lost their nuclei and cytoplasmic organelles (H&E, 60×)

Fig. 34.9  Skin at 37  weeks gestation. Special staining can readily identify melanocytes along the basal layer (Mart-1, 40×)

Fig. 34.10  Skin at 37 weeks gestation. Melanosomes are transferred from melanocytes to keratinocytes by the end of the second trimester (H&E, 60×)

Throughout life, keratinocytes predominate as the primary cells in the epidermis, but melanocytes are readily identifiable. Melanocytes can be detected in the epidermis in the embryologic period, but density is highest in the first trimester, when stratification begins and appendages develop. Melanocytes begin producing melanin at about 3–4 months gestation, and melanosomes transfer to keratinocytes at about 5 months (Figs. 34.9 and 34.10).

between the dermis and subcutaneous tissue [11, 12]. Between 9 and 12 weeks, though the thickness of the dermis stays the same, the cells appear more fibroblastic, and the collagen content increases, with fibrils associating into bundles (see Fig. 34.2) [11]. After 13 weeks gestation, the dermis becomes distinctly fibrous, and by 14  weeks, the papillary and reticular dermal regions are distinguishable from each other, with the papillary dermis directly beneath the epidermis containing a higher density of fibroblasts and finer collagen fibers (Fig. 34.11) [11, 12]. Between 14 and 21 weeks gestation, fibroblasts are the most abundant cells in the dermis, but perineural cells, pericytes, melanoblasts, Merkel cells, and mast cells can be individually identified [1]. Elastic fibers composed of elastin on a microfibrillar network are identified in the dermis by 6 months gestation, though they do not become mature in structure until postnatal life (Fig. 34.12) [11].

Dermis In the embryologic period (5–8 weeks gestation), the dermis is very cellular, composed of abundantly glycogenated mesenchymal cells in a hyaluronic acid-rich extracellular matrix [1]. By 9  weeks gestation, the dermis contains an increased amount of collagen, and there is delineation

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Fig. 34.11  Skin at 14  weeks gestation. The dermis has become distinctly fibrous. The papillary and reticular dermal regions can be distinguished from each other by the higher density of cells in the papillary dermis beneath the epidermis. Simple, tube-like dermal vessels can be seen running parallel to the epidermis (H&E, 20×)

Dermal vasculature development begins in the embryonic stage with joining of endothelial cells in the presumptive dermis to form simple, tube-like vessels in a single plane ­parallel to the epidermis and dermal-subcutaneous interface [12]. Between 7 and 8  weeks gestation, two closely positioned planes of vessels are identifiable, and there is heterogeneity in vessel diameter [12]. By 20 weeks, the small vessels closest to the epidermis show increasing wall thickness, more perivascular cells, and increased collagen (see Fig.  34.5) [12]. The presence of nerves in the dermis appears to precede vessels, with migration occurring in closer proximity to the epidermis [12]. The mature dermis is a supporting layer of collagen and elastic fibers embedded in a ground substance containing hydrated and sulfated glycosaminoglycans and associated with a rich blood supply and a cellular component composed of fibroblasts, mast cells, and histiocytes [1, 11]. The termination of dermal maturation is signaled by increased tensile strength and the evolution from a non-scarring to a scarring response to wound healing [10].

Adnexa Hair Follicles Hair follicle development occurs in multiple stages beginning with the pregerm stage and progressing through the germ, hair peg, and bulbous hair peg stages. The timing and trajectory of these changes can vary with body site. In the pregerm stage of hair development, there is crowding of nuclei in the stratum germinativum, with early invagination into the underlying dermis [1, 3]. In the subsequent germ stage, the basal cells elongate and grow downward to form the hair placode [1, 3, 9]. At this time, adjacent mesenchymal cells and

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Fig. 34.12  Skin at 37 weeks gestation. By the third trimester, the dermis contains fibers of elastin in a microfibrillar network (elastic Van Gieson, 60×)

Fig. 34.13  Skin at 15 weeks gestation. Early hair follicle formation is identified by crowding and elongation of basal nuclei with invagination into the underlying dermis, forming the hair placode. The subjacent dermal mesenchymal cells begin to increase in number and condense, forming the rudiments of the hair papilla (H&E, 40×)

­ broblasts increase in number and condense to form the rudifi ments of the hair papilla and hair peg [1, 3] (Fig. 34.13). The hair peg rapidly grows downward obliquely, with the deepest portion becoming bulbous and sitting on top of the condensing mesenchymal cells, which have formed the dermal papilla [1]. At this stage, the hair peg is still solid, with the outer cells arranged radially to the long axis. The advancing end is ­composed of self-renewing progenitor cells, which are distinguishable by being taller and narrower than those around the periphery of the developing follicle [10] (Fig.  34.14). These progenitor cells differentiate as they move toward the skin surface, creating the inner root sheath that lines the canal for the protruding hair [10]. The next stage, the bulbous hair peg stage, is defined by the appearance of two epithelial

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bulges on the outer wall of the developing follicle: the upper bulge represents the rudimentary sebaceous gland, and the lower bulge will become the site of arrector pili muscle attachment [1]. By this time, the dermal papilla has been enveloped by the advancing end, and hair can be seen in the canal (Fig. 34.15).

Sebaceous Glands The rudimentary sebaceous gland present in the bulbous hair peg stage is composed of sebaceous gland progenitor cells

Fig. 34.14  Skin at 17  weeks gestation. As hair follicle development advances, the hair peg grows downward into the dermis, with the outer cells arranged radially to the long axis. Note that the “hair canal” is still solid. The cells at the advancing end are taller and narrower than those around the periphery of the developing follicle. At the same time, the condensing mesenchymal cells have formed the dermal papilla (H&E, 20×)

Fig. 34.15  Skin at 22 weeks gestation. In the later stages of hair development, the follicle has a partially differentiated sebaceous gland above a “bulge” that will become the attachment site for the developing arrector pili muscle. The dermal papilla has become enclosed at the advancing end, and a hair can be seen in the canal (H&E, 20×)

M. C. Isales et al.

that initially contain a moderate amount of glycogen. Soon after development, the central cells become large and foamy, as the glycogen is replaced by lipid droplets (sebocytes) [1]. Sebaceous glands are differentiated by 13–15 weeks gestation and function through a continual flux of proliferating, differentiating, and disintegrating sebocytes that release oils and sebum into the hair canal [1, 3]. The sebum produced becomes a large component of the vernix caseosa [1]. Before birth, fetal sebaceous glands are large and well differentiated, but after birth they rapidly decrease in size and become functional again only during puberty [1].

Eccrine Glands Eccrine gland rudiments are recognized by undulations of the stratum germinativum at regularly spaced intervals [1]. These rudiments are first identified on the palmoplantar surfaces by 12 weeks gestation and are seen over the rest of the body by 20 weeks [1, 9]. In contrast to the hair follicle buds, the cells forming the eccrine buds are more eosinophilic and cylindrical and are not accompanied by underlying mesenchymal proliferation [9] (Fig. 34.16). By 14–15 weeks gestation on the palms and soles and 23 weeks over the body, the eccrine glands have penetrated deep into the dermis, to the level of transition from the dermis to fatty tissue [1, 9]. By 25 weeks gestation, the eccrine glands over the body have begun to form terminal coils, with complexity significantly increasing by 29–30 weeks, so that histologic examination shows multiple transverse sections deep in the dermis [9] (Fig. 34.17). Apocrine Glands Unlike eccrine glands, which arise independently, apocrine sweat glands bud from the hair follicle during the 20th week

Fig. 34.16  Skin at 27 weeks gestation. In contrast to hair follicle buds, eccrine duct buds are composed of eosinophilic cells with no underlying mesenchymal condensation (H&E, 40×)

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Fig. 34.17  Skin at 37 weeks gestation. Eccrine glands have become complex coiling structures that penetrate deep into the dermis. The terminal coil is so complex that it is seen histologically as multiple transverse cross sections (H&E, 10×)

of gestation. The clear cells and mucin-secreting cells of the apocrine gland are distinguishable by 28 weeks of gestation. The apocrine gland is initially functional in the third trimester, while the eccrine glands are nonfunctional. Postnatally, the apocrine glands become quiescent, and the eccrine glands reach functional maturity [10].

Nails Before hair follicle formation, at 8–10 weeks, the dorsal digit tips develop a flat surface, which represents the future nail bed. The proximal nail fold develops from a portion of ectoderm that spreads inward at an oblique angle. The nail matrix cells are located on the ventral aspect of this ectodermal invagination. From 16 to 24 weeks, the nail plate replaces the embryonic cornified layer and grows distally from the proximal fold. Although the keratinization of the nail and epidermis is similar, the keratin expression profile of the nail is distinct, providing greater stability and rigidity [10].

Fig. 34.18  Skin at 23 weeks gestation. The hypodermis/subcutaneous tissue is delineated from the overlying dermis but has not yet organized into lobules (H&E, 4×)

Hypodermis Subcutaneous tissue, also known as the hypodermis, is formed by adipocytes of mesenchymal origin. It consists of adipocytes surrounded by connective tissue with vascular elements and nerve branches. By 7–8  weeks gestation, there is a distinct separation between the hypodermis and the overlying dermis, which is separated by a level of thin-walled vessels (Fig. 34.18). In the 12th week, preadipocytes derived from the mesenchyme begin to differentiate and accumulate lipids (Fig. 34.19). After 24 weeks, the adipocytes become aggregated into lobules that are divided by fibrous septa [10] (Fig. 34.20).

Fig. 34.19  Skin at 23  weeks gestation. Higher power view of the hypodermis shows preadipocytes that are beginning to differentiate and accumulate lipid (H&E, 60×)

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Fig. 34.20  Skin at 31 weeks gestation. After 24 weeks gestation, adipocytes in the hypodermis have accumulated into lobules separated by fibrous septa (H&E, 4×)

References 1. McGrath JA, Eady RAJ, Pope FM. Anatomy and organization of human skin. In: Burns T, Breathnach S, Cox N, Griffiths C, editors. Rook’s textbook of dermatology. 7th ed. Malden: Blackwell Publishing; 2004. 2. Helmo FR, Machado JR, Oliveira LF, Rocha LP, Cavellani CL, Teixeira Vde P, et al. Morphological and inflammatory changes in

M. C. Isales et al. the skin of autopsied fetuses according to the type of stress. Pathol Res Pract. 2015;211:858–64. 3. Fuchs E.  Scratching the surface of skin development. Nature. 2007;445:834–42. 4. Koster MI, Loomis CA, Koss T, Chu D. Skin development and maintenance. In: Bolognia JL, Jorizzo JL, Schaffer JV, editors. Dermatology. 3rd ed. Philadelphia: Elsevier Saunders; 2018. p. 56–65. 5. Kim DK, Holbrook KA. The appearance, density, and distribution of Merkel cells in human embryonic and fetal skin: their relation to sweat gland and hair follicle development. J Invest Dermatol. 1995;104:411–6. 6. Van Keymeulen A, Mascre G, Youseff KK, Harel I, Michaux C, De Geest N, et al. Epidermal progenitors give rise to Merkel cells during embryonic development and adult homeostasis. J Cell Biol. 2009;187:91–100. 7. Foster CA, Holbrook KA, Farr AG. Ontogeny of Langerhans cells in human embryonic and fetal skin: expression of HLA-DR and OKT-6 determinants. J Invest Dermatol. 1986;86:240–3. 8. Schoenwolf GC, Bleyl SB, Brauer PR, Francis-West PH. Development of the skin and its derivatives. In: Larsen’s human embryology. 5th ed. Philadelphia: Elsevier/Churchill Livingstone; 2015. p. 155–71. 9. Ersch J, Stallmach T.  Assessing gestational age from histology of fetal skin: an autopsy study of 379 fetuses. Obstet Gynecol. 1999;94:753–7. 10. Hoath SB, Mauro T.  Fetal skin development. In: Eichenfeld LF, Frieden IJ, editors. Neonatal and infant dermatology. London: Elsevier Saunders; 2015. p. 1–13. 11. Smith LT, Holbrook KA. Embryogenesis of the dermis in human skin. Pediatr Dermatol. 1986;3:271–80. 12. Johnson CL, Holbrook KA. Development of human embryonic and fetal dermal vasculature. J Invest Dermatol. 1989;93:10S–7S.

35

Mammary Gland Dale S. Huff

Introduction The prenatal and neonatal development of the mammary gland occurs in three stages: (1) the primordia, from 6 to 8  weeks postmenstrual age; (2) the nipple, from 8 to 19 weeks; and (3) the ducts of the mammary gland proper, from 19 weeks to the neonatal period. The developing breast is composed of specialized epithelium and mesenchyme. The specialized epithelium will form the ectodermal nipple and the mammary ducts. The specialized mesenchyme will form the stroma and smooth muscle of the nipple and areola, the intralobular stroma, and the interlobular septa, as well as the supporting connective tissues, including the capsule, Cooper ligaments, and pectoral ligaments. Cooper ligaments anchor the breast to the overlying dermis. Pectoral ligaments anchor the breast to the underlying pectoralis fascia. Interactions between the mesenchyme and epithelium result in the synchronous differentiation of the epithelial basal cells into myoepithelium and of the mesenchyme into its various components. The early development of the specialized mesenchyme may be androgen-dependent. The prenatal development of the nipple and mammary ducts is independent of hormonal control until maternal and placental hormones stimulate the hyperplasia and secretory activity of the ducts, beginning at 35 weeks. Fetal prolactin stimulates the neonatal secretion of milk, which can lead to a transient discharge of a thin, milky liquid sometimes referred to as “witches’ milk” from the nipple at 5–7 days of age. The prenatal and neonatal mammary gland is similar in females and males. The postmenstrual gestational ages given for the timing of the developmental events are estimates. The ages at which

D. S. Huff (*) Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]

the events occur vary widely among published series, and studies large enough to define the range of gestational ages at which specific features normally appear are not available. Therefore, the definitions of normal, delayed, or advanced development for a given gestational age have not been established.

Embryology The Primordia: 6 2/7 to 8 0/7 Weeks The primordia of the breast are the mammary band, mammary crest, and primordial nipple. The mammary band is also known as the intermembral segment of the ectodermal ring [1], which is described in the Part IV Introduction, Early Embryology of the Genitourinary Tract. The mammary band is the segment of the ectodermal ring between the upper and lower limb buds. It appears early in the sixth week. The mammary band is present in nonmammalian as well as mammalian vertebrates, so it is not specific for the mammary gland. The mammary crest, which develops within the mammary band, is the primordium that is specific for the mammary gland [1, 2]. The mammary crest appears at 7 weeks as a narrow, microscopic thickening of the ectoderm within the mammary band, adjacent to the caudal edge of the upper limb bud. It grows caudally as a narrow, linear crest for a variable distance toward the lower limb bud. The mammary crest stops its caudal growth and then shortens by regression beginning at its caudal end and extending cranially. By 8 weeks, all that remains of the crest is a round, lens-shaped placode of thickened ectoderm near the upper limb bud, in the region that will become the axilla. The mesoderm under the ectodermal placode becomes specialized and differentiates into two layers, a dense band under the ectodermal basement membrane and a deeper, looser, well-vascularized layer. This ectodermal placode and the underlying specialized mesoderm are the primordium of the nipple [1, 2].

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_35

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Histology The Nipple: 8–19 Weeks During the 8th week, the nipple is a microscopic, lens-­shaped, nodular placode in the lateral ectoderm at the caudal edge of the upper limb bud. It bulges upward above the surface of the surrounding epidermis and downward into the underlying specialized mesoderm. The low, cuboidal basal cells of the surrounding epidermis differentiate into the specialized basal cells of the nipple, which are tall, columnar cells with apical nuclei and subnuclear vacuoles. The other cells of the nipple, the central cells, are large, polygonal, and vacuolated; they have small, eccentric nuclei. The two-­ layered, specialized mesenchyme differentiates further into four concentric layers stretched around the deep surface of the nipple. The most superficial layer will become the dense, dermis-like stroma of the nipple and areola. The second layer will become the layer of smooth muscle of the nipple and areola. The third layer will become the intralobular stroma. The deepest layer will form the dense, supporting connective tissues of the breast, including the interlobular septa, the capsule of the breast, the Cooper ligaments, and the pectoralis ligaments [2]. In routine autopsy material, some of these layers may be indistinct. During the ninth week, the position of the nipple begins to shift from the lateral to the ventral aspect of the body. The surface of the nipple flattens and becomes flush with the surrounding epidermal surface. The deep portion of the nipple enlarges and pushes into the underlying mesenchyme, forming an enlarging hemispheric ectodermal nodule. The specialized mesoderm completely surrounds the globular nipple and extends laterally under the immediately adjacent epidermis that will become the areola [2]. During the tenth week, the specialized mesenchyme becomes more distinct and proliferates, pushing the nipple up above the level of the adjacent epidermis. A shallow pit, the mammary pit, appears in the center of the ectodermal surface of the nipple, forming the inverted nipple characteristic of the prenatal breast. By the 14th week, the immunohistochemical phenotypes of the basal and central cells of the nipple are distinct from each other and from their counterparts in the adjacent epidermis [3]. The nipple is in its final position on the ventral body wall. Beginning at the 15th week, the deep border of the nipple becomes irregular and lobulated (Figs. 35.1 and 35.2). The specialized mesenchyme continues to proliferate, forming a mound at the top of which sits the inverted nipple. The mammary pit deepens. The tall, columnar basal cells of the nipple remain distinct from the small, cuboidal basal cells of the adjacent epidermis. The areola becomes identifiable grossly and microscopically because it lacks the pilosebaceous units and sweat glands that appear in the adjacent skin at this time. Lobules of fat appear in the subcutaneous tissue of the adjacent skin but not in the specialized mesoderm of the nipple

Fig. 35.1  Lobular phase of the development of the nipple: gestational age 16 weeks. The diameter of the ectodermal nipple is 0.25 mm. The narrow arrows close to the edges point to pilosebaceous units at the junction of the areola and adjacent epidermis. The thick arrows point to the edges of the ectodermal nipple. The arrowhead points to the shallow mammary pit. The ectodermal nipple sits on top of the mound of specialized mesenchyme; this picture persists throughout prenatal development. Note the irregular indentations of the deep border of the nipple, the absence of pilosebaceous units in the areola, the presence of pilosebaceous units in the adjacent skin, the concentric layers of specialized mesenchyme around the nipple and under the areola, and the underlying pectoralis muscle (hematoxylin and eosin (H&E), 4×)

Fig. 35.2  Lobular phase of the development of the nipple: gestational age 16  weeks. This is higher magnification of Fig.  35.1. The arrows point to the edges of the nipple. The arrowhead points to the mammary pit. Keratinization of the mammary pit is beginning. The small, inconspicuous basal cells of the adjacent epidermis differentiate into the specialized tall, columnar basal cells of the nipple. Note the large, clear central cells. The concentric layers of the specialized mesenchyme around the nipple and under the areola are easily seen (H&E, 10×)

and areola. At 16  weeks, keratinization begins, and the ­mammary pit becomes filled with keratin. By 17 weeks, the basal cells of the nipple are immunophenotypically incompletely differentiated epithelial precursor cells [3, 4]. Sensory end organs appear in the specialized mesenchyme.

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 he Mammary Ducts: 19 Weeks T to the Neonatal Period Until the 19th week, the development of the breast has involved only the nipple, areola, and specialized mesoderm. The development of the mammary ducts begins at 19 weeks, when the basal cells of the nipple at a few discrete foci differentiate into pseudostratified placodes of thin, columnar cells with scant cytoplasm and long, hyperchromatic nuclei. These placodes are the beginnings of the primary mammary ducts (Figs. 35.3 and 35.4). A thick layer of smooth muscle appears in the second layer of the specialized mesoderm under the nipple and areola (Fig. 35.5). The orientation of the muscle is predominately circular, but vertically oriented fibers are also present, most noticeably parallel to the primary ducts under the nipple. Glands of Montgomery, sometimes called tubercles of Montgomery, appear in the areola. Glands of Montgomery are composed of a large complex of sebaceous units associated with a single duct that is similar to a lactiferous duct, which opens onto the surface of the areola, often near its periphery. Although the ducts are difficult to distinguish from lactiferous a

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ducts, the glands of Montgomery end blindly and do not contribute to the ductal–lobular parenchyma of the mammary gland. At 19 weeks, the immunophenotype of the basal cells has differentiated into myoepithelial cell precursors and that of the central cells into luminal epithelial cell precursors [3, 4]. As the first mammary duct placodes elongate into solid primary mammary ducts, new placodes form and then elongate until a total 15–20 solid, straight primary ducts are present. These are the future lactiferous ducts of the major segments of the adult breast; they penetrate perpendicularly into the first two layers of dense, specialized mesenchyme. Each has a long, straight stem or stalk and an end bud. The end buds are bulbous and solid. The stalks develop lumens and are lined by two layers of cells—basal cells and luminal cells. The basal cells are cuboidal (in contrast to the tall, columnar basal cells of the nipple), and they lie upon a thick, continuous basement membrane. The luminal cells are small and nonvacuolated (in contrast to the large, vacuolated central cells of the nipple). The solid end buds lack a distinct basement membrane and are composed of undifferentiated basal and central epithelium (Figs. 35.6, 35.7, and 35.8). By 22 weeks, basal cells at the junction of the stalks and end buds differentiate into smooth muscle actin-positive, t­erminally differentiated myoepithelial cells, and the central cells differentiate into terminally differentiated luminal epithelial cells [3–5]. By the 24th week, the end buds of the elongating ducts penetrate through the dense layer of smooth muscle and into the underlying layer of loose mesenchyme. A sheath of the loose mesenchyme surrounds each duct and will become the intralobular stroma. The ducts with their loose stromal

b

Fig. 35.3  Earliest appearance of the buds of the mammary ducts: gestational age 21 3/7 weeks. The diameter of the ectodermal nipple has not enlarged and is 0.25 mm. (a) Buds of mammary ducts. This breast is very similar to that shown in Figs. 35.1 and 35.2, except for the presence of the buds of the mammary ducts. The arrows point to pilosebaceous units at the junction of the areola and adjacent epidermis. The differences between the adjacent skin and the areola are depicted, with numerous pilosebaceous units in the adjacent skin but none in the areola. Fetal fat is now abundant in the subcutaneous tissue under the adjacent skin but is sparse under the periphery of the areola and absent under the nipple and central areola. The dense, specialized mesenchyme extends to the pectoralis fascia. Sensory end organs have appeared in the specialized mesenchyme of the nipple and areola, and dilated capillaries surround the nipple (H&E, 2×). (b) At a higher magnification of the same specimen, the mammary pit is tangentially sectioned and appears as tubular cross sections. Two early buds of mammary ducts are seen, one arising from each shoulder of the nipple. The horizontal arrow points to the bud on the left. The arrowheads point to sensory end organs (H&E, 4×)

Fig. 35.4  Earliest appearance of the buds of the mammary ducts: gestational age 21 3/7 weeks. Higher magnification of Fig. 35.3 demonstrates details of the differentiation of the basal cells from the low, inconspicuous cells of the adjacent epidermis to the tall, specialized basal cells of the nipple and then to the pseudostratified, columnar, hyperchromatic cells of the bud of the mammary duct on the left. The large, vacuolated central cells of the nipple blend into the midzone of the epidermis (H&E, 20×)

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sheaths are separated by septa, the interlobular septa, arising from the fourth dense, deep layer of specialized mesenchyme. The end buds begin to grow by branching morphogenesis within the loose intralobular stroma [6]. The end buds are the active sites of proliferation, invasion, branching, and differentiation of the ducts. Sebaceous glands appear at the origins of the ducts and glands of Montgomery (Figs.  35.9 and 35.10). Lumens extend into the end buds, the mammary pit becomes wider and

b

Fig. 35.5  Smooth muscle in the second layer of specialized mesenchyme: gestational age 23 0/7 weeks. The diameter of the ectodermal nipple is still 0.25 mm. (a) Concentric rings of smooth muscle surround the nipple and extend like wings into the mesenchyme of the central areola. The arrows point to pilosebaceous units at the junction of the areola and adjacent epidermis. One bud of a mammary duct arises from the top right side of the nipple. Fetal fat is not seen under the nipple and areola (smooth muscle actin immunostain, 2×). (b) Higher magnification of (a). Note the first layer of dense mesenchyme under the nipple and areola, the smooth muscle in the second layer, and the undifferentiated deeper layers of mesenchyme extending to the pectoralis fascia (smooth muscle actin immunostain, 4×)

Fig. 35.6  Ducts penetrating into the first two layers of specialized mesenchyme: gestational age 21 5/7  weeks. The diameter of the nipple is 0.25 mm. This photograph includes the ectodermal nipple, the specialized mesenchyme, and the central areola, but not the periphery of the areola. A corona of dilated capillaries surrounds the nipple, and the mammary pit is deepening. The histologic phenotype of the basal cells of the nipple remains distinct from that of the adjacent epidermis. An elongating duct arising from the lower right corner of the nipple is pushing into the mesenchyme (H&E, 10×). The next two figures are from the same breast

Fig. 35.7  Ducts penetrating into the first two layers of specialized mesenchyme: gestational age 21 5/7 weeks. Higher magnification demonstrates that the solid duct is composed of basal and luminal epithelial cells. The cuboidal basal cells are precursors of myoepithelial cells and are distinct from the columnar basal cells of the nipple. The luminal cells are squamous-like (H&E, 20×)

Fig. 35.8  Ducts penetrating into the first two layers of specialized mesenchyme: gestational age 21 5/7 weeks. In this deeper section, two penetrating ducts join before entering the nipple. Each has a bulbous, solid end bud that is the site of ductal differentiation, proliferation, and elongation. The concentric layers of mesenchyme extend to the fascia of the pectoralis muscle at the bottom of the picture (H&E, 10×)

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Fig. 35.9  Early branching morphogenesis of mammary ducts: gestational age 24 4/7 weeks. The enlarging nipple has a diameter of 1 mm. The inverted nipple sits atop a mound of specialized mesenchyme, and most of the areola is included. The ducts have pushed through the layer of smooth muscle, penetrated into the underlying layer of loose mesenchyme that will become the intralobular stroma, and are elongating by branching morphogenesis. Glands of Montgomery arise from the areola lateral to the nipple. They are nearly identical to the major mammary ducts but do not have end buds and do not branch. One is associated with sebaceous glands (H&E, 4×). The next two figures are from the same breast

s­hallower, and the inverted nipple becomes cup-shaped (Figs. 35.11, 35.12, and 35.13). By 26  weeks, continued branching morphogenesis results in a complex ductal tree, a lobular pattern, and an enlarging breast bud. Secretions appear in the lumens of the end buds. Maturing interlobular septa accentuate the lobular-like architecture, and Cooper ligaments and pectoralis ligaments become more conspicuous. Adipose tissue appears within the intralobular stroma and interlobular septa, and the inverted nipple becomes larger and deeper. By 32–35  weeks, the breast bud becomes palpable but is usually only a few millimeters in diameter (Figs.  35.14, 35.15, 35.16, and 35.17). By 35 weeks gestation, ductal elongation, end bud branching, and secretion accelerate, causing enlargement of the breast bud (Figs. 35.18, 35.19, and 35.20). Extramedullary hematopoiesis is commonly seen in the intralobular stroma; it is seen less often in the interlobular septa. The amount varies widely and tends to be more extensive near term than earlier in development. In some cases, the myoepithelial cells assume the phenotype of mature smooth muscle cells (Fig. 35.21). Squamous metaplasia of large ducts, apocrine metaplasia (Fig.  35.22), and giant cells in the interlobular stroma may be seen, as have been described in adults (Fig. 35.23). At term, the breast bud is usually 8 mm in diameter. Secretion of “witches’ milk” and oozing of milk from the nipple may be apparent by 5–7 days of life. The lobular architecture, branching end buds, and secretory changes mimic but are probably not identical to the terminal duct

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Fig. 35.10  Early branching morphogenesis of mammary ducts: gestational age 24 4/7 weeks (higher magnifications of Fig. 35.9). (a) The enlarging crater of the inverted nipple is filled with keratin. The major ducts are lined by a layer of cuboidal basal cells and a layer of flat, luminal cells with eosinophilic cytoplasm (H&E, 10×). (b) The terminal ducts enter the intralobular stroma and branch into solid terminal end buds. The terminal ducts are lined by a basal layer of terminally differentiated myoepithelial cells and a luminal layer of terminally differentiated luminal epithelial cells. The solid end buds are lined by basal precursors of myoepithelial cells and central precursors of luminal epithelial cells. Epithelial differentiation occurs at the junction of the terminal duct and end bud. Note the difference in the phenotypes of the epithelial cells of the primary ducts, terminal ducts, and end buds (H&E, 20×)

lobular unit and alveolae of the adult. Some ducts become cystic (Fig.  35.24). The breast continues to enlarge for 2 weeks or longer after a term birth and may attain a ­diameter of a few centimeters or more. The nipple may remain inverted. Beginning at approximately 2  weeks postnatal age, the distal branches of the ductal tree begin to regress, and secretion decreases. The intralobular stroma begins to disappear. Cystic change may become extensive prior to complete regression (Figs.  35.25 and 35.26). The dense, interlobular stroma becomes more prominent, and the breast is reduced to

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Fig. 35.11  Progressive branching morphogenesis. The breast shown is only slightly more advanced than the one shown in Fig. 35.9. The gestational age is 26 5/7 weeks, and the diameter of the nipple is 1 mm. After this nipple becomes everted sometime after birth, the orifice of the duct entering the middle of the nipple in this image will be located at the tip of the everted nipple, whereas those entering the lateral border here will be located at sides of the everted nipple. The layer of smooth muscle is prominent (H&E, 4×). The next two figures are from the same breast

Fig. 35.13  Progressive branching morphogenesis: higher magnification of the lobule on the right in Fig. 35.12. The lumen extends into the terminal branches of the duct. The myoepithelial and luminal epithelial cells demonstrate progressive differentiation (H&E, 20×)

D. S. Huff

Fig. 35.12  Progressive branching morphogenesis: higher magnification of Fig. 35.11. The branches of each primary duct form a simple lobule invested in loose intralobular stroma. The lobules are separated by slightly more dense interlobular connective tissue, which is derived from the fourth layer of the specialized mesenchyme. Undifferentiated end buds are seen on the left, and sensory end organs are prominent (H&E, 10×)

Fig. 35.14  Complex branching of mammary ducts: gestational age 34 4/7  weeks. The diameter of the breast bud is 3  mm. The complex branching results in an enlarging, palpable breast bud, and the primary ducts entering the nipple are lined by squamous epithelium. Each primary duct has several generations of branches that form complex lobules surrounded by intralobular stroma. The interlobular septa are becoming more dense, and the irregular capsule, Cooper ligaments, and pectoral ligaments are becoming defined. The large arrows point to Cooper ligaments, and the small arrows point to pectoral ligaments (H&E, 2×). This figure, 35.15, 35.16, and 35.17 are from the same breast

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the 15–20 segmental lactiferous ducts and a few major branches (Fig. 35.27). The breast decreases in size but may be palpable and up to 1 cm in diameter at 6 months of age [7, 8]. The definitions of the normal size of the breast bud, the duration of the stage of hyperplasia and secretion, and the duration and severity of cystic change are not well estabFig. 35.15  Complex branching of mammary ducts: gestational age 34 4/7 weeks: higher magnifications of Fig. 35.14. Note the continuing differentiation of the loose intralobular stroma and wisps of denser interlobular stroma in this figure and the following figures (a H&E, 4×; b H&E, 10×)

a

b

lished. Cystic change may become severe and persistent, and secretory changes may persist in the thin epithelial linings of the cysts. The distinction between persistent cystic regression and galactocele may be difficult. After neonatal regression is complete, the breast remains inactive until thelarche.

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Fig. 35.16  Higher magnification of primary ducts. (a) This segment of the primary duct is superficial to the layer of smooth muscle. The proximal end of the duct (top) is lined by squamous epithelium. Distally (bottom), the myoepithelium transforms from an inconspicuous layer of flattened cells to a prominent layer of cuboidal cells, and the luminal epithelium is composed of one or more layers of flattened, eosinophilic

a

Fig. 35.17  Primary duct entering the intralobular stroma. (a) Primary duct deep to the layer of smooth muscle. The morphology of the duct changes dramatically as it enters the intralobular stroma beneath the layer of smooth muscle. The myoepithelial cells are flat and inconspicuous, whereas the luminal epithelial cells are tall and columnar and dem-

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cells (H&E, 20×). (b) This is the same duct as it penetrates through the layer of smooth muscle. Its proximal end (top) overlaps the distal end of the duct in (a). The myoepithelial cells transform from the cuboidal cells in (a) to inconspicuous, flat cells, and the luminal epithelial cells become prominent, cuboidal cells (H&E, 20×)

b

onstrate apical projections and blebs. The lumen contains secretory material. There is mild extramedullary hematopoiesis in the intralobular stroma (H&E, 20×). (b) Branching ends of an intralobular duct. The end buds have lumens containing secretions, and the myoepithelial cells are prominent (H&E, 20×)

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Fig. 35.19  Neonatal hyperplasia and secretion: gestational age 38 6/7 weeks: higher magnification of Fig. 35.18. Intralobular branching of the ducts is extensive and complex. Extramedullary hematopoiesis is prominent in the intralobular stroma (H&E, 10×)

Fig. 35.18  Neonatal hyperplasia and secretion: gestational age 38 6/7 weeks. The diameter of the breast bud is 7 mm. The lobular appearance is exaggerated by the enlargement of the lobules and the broad, dense bands of interlobular stroma. The capsule of the breast, Cooper ligaments, and pectoral ligaments are more prominent. Fat is present in the lobules and interlobular septa. The large arrows point to Cooper ligaments, and the small arrows point to pectoral ligaments (H&E, 1×). The next two figures are from the same breast

Fig. 35.20  Neonatal hyperplasia and secretion: gestational age 38 6/7 weeks: higher magnification of the terminal branches of the intralobular ducts shown in Fig.  35.19. Secretory changes are prominent (H&E, 20×)

Fig. 35.21  Additional variation of the morphology of myoepithelial cells: gestational age 35 0/7  weeks. This primary duct is deep to the layer of smooth muscle of the nipple and areola. The spindle-shaped myoepithelial cells with eosinophilic cytoplasm display distinct myoid differentiation. To the right of and parallel to the duct is a band of myoid myoepithelial cells from the adjacent duct cut en face (H&E, 20×)

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Fig. 35.22 Apocrine metaplasia: gestational age 36 5/7  weeks. Apocrine metaplasia of mammary ducts is common in near-term breasts and becomes more extensive prior to and during regression (H&E, 4×)

Fig. 35.23  Atypical giant cells in the interlobular stroma: gestational age 41 2/7 weeks. The frequency and significance of this change is not known. Extramedullary hematopoiesis is also present (H&E, 20×)

Fig. 35.24  Near-term hyperplasia with early regression: gestational age 41 0/7 weeks. The baby lived 9 days. The diameter of the breast bud is 1.5  cm. Cystic change is an early indication of regression (H&E, 0.5×)

Fig. 35.25  Neonatal regression: gestational age 42 0/7  weeks. The baby lived 9 days. The diameter of the breast bud is 1.2 cm. Diffuse cystic change is common during regression (H&E, 0.5×)

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Fig. 35.26  Exaggerated cystic change during regression: gestational age 39 0/7 weeks. The baby lived 39 days. The diameter of the breast bud is 1.7 cm. The difference between normal and abnormally exaggerated cystic change during regression is not well defined, but this case is unusual in both the severity and duration of the cystic change. This degree of exaggerated cystic change of regression may be difficult to distinguish from a galactocele (H&E, 0.5×)

References 1. O’Rahilly R, Müller F.  The origin of the ectodermal ring in staged human embryos of the first 5 weeks. Acta Anat (Basel). 1985;122:145–57. 2. Hughes ESR. The development of the mammary gland. Ann R Coll Surg. 1950;6:3–23. 3. Friedrichs N, Steiner S, Buettner R, Knoepfle G. Immunohistochemical expression patterns of AP2a and AP2g in the developing fetal human breast. Histopathology. 2007;51:814–23.

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Fig. 35.27  Neonatal regression: gestational age 25 5/7  weeks. The baby lived 7 months. The diameter of the breast bud is 2 mm. The distal branches, end buds, and intralobular stroma disappear, leaving only primary lobular ducts and dense interlobular stroma. Cooper and pectoral ligaments remain. The large arrows point to Cooper ligaments, and the small arrows point to pectoral ligaments (H&E, 1×)

4. Sternlicht MD.  Key stages in mammary gland development. The cues that regulate ductal branching morphogenesis. Breast Cancer Res. 2006;8:201–11. 5. Jolicouer F, Gaboury LA, Oligny LL. Basal cells of second trimester fetal breasts: immunohistochemical study of myoepithelial precursors. Pediatr Dev Pathol. 2003;6:398–413. 6. Jolicouer F. Intrauterine breast development and mammary myoepithelial lineage. J Mammary Gland Biol Neoplasia. 2006;10:199–210. 7. McKiernan JF, Hull D.  Breast development in the newborn. Arch Dis Child. 1981;56:525–9. 8. Anbazhagan R, Bartek J, Monaghan P, Gusterson BA.  Growth and development of the human infant breast. Am J Anat. 1995;192:407–17.

Part X Placenta

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Placenta Linda M. Ernst and Chrystalle Katte Carreon

Introduction The placenta is the organ of fetal-maternal exchange during gestation. It is a fetal organ by embryologic origin but has close association with the maternal tissues of the endometrium. Thus, it is unique among the fetal organs in that maternal tissues are also a significant part of the histological component of the placenta. An examination of the placenta is an important component of a thorough investigation into the cause of fetal death and may help with an understanding of neonatal morbidities. The histology and pathology of the placenta are best understood using a systematic approach to the several anatomic/ histological compartments of the placenta. These include the extraplacental membranes, the umbilical cord, the chorionic plate or fetal surface, the basal plate or maternal surface, and the villous parenchyma. These regions correspond to the usual samples of the placenta submitted for microscopic examination in a thoroughly studied placenta. Knowledge of the normal histological appearance of each of these compartments at a variety of gestational ages is necessary to interpret any pathologic changes.

Embryology The earliest trophoblastic cells of the placenta are recognizable by the 12- to 16-cell stage as the outer cell mass of the developing cellular sphere known as the morula (Fig. 36.1a). L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected] C. K. Carreon Department of Pathology and Laboratory Medicine, The Children’s Hospital of Philadelphia and Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA e-mail: [email protected]

By the blastocyst stage, the trophoblast cells of the outer cell mass expand and flatten to form the outer layer of the blastocyst that encloses the inner cell mass, now referred to as the embryoblast, at one pole (Fig. 36.1b). At the end of the first week of gestation, the trophoblast cells situated along the embryonic pole of the blastocyst reach and begin to invade the endometrial lining. Shortly after beginning to invade the endometrium, differentiation of cellular phenotypes is seen within the trophoblast layer (Fig. 36.1c). An outer layer of syncytiotrophoblast and inner layer of cytotrophoblast are formed [1, 2]. The cytotrophoblast cells represent the replicative layer of trophoblast and maintain a simple layer of epithelial cells, whereas the syncytiotrophoblast layer is a multinucleated syncytium formed by fusion of numerous trophoblastic cells. The syncytiotrophoblast layer is thickest at the embryonic pole and decreases in thickness around the circumference of the blastocyst (see Fig. 36.1c). With continued blastocyst invasion into the endometrium, lacunae begin to form within the syncytiotrophoblast layer (Fig.  36.1d). These lacunae begin to coalesce and form an anastomosing network of sinusoidal spaces, which become filled with maternal blood, as the leading invasive trophoblast cells reach and invade maternal endometrial blood vessels. Thus, the beginning of the maternal circulation to the placental tissues is established at approximately 12 days gestation [1, 2]. By the end of the second week of development, the formation of primary chorionic villi begins, as cytotrophoblast cells proliferate as short columns protruding outward, still covered by the syncytiotrophoblast. In the third week of development, mesodermal cells, derived from the extraembryonic mesoderm, infiltrate the cytotrophoblastic cores of the primary villi and produce structures known as secondary villi, which are characterized by a mesodermal core lined by cytotrophoblast cells, all covered by syncytiotrophoblast. The next step in villous development is the formation of a vascular network within the mesodermal core, the characteristic of tertiary villi (Fig.  36.2). The blood vessels formed within the tertiary villi become linked to blood vessels developing in the

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_36

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Fig. 36.1  Early placedevelopment. (a) The morula at 3–4 days after fertilization is characterized by an inner cell mass that will form the embryo (blue) and outer cell mass that will form the placenta (green). (b) During the blastocyst stage, the cells of the outer cell mass, the trophoblast (green), will form the placenta. (c) Trophoblast differentiation

Fig. 36.2  Comparison of the components of primary, secondary, and tertiary villi. BV fetal blood vessels, C cytotrophoblast, M mesenchyme, S syncytiotrophoblast

d occurs shortly after the blastocyst implants on the endometrium. The outer layer is the syncytiotrophoblast (dark green), and the inner layer is the cytotrophoblast (light green). (d) As the blastocyst continues to invade the endometrium, lacunae form within the syncytiotrophoblast layer and contain maternal blood (red)

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chorionic plate and connecting stalk to complete the formation of the fetoplacental circulation. The formation of the amnionic and chorionic cavities (Fig. 36.3) occurs simultaneously with villous development. At the beginning of the second week of development, the amnionic cavity begins as a small cavity within the epiblast layer of the bilaminar germ disc formed from the embryoblast [1, 2]. The cells lining the roof of the amnionic cavity are in continuity with the cytotrophoblast shell surrounding the embryo and are called amnioblasts. The cells of the other layer of the bilaminar embryo, the hypoblast, form the lining of a second cavity, the primitive yolk sac. The lining of the primitive yolk sac is also initially in contact with the cytotrophoblast. Subsequently, mesodermal cells derived

Secondary villi

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from the primitive yolk sac begin to proliferate and fill the space between the two existing sacs (amnionic and primitive yolk sacs) and the cytotrophoblast (see Fig. 36.3a). This mesodermal layer is known as the extraembryonic mesoderm. Later, cavities begin to form within the extraembryonic mesoderm, and these numerous cavities coalesce to form a new cavity, the chorionic cavity (or extraembryonic coelom), which surrounds the amnionic cavity and yolk sac cavity except where the connecting stalk—the future umbilical cord linking fetal and placental circulations—is present (see Fig. 36.3b). With this development, both the amnionic and chorionic linings contain a mesodermal layer of connective tissue derived from the extraembryonic mesoderm, also referred to as the extraembryonic somatopleuric meso-

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c

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Fig. 36.3  Formation of the amnionic cavity, chorionic cavity, and umbilical cord. (a) Day 12. The amnionic cavity (AC) begins as a small cavity within the epiblast layer (E). The cells lining the cavity will form the amnion (A). The cells of the hypoblast form the lining of the primitive yolk sac (PY). The extraembryonic mesoderm (EM) fills in the space between the two existing sacs (amnionic and primitive yolk sacs) and the cytotrophoblast (T). (b) Day 14. A cavity has now formed within the extraembryonic mesoderm, the chorionic cavity (CC) or extraembryonic coelom, and the extraembryonic mesoderm (EM) is present on the inner surface of the trophoblast and surrounds the yolk sac and amnionic sac

(AC). The connecting stalk (CS) is also beginning to develop. (c) Day 28. As the amnionic sac expands to fill the chorionic cavity, the mesodermal tissues of the amnion (AM) and chorion (CM) come in close contact. The secondary yolk sac (SY), allantois (AL), and connecting stalk (CS) are developing. The placental tissue is forming villous structures in the chorion frondosum (CF). (d) Day 40. As the trophoblasts continue to develop and invade the endometrium, the placental disc is becoming more obvious and the villi of the chorion laeve (CL) atrophy. Note the continued maturation of the amnionic cavity (AC) and chorionic cavity (CC), which surround the connecting stalk as the embryo folds

derm. This mesoderm will form the definitive connective tissue layers of the membranes and chorionic plate. As the amnionic sac expands to fill the chorionic cavity, the mesodermal tissues of the amnion and chorion are in close contact (see Fig. 36.3c). The primitive yolk sac is replaced by the smaller definitive or secondary yolk sac; in the process, small exocoelomic cysts are produced and persist in the chorionic cavity

between the chorion mesoderm and the amnion mesoderm. Around the fifth week of gestation, the yolk sac stalk containing the vitelline vessels becomes incorporated into the connecting stalk, the future umbilical cord along with the allantois, and the umbilical vessels (see Fig. 36.3d) [1, 2]. In embryonic life, intestinal loops also protrude into the connecting stalk along the opening between the midgut and the yolk sac, the omphalomesenteric duct. By the end

402 Fig. 36.4  Placental anatomy. Anatomy of the placenta, including the extraplacental membranes (EPM), composed of amnion (A), chorion (C), and parietal decidua (PD); the umbilical cord; the chorionic plate; the basal plate; and the villous parenchyma

L. M. Ernst and C. K. Carreon PD C A

EPM Umbilical cord Chorionic plate

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of the embryonic period, however, the yolk sac is obliterated, and the intestines return to the abdomen. The umbilical cord then contains the umbilical vessels surrounded by Wharton’s jelly, but remnants of the embryonic structures within the umbilical cord are seen commonly in fetal life.

Histology By the beginning of the midtrimester, the placental anatomy is established, but the tissues continue to undergo maturational changes throughout the fetal period. Figure 36.4 reviews the structural anatomy of the compartments of the placenta: the extraplacental membranes, the umbilical cord, the chorionic plate, the basal plate, and the villous parenchyma.

Extraplacental Membranes Histological sections of the human extraplacental membranes are composed of three main tissue layers that surround the amnionic sac: the fetally derived amnion, the fetally derived chorion, and the maternally derived decidua. Over the gestational period, all three layers remain present, with only minor changes in thickness or epithelial appearance (Figs. 36.5, 36.6, and 36.7). The amnion consists of a simple squamous to cuboidal epithelial layer, which is in contact with the amnionic fluid. The epithelium has an underlying basement membrane, and a thin connective tissue layer is present beneath the basement membrane. The paucicellular connective tissue layer of the amnion is composed of collagen and elastic fibers, with a few identifiable fibroblasts and occasional macrophages. These macro-

Fig. 36.5  Membranes at 15 weeks gestation. Note the simple, nearly squamous epithelial layer of the amnion at the upper limit of the image. The amnion and chorionic connective tissues are back-to-back, and there is a somewhat thin trophoblastic layer in the chorion. Note the atrophic villi of the chorion laeve (arrows). The decidua is the layer at the lower edge of the image (hematoxylin and eosin (H&E), 10×)

phages will ingest pigment in abnormal situations such as intra-amnionic meconium or blood. No blood vessels, nerves, or lymphatics are present in the connective tissue of the amnion. As gestation approaches term, the amnion epithelial cells may appear slightly more columnar and reactive. Intraamnionic irritants, such as meconium, can cause marked and bizarre amnion epithelial cell alterations (Figs. 36.8 and 36.9). The amnion and chorion are imperfectly fused [3, 4], and a potential space exists between the two, which can be seen histologically as a gap between the amnion and chorion connective tissue layers. Embryologically, this space represents the remnant of the extraembryonic coelom. This potential space is referred to as the spongy layer and is composed of

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Fig. 36.6  Membranes at term. The amnion is slightly more cuboidal at this gestational age but is still relatively flattened. The chorionic trophoblast is somewhat thicker but still shows the atrophic villi of the chorion laeve. Note that vacuolation/clearing of the cytoplasm of some of the chorionic trophoblast cells is seen. A clear distinction can be made between the decidual cells, with polygonal, pale cytoplasm, and the more amphophilic cytoplasm of the chorionic trophoblast (H&E, 10×)

Fig. 36.7  Membranes at term. Higher-power view of the membrane components. Note the potential space between the amnion and chorion, with the spongy layer containing loose collagen fibers. Trophoblast cells of the chorion have variable sizes and shapes, and occasional multinucleated cells are seen (H&E, 20×)

Fig. 36.8 Membranes at 32  weeks gestation with reactive amnionic changes. The amnion epithelium is columnar, and many of the amnionic nuclei are apically placed, instead of the normal basal location. This is a reactive change that is commonly seen in the amnion; in this case, numerous hemosiderin-laden macrophages, indicative of previous hemorrhage, are noted in the amnion and chorionic connective tissue (H&E, 20×)

Fig. 36.9  Higher-power view of reactive amnionic changes. Note that several of the nuclei are apically placed, whereas others remain at the basal aspect of the cell, giving the epithelium a somewhat pseudostratified appearance (H&E, 60×)

loosely arranged collagen fibers (see Fig. 36.7), which sometimes have a myxoid or edematous appearance and contain scattered fibroblasts and macrophages [3]. The chorionic layer of the membranes is known as the chorion laeve. The chorion laeve, in general, is also composed of epithelial and connective tissue layers, but compared with the amnion, the layers in the chorion are reversed in orientation. Therefore, the connective tissue layer of the chorion is adjacent to the connective tissue layer of the amnion. The chorionic connective tissue is slightly thicker than the amnionic

connective tissue, but it has a similar appearance of paucicellular connective tissue composed of collagen and elastic fibers with occasional macrophages. Although the chorion laeve is in continuity with the vascular chorionic plate, the chorion laeve itself is avascular. Therefore, all fetally derived tissues in the extraplacental membranes are avascular. The epithelial layer of the chorion has several layers of trophoblastic cells, and the thickness of the trophoblastic cell layer is variable. Clearing of the cytoplasm of the trophoblastic cells of the chorion is not uncommon and sometimes

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Fig. 36.10  Decidua parietalis at term. Note the muscularized decidual vessels (arrows). Because these vessels are not normally remodeled by extravillous trophoblast cells, they frequently retain a thin coat of smooth muscle (H&E, 10×)

imparts a vacuolated appearance to this layer. The trophoblastic cells of the extraplacental membranes are also known as chorionic-type intermediate trophoblast cells and have a distinct immunohistochemical phenotype, which includes expression of p63, human placental lactogen (hPL), HLA-G, CD146 (Mel-CAM), mucin-4, and cyclin E [5]. In addition, the chorionic epithelial layer contains atrophied villi, which appear as well-circumscribed, slightly basophilic nodules throughout the layer (see Figs. 36.5 and 36.6). The maternally derived tissue layer of the extraplacental membranes is the decidua. Until about 20 weeks, this layer is referred to as decidua capsularis, the initial abembryonic decidual covering of the implanted blastocyst. When the decidua capsularis fuses with the decidua parietalis of the opposite side of the uterus, the decidua is then referred to as decidua parietalis, but there is no distinct histological difference [3]. The decidua of the membranes is the only vascularized tissue of the membranes, and it has both a superficial and a deep vascular component. The maternal vasculature in the decidua parietalis does not undergo invasion and remodeling by extravillous trophoblast cells. Therefore, parietal decidual vessels can and frequently do contain smooth muscle (Fig.  36.10). However, excessive mural thickening or hypertrophy of these membranous maternal vessels, defined as mean wall diameter more than one third the mean circumference, is considered one of the pathologic features of maternal vascular malperfusion [6, 7]. Other changes, such as fibrinoid necrosis, acute atherosis, and thrombosis, should not be considered normal within the parietal decidual vessels. Lastly, inflammatory cells are also seen in the decidua, with occasional macrophages and foci of lymphocytes as normal components of the decidua (Fig.  36.11), but plasma cells are not a normal component of the decidua.

L. M. Ernst and C. K. Carreon

Fig. 36.11  Decidua parietalis at term. Medium-power view of the decidua in the membranes. Note the fairly thin-walled vessels in both the superficial and deep layers of the decidua. Also note the scattered lymphocytes, which are a normal component of the decidua. The lower edge of the image contains a few adherent myometrial fibers (H&E, 20×)

Umbilical Cord The umbilical cord is the conduit for the fetal vasculature between the placenta and the fetus. The main components of the umbilical cord are the fetal umbilical vessels, two umbilical arteries, and a vein, surrounded by a gelatinous, shock-­ absorbing substance known as Wharton’s jelly (Fig.  36.12). The outermost covering of the umbilical cord, which is in contact with amnionic fluid and is firmly attached to the cord [8], is histologically the same as the amnion epithelium with its underlying basement membrane (Fig. 36.13). However, squamous metaplasia of the umbilical cord epithelium is considered normal (Fig. 36.14) and is especially common near the fetal body wall. The Wharton’s jelly (see Figs. 36.13 and 36.14) consists of mucopolysaccharides, such as hyaluronic acid and chondroitin sulfate, which give a slightly basophilic appearance to the jelly on H&E-stained sections. Sparse collagen fibers and other microfibrils are also present, produced predominantly by myofibroblasts located at the periphery of the umbilical vessels and beneath the basement membrane of the epithelium [8]. Scattered macrophages and mast cells are present within the substance of Wharton’s jelly [8]. Hemorrhage is not a normal finding in Wharton’s jelly, but iatrogenic umbilical vessel injury frequently occurs secondary to umbilical cord clamping or postpartum sampling for umbilical cord blood. Consequently, acute perivascular hemorrhage is a frequent artifact seen in routine sections of the umbilical cord (Fig. 36.15). Organization of the hemorrhage, including layering of the clot (lines of Zahn) or neutrophil infiltration, is not seen with artifactual hemorrhage.

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Fig. 36.12  Umbilical cord at 16  weeks gestation. Low-power view showing the single umbilical vein (UV) and two umbilical arteries (UA). A small allantoic remnant (arrow) is also present between the two arteries (H&E, 2×)

Fig. 36.13  Umbilical cord at 24 weeks gestation. Medium-power view of the external surface of the cord, showing the amnion epithelium lining the cord and the paucicellular Wharton’s jelly beneath it (H&E, 20×)

Fig. 36.14  Umbilical cord at 36 weeks gestation. Medium-power view of the external surface of the cord, showing squamous metaplasia of the external surface of the umbilical cord and the paucicellular Wharton’s jelly beneath it (H&E, 20×)

Fig. 36.15  Umbilical cord at 40 weeks gestation. Low-power view of the umbilical cord showing acute perivascular hemorrhage surrounding an umbilical artery, a common artefactual finding. Note the lack of any layering or organization of the blood clot (H&E, 10×)

Selection of samples of the umbilical cord should avoid areas with umbilical cord clamp marks or obvious puncture marks. The umbilical arteries possess two smooth muscle layers, an outer circular layer and an inner spiral longitudinal layer, so they have generally thicker walls than the vein, which has only a single layer of circular smooth muscle (Figs. 36.16 and 36.17). In the arteries, occasional elastic fibers may be seen in the media, but the umbilical arteries generally lack a well-formed internal elastic lamina. The vein, however, has a well-formed internal elastic lamina (Figs.  36.18, 36.19, and 36.20) [8, 9]. There are no vasa vasorum, lymphatic channels, or nerves within the umbilical cord [8]. There have been descriptions of ganglion cells [10] and nerve fibers [10, 11] at the fetal end of the cord, however.

The smooth muscle cells of the umbilical vessels may have nuclei that are somewhat squiggle-shaped, and care should be taken not to mistake these cells for neutrophils and overdiagnose acute umbilical vasculitis; this can be especially tricky in macerated midtrimester fetal umbilical cords (Fig.  36.21). One helpful clue is that neutrophil nuclei are typically seen between the smooth muscle cells in the interstitial space. The normal umbilical cord commonly contains embryonic remnants of either the allantoic duct or the vitelline duct. Vitelline (omphalomesenteric) duct remnants are commonly present at the fetal end of the cord, but they can also be seen at the placental end (Fig. 36.22). These remnants are composed of endodermally derived epithelial elements of the vitelline duct or of the vessels associated with the duct. Epithelial remnants usually possess a lumen lined by mucin-­ producing cuboidal to columnar epithelial cells, sometimes

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Fig. 36.16  Umbilical cord at 16 weeks gestation. Comparison of the layers of umbilical vein (a) and umbilical artery (b). Note that, at this gestational age, the muscle layer is relatively thin, and distinction between vein and artery can be difficult (H&E, 10×)

Fig. 36.17  Umbilical cord at 35 weeks gestation. Comparison of the layers of umbilical vein (a) and umbilical artery (b). The umbilical artery has two smooth muscle layers and a slightly thicker wall than the vein, which has single layer of circular smooth muscle (H&E, 10×)

Fig. 36.18  Umbilical vein at term, showing prominent internal elastic lamina (Verhoeff Van Gieson, 4×)

Fig. 36.19  Umbilical vein at term. Higher-power view showing prominent internal elastic lamina (Verhoeff-Van Gieson, 10×)

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Fig. 36.20  Umbilical artery at term. Umbilical artery with scattered peripheral elastic fibers but no internal elastic lamina (Verhoeff-Van Gieson, 20×)

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Fig. 36.21  Umbilical cord at 16  weeks gestation, after intrauterine fetal demise. After fetal death, the smooth muscle cells of the media of the umbilical vessels undergo degenerative changes, and their cytology is altered. Note that many of the nuclei appear pyknotic and wrinkled, mimicking neutrophils. These degenerative changes should not be misinterpreted as evidence of acute inflammation (H&E, 40×)

accompanied by a muscular wall (Figs.  36.23 and 36.24), but other endodermally derived tissues such as hepatocytes may be seen [12]. Vitelline vessel remnants may also accompany the vitelline duct remnant near the fetal end of the cord and may appear as a single, thin-walled vessel or sometimes as a plexus of vessels (Figs. 36.25 and 36.26). Allantoic remnants are also typically present at the fetal end of the cord between the two umbilical arteries (Fig. 36.27). They are composed of transitional-type epithelial cells arranged in a spherical configuration, usually without a recognizable lumen.

Chorionic Plate The chorionic plate, the fetal surface of the placenta, is covered by a layer of amnionic tissue that is identical to the amnion of the extraplacental membranes. As stated earlier, the amnion and chorion are imperfectly fused, and the amnion layer is easily stripped from the surface of the chorionic plate, so sometimes it is not present in histological sections taken from the fetal surface. Particularly on the fetal surface of the placenta, this potential space between the amnion and chorion may become filled with fresh hemorrhage known as a subamnionic hemorrhage or hematoma. Usually, this represents artifactual fetal hemorrhage associated with tension placed on the umbilical cord in the third stage of labor, with tearing of the fetal vessels near the umbilical cord insertion site. The chorionic plate represents a segment of the continuous layer of chorionic tissue surrounding the fetus. It dif-

Fig. 36.22  Gross picture of the umbilical cord with a vitelline vessel remnant. This gross picture shows a remnant of the vitelline vessel extending from the yolk sac remnant (arrow) on the fetal surface of the placenta, coursing over the chorionic plate and into the Wharton’s jelly of the umbilical cord (arrowhead)

fers from the chorion laeve because the chorionic plate has a thicker connective tissue containing fetal vessels that ramify from the umbilical cord onto the fetal surface (Figs. 36.28, 36.29, 36.30, 36.31, 36.32, and 36.33). The fetal vasculature of the chorionic plate consists of muscularized fetal vessels, but distinguishing arteries from veins based on morphologic histological features on routine H&E stains is not consistently possible [4]. Fetal vessels of the chorionic plate frequently have asymmetrical thinning of the muscular coat, especially in superficial segments of the vessels (see Fig. 36.30). In addition, it is not

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Fig. 36.23  Umbilical vitelline duct remnant. Low-power view of an umbilical cord demonstrating a remnant of the vitelline/omphalomesenteric duct (arrow) located centrally in the cord between the umbilical vein (UV) and the two umbilical arteries (UA) (H&E, 4×)

Fig. 36.24  Umbilical vitelline duct remnant. This higher-power view of the vitelline/omphalomesenteric duct remnant shows cuboidal to columnar lining cells with focal mucin production, resembling the epithelium of the gut (H&E, 20×)

Fig. 36.25  Umbilical vitelline vessel remnant. Low-power view of the vitelline/omphalomesenteric vessel remnant, showing two large, thin-­ walled vascular structures and a plexus of smaller vessels (H&E, 10×)

Fig. 36.26  Umbilical vitelline vessel remnant. Higher-power view of the vitelline/omphalomesenteric vessel remnant present at term (H&E, 40×)

unusual to see myxoid intimal cushions in fetal chorionic vessels, but in the absence of subendothelial/intramural fibrin deposition or calcification, these myxoid cushions are not considered histological evidence of fetal vascular obstruction [7, 13]. The connective tissue of the chorionic plate, which surrounds the fetal vessels, is composed of collagen layers and is generally paucicellular. The few cells present consist of fibroblasts and occasional macrophages. Beneath the connective tissue of the chorionic plate is a layer of trophoblastic cells, which varies in thickness but is usually just a few cells thick. These trophoblastic cells line the subchorionic intervillous space. A subchorionic fibrin layer (known as Langhans’ fibrin), also of variable thickness, forms beneath the trophoblastic layer [14]. In general, with

increasing gestational age, the subchorionic fibrin layer shows areas of increasing thickness forming subchorionic fibrin plaques (see Figs.  36.31 and 36.32; compare with Figs. 36.29 and 36.30). Stem villi arise from the lower surface of the chorionic plate (see Fig. 36.33) and give rise to the remainder of the villous parenchyma beneath the chorionic plate.

Basal Plate The basal plate is present on the maternal aspect of the delivered placenta and is a relatively thin layer, measuring up to 0.15 cm in thickness [14]. It covers the maternal aspect of the placenta and has a smooth, grayish gross appearance. The basal plate represents an intimate mixture of both maternal

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Fig. 36.27  Allantoic remnant in the umbilical cord (H&E, 20×)

Fig. 36.28  Chorionic plate at 21 weeks gestation. The amnionic layer is present (top). The chorionic connective tissue layer is thicker than in the membranes and contains the fetal chorionic vessels (H&E, 4×)

Fig. 36.29  Chorionic plate at 21 weeks gestation. This higher-power image shows the trophoblastic lining of the lower border of the chorionic plate at the interface with intervillous space. Subchorionic fibrin is not prominent at this gestational age (H&E, 10×)

Fig. 36.30  Chorionic plate at 32 weeks gestation. Low-power view of the chorionic plate (fetal surface of the placenta). The amnionic layer is present, and subchorionic fibrin is more prominent. Note that there is frequently asymmetry of the chorionic plate vessel wall, with thinning noted along the upper surface of the vessels (H&E, 2×)

and fetal tissues at their closest point of contact. With careful gross inspection, maternal decidual arterioles can sometimes be identified on the surface of the basal plate as curlicue-­ shaped structures (Fig. 36.34). An understanding of the normal components of the basal plate is important for understanding and interpreting pathologic alterations in this important anatomic compartment. The main components or layers of the basal plate are listed in Table 36.1 and shown in Fig. 36.35. The inner layer of the basal plate is adjacent to the intervillous space and is lined by syncytiotrophoblast cells focally, with maternal endothelial cells where the trophoblasts are absent. There are two fibrinoid layers on either side of the principal layer of the basal plate. Both of these layers Fig. 36.31  Chorionic plate at term. Low-power view of the chorionic plate between fetal vessels. Note the presence of a fairly thick layer of subchorionic fibrin (H&E, 10×)

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Fig. 36.32  Chorionic plate at term. Higher-power view of the chorionic plate between fetal vessels. Note the presence of a fairly thick layer of subchorionic fibrin, with trophoblastic cells present within the fibrin. Inflammatory cells are not normally present within this fibrin layer (H&E, 20×)

appear as discontinuous or interrupted, pale, eosinophilic acellular layers in routine H&E-stained sections at term. The innermost fibrinoid layer is known as Rohr’s fibrinoid, and the outer layer is Nitabuch’s fibrinoid [14]. In some areas of the inner surface of the basal plate, Rohr’s fibrinoid is exposed and in contact with the components of the intervillous space; it is therefore continuous with perivillous fibrin (Fig. 36.36). Nitabuch’s fibrinoid is on the opposite side of the principal layer and is considered the boundary between maternal and fetal tissues of the placenta because it separates the most superficial extravillous trophoblasts of the principal layer from the maternal decidual cells. Although Nitabuch’s fibrinoid represents the maternal-fetal boundary, it is by no means a blockade to fetal extravillous trophoblast cells. Trophoblast cells are frequently seen well beyond Nitabuch’s layer [14]. Additional tissue is usually attached to the delivered placenta deep to Nitabuch’s fibrinoid; this tissue is known as the separation zone [14]. It contains predominantly decidualized endometrial stromal cells, with occasional extravillous trophoblast cells (Fig. 36.37). The principal layer of the basal plate is the thickest layer, occupying 50–75% of the basal plate at term [14]. Within this layer, the anchoring villi are seen as somewhat atrophic-­ appearing villous structures embedded in fibrin. Anchoring villi are the most distal extensions of stem villi, and their “heads” are anchored in the basal plate. From the abembryonic surface of anchoring villi, trophoblast cells proliferate and extend into the basal plate as the so-called trophoblast cell columns. Extravillous trophoblast cells emanate from these cell columns and then migrate through the basal plate, endometrium, and myometrium to perform their main duty of maternal vascular remodeling (see Fig. 36.37). Maternal tissues are also a component of the principal layer of the basal plate, with the main cell consisting of the decidualized endometrial stromal cell. Other endometrial stromal cells are also present, including the vasculature of the endometrial stroma, fibroblasts, and occasional macrophages. Scattered

L. M. Ernst and C. K. Carreon

Fig. 36.33  Chorionic plate at term. Higher-power view of the lower chorionic plate, showing the origin of a stem villus from the chorionic plate (H&E, 20×)

Fig. 36.34  Basal plate at 28  weeks gestation, with visible maternal basal decidual arterioles. This gross image is a close-up of the basal surface of a preterm placenta. Note the overall shiny gray appearance of the basal plate. The yellow regions represent areas of underlying villous infarction. Curlicue-shaped structures are seen in clusters and represent the maternal spiral arterioles of the endometrium, which are remodeled by extravillous trophoblast cells Table 36.1  Components of the basal plate from intervillous space to endometrium Lining of maternal endothelial cells and syncytiotrophoblast Rohr’s fibrinoid Principal layer  Fetal tissues:    Extravillous trophoblast cells    Nonproliferative trophoblast cells emanating from cell columns    Anchoring villi    Buried cell columns  Maternal tissues:   Decidual cells   Connective tissue (endometrial stroma, vessels, fibroblasts, macrophages) Nitabuch’s fibrinoid Separation zone  Decidua and other components of endometrial stroma  Occasional extravillous trophoblast cells  Multinucleated extravillous trophoblast (placental site giant cells)

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Fig. 36.35  Basal plate at term. Low-power view of the basal plate showing the major constituents of the maternal-fetal interface. The intervillous space (at top of basal plate) is lined by maternal endothelial cells and syncytiotrophoblast. Top arrow indicates Rohr’s fibrinoid. The principal layer (PL; between the two arrows) contains anchoring villi (av) from which the trophoblast cells emanate. Other cells present include fibroblasts, macrophages, and decidua. Lower arrow indicates Nitabuch’s fibrinoid. The separation zone (SZ) contains decidua and other stromal components, with occasional trophoblast cells (H&E, 4×)

lymphoid and natural killer cells are also seen in the normal basal plate [14], but plasma cells are not considered a normal component of the decidua. The process of maternal vascular remodeling by the trophoblasts is essential for obtaining adequate perfusion to the intervillous space and, thus, to the fetus. The effects of this remodeling process are evident in histological samples of the basal plate, and alterations from the expected normal appearance of these vessels may be indicative of a pathologic process. Extravillous trophoblast cells first reach the spiral arterioles of the endometrium in the first trimester, between 6 and 12  weeks gestation (the first wave of invasion) [15– 17]. Trophoblast cells enter the vascular lumen and produce factors that degrade the smooth muscle and elastic tissue of the vascular media. The vascular infiltration of the spiral arterioles by the extravillous trophoblast cells is so profound in this stage as to temporarily plug the distal vessel lumens, thereby minimizing the maternal blood flow to the developing placenta and reducing oxidative damage at this critical point in embryologic development [18, 19]. During the fetal period, especially in the midtrimester, the process of vascular remodeling is ongoing, with the second wave of trophoblast invasion occurring in the early midtrimester (16–22 weeks; Fig. 36.38) [15–17]. The extravillous trophoblast cells reach as far as the myometrial segments of the spiral arterioles by this time [17]. Therefore, in the midtrimester, it is not unusual to see partially remodeled arterioles or arterial lumens containing endovascular trophoblast cells (Fig. 36.39). As gestation progresses, however, evidence of such ongoing remodeling should be less frequent, and most maternal vessels should appear as completely remodeled spiral arterioles (Fig.  36.40). The degraded medial tissues are replaced by fibrinoid material containing embedded extravillous trophoblast cells but devoid of smooth muscle or elastic tissue. With the degradation of the medial tissues, there is also a

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Fig. 36.36  Basal plate at term. Inner aspect of the basal plate, showing the lining of the intervillous space, with maternal endothelial cells, syncytiotrophoblast, and bare Rohr’s fibrinoid/intervillous fibrin (H&E, 20×)

Fig. 36.37  Basal plate at term. Medium-power view of basal plate showing the main constituents of the principal layer and separation zone. Note the extravillous trophoblast cells emanating from the anchoring villus (av). The cytoplasm of the trophoblast is more amphophilic than the cytoplasm of the decidua; this difference can aid in distinguishing between the two cell types. Note that extravillous trophoblast cells are seen beyond Nitabuch’s fibrinoid (H&E, 10×)

marked increase in the diameter of the vascular lumen. The cells lining these vascular channels are extravillous trophoblast cells, which acquire an endothelial phenotype (Fig. 36.41) [14, 15].

Villous Parenchyma The tissues between the chorionic plate (fetal surface) and basal plate (maternal surface) comprise the villous parenchyma. The villous parenchyma is surrounded by the intervillous space, which contains maternal blood, but the remainder of the tissue between the two placental surfaces is fetally derived chorionic villi. The basic components of all chorionic villi are an outer trophoblastic covering, vil-

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a

b

Fig. 36.38  Basal plate at 14  weeks gestation with remodeling of maternal vasculature. (a) This image shows a basal maternal decidual vessel undergoing remodeling by trophoblastic cells. Note a portion of the circumference of the vessel wall contains remnants of smooth muscle within the media (arrow), but the opposite side no longer possesses smooth muscle. (b) This image shows ongoing remodeling of maternal vasculature by extravillous trophoblast. The media of the spiral arterioles has been predominantly destroyed, but numerous extravillous trophoblast cells remain within the vascular lumina (H&E, 10×)

lous stroma, and fetal vessels. These components vary in character depending on the developmental stage of the placenta, which accounts for the dramatic histological changes seen in cut sections of the villous parenchyma over the course of normal gestation. The main alterations over time occur in the types of chorionic villi present at each developmental stage and the percentage of the various types of villi. The main villous types are mesenchymal villi, immature intermediate villi, mature intermediate villi, terminal villi, and stem villi (Fig.  36.42). Mesenchymal villi are present throughout gestation but predominate in the first trimester. Mesenchymal villi give rise to immature and

Fig. 36.39  Basal plate at 23 weeks gestation. Note the extravillous trophoblast cells emanating from the anchoring villus (AV) and the extravillous trophoblast cells within the principal layer of the basal plate. The media of the spiral arterioles has been predominantly destroyed, and a few extravillous trophoblast cells remain in the decidual vessel (DV) lumina (H&E, 10×)

Fig. 36.40  Basal plate at term. Fully remodeled maternal vessel with large luminal size; the wall is composed of fibrin and lacks smooth muscle. No residual intravascular trophoblast cells are seen (H&E, 4×)

mature intermediate villi. Immature intermediate villi become stem villi and also give rise to new mesenchymal villi. Terminal villi, the mature functional end of the villous tree, arise from mature intermediate villi. The types of chorionic villi and the major characteristics of their basic components are listed in Table  36.2 [20, 21]. The types and distribution of chorionic villi at various gestational ages are listed in Table 36.3 [20, 21]. Each of the different types of chorionic villi present in the fetal period develops from the mesenchymal villus, which is the tertiary villus of the embryologic period (see Table  36.2) and predominates in the embryonic placenta.

36 Placenta

Fig. 36.41  Basal plate at term. Higher-power view of the remodeled maternal vessel. Note the flattened lining cells and extravillous trophoblast cells embedded in the fibrin wall (H&E, 10×)

Mesenchymal villi are present in smaller numbers throughout gestation and act as the reservoir for continued villous growth and development [21]. Mesenchymal villi appear as long, slender villous structures with primitive-appearing reticular stroma with centrally placed blood vessels and a double layer of trophoblast cells, consisting of inner cytotrophoblast cells and outer syncytiotrophoblast cells (Figs. 36.43, 36.44, and 36.45). In the fetal period, they are found at the tips and along the surfaces of immature and mature intermediate villi. Stem villi are also present at all gestational stages; they represent the conduits for the larger fetal vessels and serve as the support structure for the villous tree. They extend from the main trunks beneath the chorionic plate (truncus chorii) to the basal plate, where they form the anchoring villi. Stem villi also send out side branches (rami and ramuli chorii), which extend into the placental lobule [21]. All stem villi are characterized by densely collagenized stroma and the presence of arteries and veins containing media and/or adventitia. The large stem villous trunks have the largest arteries and veins with the thickest muscular walls. The outer layer of trophoblast is typically thick and in many instances is replaced by fibrin. In the early midtrimester, the most numerous villus type is the immature intermediate villus, which represents the first generation of villi, with vessels lacking vascular media or adventitia arising from the stem villi (Fig.  36.46). Immature intermediate villi typically have a thick trophoblast layer with cytotrophoblast cells present over more than 50% of the circumference (Fig.  36.47) [21]. The stroma is described as reticular with numerous fluid-filled spaces, many containing placental macrophages or Hofbauer cells [21]. These villi generally contain centrally

413

placed fetal vessels without the formation of vasculosyncytial membranes (Fig. 36.48). By the late midtrimester, many of the immature intermediate villi have transformed into stem villi, and mature intermediate villi are formed (Figs.  36.49, 36.50, 36.51, 36.52, 36.53, 36.54, and 36.55). Mature intermediate villi exhibit a more fibrous stroma than immature intermediate villi, and they generally are slightly smaller. They also have better vascularization than immature intermediate villi, but they still have less than 50% of their stroma occupied by capillaries. Terminal villi also begin to form at the surface of mature intermediate villi in the late midtrimester. Terminal villi represent the end of the villous tree and have the greatest capacity for maternal-fetal exchange because they are highly vascularized, with more than 50% of their stroma occupied by capillaries. The number of terminal villi in a cross section of placental parenchyma increases with increasing gestational time throughout the third trimester. For instance, at 27–28  weeks gestation, mature intermediate villi still predominate, and terminal villi occupy less than 50% of the villous volume (Figs.  36.56, 36.57, 36.58, and 36.59). By 32 weeks gestation, the number of terminal villi is roughly equal to the number of mature intermediate villi (Figs. 36.60, 36.61, 36.62, and 36.63). By the late third trimester, terminal villi are the most numerous type; at term, terminal villi represent 60% of the placental cross section (Figs. 36.64, 36.65, 36.66, and 36.67) [21]. In addition to the numerous capillaries in the villous stroma, terminal villi have significant thinning of the trophoblastic layer, accompanied by sinusoidal dilatation of fetal capillaries, allowing for reduction of the diffusion distance between the fetal and maternal circulations. Vasculosyncytial membranes, which represent the diffusion barrier in terminal villi, are composed of ­ ­syncytiotrophoblast cell cytoplasm, fused basement membrane of the syncytiotrophoblast cell and fetal endothelial cell, and the fetal capillary endothelial cell cytoplasm (Fig. 36.68). Vasculosyncytial membranes appear as the thin, eosinophilic barrier, devoid of nuclei, between the dilated fetal capillary and intervillous space in H&E-stained s­ ections (see Figs.  36.61 and 36.65). Vasculosyncytial membranes begin to become noticeable in the early third trimester and increase in number as the pregnancy approaches term. At term, approximately 40% of the villous surface is composed of vasculosyncytial membranes [21]. Syncytiotrophoblastic knots or syncytial knots, defined as clumps of syncytiotrophoblastic nuclei along the villous perimeter, can be noted throughout the fetal period. Their etiology and pathogenesis is not clear, but the accumulation of these syncytial nuclei does increase with increasing gestational age (see Table 36.3) [22]. The peripheral dis-

414 Fig. 36.42  Diagram of the main villous types of the placenta. (a) Mesenchymal villus contains abundant mesenchymal cells and exhibits syncytial sprouting, a thick double-layered trophoblast, primitive stroma, and rare vessels. (b) Immature intermediate villus contains numerous macrophages (Hofbauer cells) within stromal channels, reticular stroma, and poorly developed capillaries. (c) Mature intermediate villus contains smaller vessels and capillaries occupying less than 50% of the villous stroma that is beginning to become collagenous. (d) Terminal villus has a thin trophoblast layer with vasculosyncytial membrane formation representing reduced diffusion distance between the fetal and maternal circulations; fetal capillaries occupy more than 50% of the villous stroma. (e) Stem villus, the largest of the villous types, contains large muscularized arteries and veins and rare capillaries within a densely collagenous stroma

L. M. Ernst and C. K. Carreon

a Mesenchymal villus Fetal capillary Syncytiotrophoblast Cytotrophoblast

Mesenchymal cells Syncytial sprouting

b Immature intermediate villus

Syncytiotrophoblast Cytotrophoblast Fetal capillary Stromal channels, some containing Hofbauer cells

c Mature intermediate villus

Syncytiotrophoblast Cytotrophoblast Fibrous stroma Fetal capillaries

d Terminal villus

Fetal capillary

Vasculosyncytial membrane

e Stem

villus

Fetal capillary Syncytiotrophoblast Cytotrophoblast Fibrous stroma Large muscularized artery and vein

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415

Table 36.2  Villous types and characteristics Villous type Mesenchymal villi

Size (μm) 60–100

Trophoblast Thick, double layer

Stroma Primitive stroma

Vessels/capillary density Few vessels

Immature intermediate villi

100–400

ST = 100%

Reticular stroma; numerous macrophages

Poorly developed capillaries

Fibrous stroma begins

50% capillaries

End of villous tree

Fibrous stroma

Large muscularized arteries and veins, few capillaries

Important support of villous tree

CT > 50% Mature intermediate villi Terminal villi

80–150

Stem villi

150–500

60

ST = 100% CT  mature intermediate villi Stem villi Mesenchymal villi Terminal villi 60% of cross section Stem villi Mesenchymal villi

% of villi with cytotrophoblast 80

Average % of villi with syncytial knots

60

5.8

50

8.6

45

13.2

35

13.1

25

22.5

20

29

Data from Benirschke [20] and Loukeris et al. [22]; with permission

placement of these nuclei may aid in the formation and functional significance of the fetomaternal diffusion barrier. Perivillous fibrin, the product of blood clotting on the villous surface, can be noted as a normal feature throughout gestation (see Figs. 36.52, 36.53, 36.56, 36.57, and 36.60). The deposition of perivillous fibrin increases with increasing

gestational age and is normally more prominent around stem villi, in the subchorionic region, and at the placental margin [23]. Therefore, caution should be exercised when ­interpreting the amount of perivillous fibrin in histological sections taken from these areas. Generally, the normal deposition of perivillous fibrin has an evenly distributed pattern. The deposition of perivillous fibrin should be considered abnormal when

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Fig. 36.43  Placental tissue at 6 weeks gestation. Note abundance of mesenchymal villi with sparse vascularity and primitive mesenchymal stroma (H&E, 4×)

Fig. 36.44  Placental tissue at 6 weeks gestation. Note abundance of mesenchymal villi with sparse vascularity and primitive mesenchymal stroma (H&E, 10×)

Fig. 36.45  Placental tissue at 6 weeks gestation. Close-up of mesenchymal villus showing an inner cytotrophoblast layer and an outer syncytiotrophoblast layer. Note the pink fuzzy outline of the syncytiotrophoblast layer, which represents the numerous microvilli that are readily identified in the earlier gestational age (also easily seen in Fig. 36.46) and are an important site of nutrient transport. Also note the primitive-appearing stroma with thin-walled vessels showing numerous nucleated erythrocytes in the fetal circulation at this gestational age (H&E, 20×)

Fig. 36.46  Chorionic villi at 14 weeks gestation. Immature intermediate villus is shown here with a thick double layer of trophoblast and reticular stroma with numerous macrophages. The stroma here is more cellular than in mesenchymal villi. Only occasional capillaries are seen within the villous stroma (H&E, 10×)

there are macroscopically visible deposits involving larger parts of the villous tree [23]. The fetal vessels of the placenta contain the elements of fetal blood. Nucleated red blood cells (NRBCs) in the villous circulation are seen commonly in the first trimester (see Fig.  36.45); there is a correlation between the number of nucleated and anucleate erythrocytes seen in the fetal circulation and the number of gestational days in the first trimes-

ter [24]. As gestation progresses, however, NRBCs are seen less frequently in the fetal circulation, and they are rare in the third trimester. Redline [25] correlated elevated NRBC counts (2.5 × 103/mm3) with placental findings and found that a threshold of 10 NRBC per 10 high-power fields (40×) in the fetal circulation of the placenta distinguished infants with elevated NRBC counts. Therefore, the finding of at least one definitive NRBC in most high-power fields of the villous parenchyma is a reasonable indicator of an increased NRBC count in the infant [25] and should be considered above the normal range.

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a

417

b

Fig. 36.47  Chorionic villus at 16 weeks gestation. (a) Note the continuous layer of syncytiotrophoblast highlighted by stain for human chorionic gonadotropin (HCG immunohistochemistry, 20×). (b)

Cytotrophoblast layer is discontinuous but still fairly prominent, as demonstrated by p57KIP2 staining (p57KIP2 immunohistochemistry, 40×)

Fig. 36.48  Chorionic villi at 16 weeks gestation. The vasculature of the villi is highlighted with CD31 immunohistochemical stain. The ratio of capillaries to stroma is quite low, and the capillaries are fairly centrally placed. The villous stroma appears cellular. This appearance is appropriate for an immature intermediate villus (CD31 immunohistochemistry, 20×)

Fig. 36.49  Chorionic villi at 21 weeks gestation. Note the formation of early stem villus (SV; right) and the presence of mature intermediate villi (arrows), with a more fibrous stroma than immature intermediate villi. Capillary formation does not occupy a large percentage of the stroma (H&E, 10×)

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Fig. 36.50  Chorionic villi at 21 weeks gestation. Mature intermediate villus with a more fibrous stroma than immature intermediate villi. Capillary formation does not occupy a large percentage of the stroma (H&E, 40×)

Fig. 36.51  Chorionic villi at 20  weeks gestation. CD31 stain highlights capillary formation occupying less than 50% of the stroma (CD31 immunohistochemistry, 20×)

Fig. 36.52  Chorionic villi at 23–24 weeks gestation. Low-power view of a stem villus (SV) surrounded predominantly by mature intermediate villi. Capillary formation is not prominent at this low power (H&E, 4×)

Fig. 36.53  Chorionic villi at 23–24  weeks gestation. Mostly mature intermediate villi are seen. Terminal villi are not a prominent feature in this image, although a few may be present at this gestational age (H&E, 10×)

Fig. 36.54  Chorionic villi at 23–24 weeks gestation. P57KIP2 is typically expressed in cytotrophoblast cells (arrows) and stromal cells of the chorionic villi. At this gestational age, p57KIP2 stain highlights the cytotrophoblast cells of mature intermediate villi, which occupy less than 50% of the villous surface (p57KIP2 immunohistochemistry, 20×)

Fig. 36.55  Chorionic villi at 23–24 weeks gestation. CD31 stain highlights villous vessels. Most villi have less than 50% stroma occupied by capillaries (CD31 immunohistochemistry, 10×)

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Fig. 36.56  Chorionic villi at 27 weeks gestation. Mature intermediate villi still predominate, but terminal villi (arrows), characterized as smaller villi with more than 50% capillaries, are seen occasionally (H&E, 10×)

Fig. 36.57  Chorionic villi at 27 weeks gestation. Mature intermediate villi with occasional smaller terminal villi (arrows). Vasculosyncytial membranes are just beginning to form (arrowhead). Perivillous fibrin is present but not excessive (H&E, 20×)

Fig. 36.58  Chorionic villi at 28  weeks gestation, with CD31 stain highlighting villous vessels. The villi in the center of the image have more than 50% of their stroma occupied by capillaries, a feature of terminal villi (CD31 immunohistochemistry, 10×)

Fig. 36.59  Chorionic villi at 28  weeks gestation, with p57KIP2 stain highlighting discontinuous villous cytotrophoblast cells (p57KIP2 immunohistochemistry, 20×)

Fig. 36.60  Chorionic villi at 32 weeks gestation. Villi are smaller and more vascular than at 27–28 weeks (compare with Fig. 36.56). Stem villus (SV) is present (right) (H&E, 10×)

Fig. 36.61  Chorionic villi at 32 weeks gestation. Note the presence of more terminal villi and vasculosyncytial membranes (arrows) than at earlier gestation (H&E, 20×)

420

Fig. 36.62  Chorionic villi at 32  weeks gestation, with CD31 stain highlighting villous vessels. Many villi have more than 50% stroma occupied by capillaries, a feature of terminal villi (CD31 immunohistochemistry, 20×)

L. M. Ernst and C. K. Carreon

Fig. 36.63  Chorionic villi at 32  weeks gestation, with p57KIP2 stain highlighting villous cytotrophoblast cells (p57KIP2 immunohistochemistry, 20×)

Fig. 36.64  Chorionic villi at 39  weeks gestation. Note the still-­ Fig. 36.65  Chorionic villi at 39 weeks gestation. Higher-power view increasing number of terminal villi and well-formed vasculosyncytial of villi highlighting vasculosyncytial membranes of terminal villi membranes (H&E, 20×) (H&E, 40×)

Fig. 36.66  Chorionic villi at 39  weeks gestation, with p57KIP2 stain highlighting villous cytotrophoblast cells, which are not very prominent at this gestational age (p57KIP2 immunohistochemistry, 20×)

Fig. 36.67  Chorionic villi at 39  weeks gestation, with CD31 stain highlighting villous vessels. Many villi have more than 50% stroma occupied by capillaries, a feature of terminal villi (CD31 immunohistochemistry, 10×)

36 Placenta Fig. 36.68  Diagram of vasculosyncytial membranes. The vasculosyncytial membrane, which becomes more noticeable in the early third trimester, is composed of syncytiotrophoblast cell cytoplasm, fused basement membrane of the syncytiotrophoblast cell and fetal endothelial cell, and fetal capillary endothelial cell cytoplasm. This thinning of the trophoblast layer, along with dilatation of fetal capillaries, results in a shorter diffusion distance between the fetal capillary lumen and the intervillous space, which in turn facilitates a more efficient gas and nutrient exchange between maternal and fetal blood

421 Syncytotrophoblast

Terminal villus

Fused basement membrane Fetal endothelial cell

Intervillous space

Special Considerations Multiple Gestations The microscopic anatomy of placentas of twin pregnancies and higher-order multiple births does not differ significantly from singleton placentas in the main basic compartments of the placenta. In fact, completely separate placentas are found in some multiple-gestation pregnancies. If there is an area of

Capillary lumen

connection between any two placentas of a multiple-­gestation pregnancy, however, histological sampling of this area is required to classify and document the type of placentation. Considering twin placentas for simplicity, there are three types: dichorionic diamnionic, monochorionic diamnionic, and monochorionic monoamnionic. The presence or absence of a dividing membrane, the portion of membranes that separates the two twin sacs, and the character of that membrane will define the type of twin placenta present.

422

L. M. Ernst and C. K. Carreon

Table 36.4  Comparison of the gross characteristics of twin dividing membranes Membrane thickness Membrane character Attachment to chorionic plate (T-zone) Chorionic vessels

Dichorionic Thick/four layers

Monochorionic Thin/two layers

Atrophied villi of chorion visible Well-attached, thickened ridge visible at attachment Stop at dividing membrane

Smooth and transparent, no atrophied villi Not well-attached, easily torn away, no ridge visible at attachment

a

Traverse beneath dividing membrane

Dichorionic diamnionic twin placentas can present as two completely separate placentas (no connection between the twin placentas; no dividing membrane present), or there can be a membranous connection between the twins (dividing membrane present on extraplacental membranes), or there can be fusion of the placental discs, with the presence of a dividing membrane on the chorionic plate. Monochorionic diamnionic twins essentially always present as a single placental disc, and the dividing membrane is typically found on the fetal surface. If no dividing membrane is seen, the examiner should suspect monochorionic monoamnionic placentation, but it is important to be aware that the dividing membrane of the monochorionic diamnionic twin is very delicate and is easily avulsed from the fetal surface before examination. Other data supportive of monoamnionic placentation should be sought, such as cord entanglement, short intercordal distance, or conjoined fetuses. The gross characteristics of the dividing membrane are described in Table  36.4 and Fig.  36.69. Histologically, the monochorionic diamnionic dividing membrane is characterized simply by two amnions in back-to-back orientation (Fig.  36.70). In comparison, the dichorionic diamnionic dividing membrane is characterized by the addition of chorionic tissue between the two back-to-back amnions (Figs. 36.71 and 36.72). In multiple-gestation pregnancies, chorionicity, the number of chorions (monochorionic vs. dichorionic), is not equivalent to zygosity, the number of zygotes (monozygous vs. dizygous). The number of zygotes and thus zygosity determines whether the offspring are identical or fraternal. Monozygosity can lead to either monochorionic or dichorionic placentation. Dizygosity leads only to dichorionic placentation. Therefore, from a histologic perspective, the identification of a monochorionic dividing membrane confirms monozygosity (with rare exceptions [26]), and the identification of a dichorionic dividing membrane can represent either identical or fraternal twins. Further genetic testing is needed to determine zygosity.

b

Fig. 36.69  Comparison of the gross characteristics of twin dividing membranes. (a) Dichorionic, diamnionic dividing membrane is characterized by a thick membrane composed of two amnions and two fused chorions. It is well fixed to the chorionic plate, and the chorionic vessels of each twin terminate at the dividing membrane. (b) Monochorionic, diamnionic dividing membrane is thinner, composed of only two amnions, and it is poorly fixed to the chorionic plate and is easily disrupted. Although not visualized well in this image, the chorionic vessels of the twins cross beneath the dividing membrane and may have intertwin vascular connections

Fetus Papyraceous In some multiple gestations, one of the fetuses may either be selectively reduced or may die spontaneously and be retained until the full term of the pregnancy’s gestation. Regardless of the mechanism of death, when a fetus dies in utero and is retained in utero, it will undergo involu-

36 Placenta

423

Fig. 36.70  Monochorionic dividing membrane from a twin placenta. The dividing membrane consists of two back-to-back amnion membranes (H&E, 20×)

Fig. 36.71  Dichorionic dividing membrane from a twin placenta. This is the typical appearance of the fused membranes, with back-to-back fused chorions (C). Note that there is no significant parietal decidua between the two chorions, and an atrophied villus of the chorion laeve is seen (arrow). The amnions of each twin are adjacent to the fused chorions, with their connective tissue facing the central fused chorions (H&E, 10×)

Fig. 36.72  Dichorionic dividing membrane from a twin placenta. Fused membranes with back-to-back chorions. In this figure, there are no visible atrophied villi of the chorion laeve, which are seen in a patchy fashion in dichorionic dividing membranes (H&E, 20×)

Fig. 36.73  Gross image of fetus papyraceous within the extraplacental membranes. Note the flattened and rounded appearance of the macerated fetal tissue. The fetal rib cage can be seen faintly and is a clue to the presence of a fetus papyraceous

Fig. 36.74  Microscopic appearance of fetus papyraceous. Note the macerated fetal tissues with significant loss of nuclear basophilia. Skeletal elements tend to retain nuclear detail the longest (H&E, 2×)

tional changes. With p­ rolonged periods of retention in utero, the fetal tissue becomes atrophied and flattened; this is known as a fetus papyraceous. It is usually recognized as a somewhat thickened, ovoid area on the extraplacental membranes. Features that distinguish fetus papyraceous from thickened decidua or mere decidual necrosis include the presence of a small speck of retinal pigment in the region of the eye or the presence of parallel-oriented ribs of the thoracic cage (Fig.  36.73). If the fetus papyraceous is examined histologically, it shows variable degrees of loss of nuclear basophilia in fetal tissue; with prolonged retention, nearly all the visceral organs show loss of nuclear basophilia. Skeletal elements, such as cartilage and bone, are the easiest to recognize because of longer retention of nuclear detail in those tissues (Fig. 36.74).

424

Fig. 36.75  Gross image of yolk sac remnant. A small, ovoid, yellow-­ white structure (arrow) is noted on the chorionic plate

L. M. Ernst and C. K. Carreon

Fig. 36.76  Calcified yolk sac remnant. Low-power image of the yolk sac remnant located between the amnion and chorion of the membranes (H&E, 4×)

Yolk Sac Remnant Not infrequently, a small, ovoid, flattened calcification can be noted between the chorion and amnion, either in the extraplacental membranes or on the chorionic plate (Fig. 36.75). This represents the calcified remnant of the yolk sac, which atrophies in the embryonic period. Histological sections usually show an ovoid calcification without other epithelial elements, located between the chorion and the amnion (Fig. 36.76).

References 1. Sadler TW, Langman J.  Langman’s medical embryology. 11th ed. Philadelphia: Wolters Kluwer Lippincott Williams & Wilkins; 2010. 2. Schoenwolf GC, Larsen WJ. Larsen’s human embryology. 4th ed. Philadelphia: Elsevier/Churchill Livingstone; 2009. 3. Benirschke K, Burton GJ, Baergen RN. Anatomy and pathology of the placental membranes. In: Pathology of the human placenta. 6th ed. New York: Springer; 2012. p. 249–307. 4. Benirschke K.  Remarkable placenta. Clin Anat. 1998;11: 194–205. 5. Shih I-M, Mazur MT, Kurman RJ. Gestational trophoblastic tumors and related tumor-like lesions. In: Kurman RJ, Ellenson LH, Ronnett BM, editors. Blaustein’s pathology of the female genital tract. 6th ed. New York: Springer; 2011. p. 1075–135. 6. Redline RW, Boyd T, Campbell V, Hyde S, Kaplan C, Khong TY, Society for Pediatric Pathology, Perinatal Section, Maternal Vascular Perfusion Nosology Committee, et al. Maternal vascular underperfusion: nosology and reproducibility of placental reaction patterns. Pediatr Dev Pathol. 2004;7:237–49. 7. Khong TY, Mooney EE, Ariel I, Balmus NC, Boyd TK, Brundler MA, et al. Sampling and definitions of placental lesions: Amsterdam placental workshop group consensus statement. Arch Pathol Lab Med. 2016;140:698–713. 8. Benirschke K, Burton G, Baergen RN. Anatomy and pathology of the umbilical cord. In: Pathology of the human placenta. 6th ed. New York: Springer; 2012. p. 309–75. 9. Stehbens WE, Wakefield JS, Gilbert-Barness E, Zuccollo JM.  Histopathology and ultrastructure of human umbilical blood vessels. Fetal Pediatr Pathol. 2005;24:297–315. 10. Drut R, Quijano G.  Heterotopic neurons in the umbilical cord. Pediatr Dev Pathol. 2005;8:124–7.

11. Ellison JP. The nerves of the umbilical cord in man and the rat. Am J Anat. 1971;132:53–60. 12. Preminger A, Udassin R, Pappo O, Arad I.  Ectopic liver tis sue within the umbilical cord. J Pediatr Surg. 2001;36: 1085–6. 13. Redline RW, Ariel I, Baergen RN, Desa DJ, Kraus FT, Roberts DJ, Sander CM.  Fetal vascular obstructive lesions: nosology and reproducibility of placental reaction patterns. Pediatr Dev Pathol. 2004;7:443–52. 14. Benirschke K, Burton G, Baergen RN.  Nonvillous parts and trophoblast invasion. In: Pathology of the human placenta. 6th ed. New York: Springer; 2012. p. 157–240. 15. Salafia CM. The normal placenta immune anatomy and function. Immunol Allergy Clin N Am. 1998;18:271–89. 16. Khong TY, Sawyer IH. The human placental bed in health and disease. Reprod Fertil Dev. 1991;3:373–7. 17. Khong TY. The Robertson-Brosens-Dixon hypothesis: evidence for the role of haemochorial placentation in pregnancy success. Br J Obstet Gynaecol. 1991;98:1195–9. 18. Jauniaux E, Gulbis B, Burton GJ. The human first trimester gestational sac limits rather than facilitates oxygen transfer to the foetus – a review. Placenta. 2003;24(Suppl A):S86–93. 19. Parks WT.  Manifestations of hypoxia in the second and third trimester placenta. Birth Defects Res. 2017;109:1345–57. 20. Benirschke K, Burton G, Baergen RN.  Characterization of the developmental stages. In: Pathology of the human placenta. 6th ed. New York: Springer; 2012. p. 145–55. 21. Benirschke K, Burton G, Baergen RN. Architecture of normal villous trees. In: Pathology of the human placenta. 6th ed. New York: Springer; 2012. p. 101–44. 22. Loukeris K, Sela R, Baergen RN. Syncytial knots as a reflection of placental maturity: reference values for 20 to 40 weeks’ gestational age. Pediatr Dev Pathol. 2010;13:305–9. 23. Benirschke K, Burton G, Baergen RN.  Histologic approach to villous alterations. In: Pathology of the human placenta. 6th ed. New York: Springer; 2012. p. 395–410. 24. Salafia CM, Weigl CA, Foye GJ.  Correlation of placental erythrocyte morphology and gestational age. Pediatr Pathol. 1988;8: 495–502. 25. Redline RW.  Elevated circulating fetal nucleated red blood cells and placental pathology in term infants who develop cerebral palsy. Hum Pathol. 2008;39:1378–84. 26. Souter VL, Kapur RP, Nyholt DR, Skogerboe K, Myerson D, Ton CC, et al. A report of dizygous monochorionic twins. N Engl J Med. 2003;349:154–8.

Part XI Special Considerations for Fetal Tissues

Histological Changes Associated with Fetal Maceration

37

Linda M. Ernst and Michael K. Fritsch

Introduction The word “macerate” used as an intransitive verb is defined as “to soften and wear away especially as a result of being wetted or steeped” [1]. The collective changes that occur in the fetal organs and tissues after fetal death in utero are referred to as maceration changes, which differ from pure postmortem autolysis because they occur in utero, in fluid, and at body temperature. The changes of maceration are therefore commonly encountered by the pathologist performing fetal autopsies and examining fetal tissues at the surgical pathology bench. Maceration changes in fetal tissue can produce artifacts and loss of nuclear detail that can limit interpretation of pathology. Fortunately, the changes of maceration are somewhat predictable and once recognized can be used to help estimate the timing of in utero death.

Histology of Maceration  iming of Intrauterine Fetal Death Based T on Histology of Fetal Organs Retention in utero after fetal death leads to both gross and histologic changes in the fetal tissues. Gross changes in the fetal appearance include desquamation of the skin, red to brown discoloration of the skin, discolored sclerae, overlapping skull bones, unfused eyelids, sunken eyes, generalized loss of tissue turgor with joint laxity, serosanguineous effusions, dark red discoloration of the umbilical cord, softening

L. M. Ernst (*) Department of Pathology and Laboratory Medicine, NorthShore University HealthSystem, Evanston, IL, USA e-mail: [email protected] M. K. Fritsch Department of Pathology and Laboratory Medicine, University of Wisconsin, Madison, Madison, WI, USA

and discoloration of the fetal organs, and ultimately mummification [2]. Brain and spinal cord tissue can become extremely soft and liquefied over time, but the nuclear detail can persist even in extremely macerated brain tissue. With prolonged retention in utero, the fetus loses mass and becomes compressed and flattened, with the end stage known as fetus papyraceous. Nevertheless, it can be difficult to rely on gross features alone to assess the timing of intrauterine fetal death. At least one study has shown that in low-income countries, provider descriptions of the fetal appearance at birth may be unreliable and lead to inaccurate estimates of the timing of fetal death [3]. Histologic features of macerated fetal tissue can also provide additional information to assist in the process of estimating when fetal death occurred. A classic study of the timing of maceration changes based on histologic features of the fetal organs by Genest et al. [4] provided a framework to approach estimating the timing of fetal death (death-to-­delivery interval) using histologic features. The universal feature of maceration seen in the fetal tissues is loss of nuclear basophilia, which is characterized by a fading of the nuclear detail in an organ. An organ with maximal loss of nuclear basophilia has no nuclei visible, and the tissue appears diffusely eosinophilic. Because each fetal organ involutes differently in the postmortem in utero setting, some organs are better predictors of timing than others (Table 37.1) [4]. In our experience, the organs that are the most helpful in estimating the timing of fetal death are those that present with maximal loss of nuclear basophilia and are good predictors of timing: the myocardium (Fig. 37.1), the liver (Figs. 37.2 and 37.3), the adrenal gland (Figs. 37.4 and 37.5), and the kidney (Fig. 37.6). There is no fetal organ that provides a good estimate of the timing of fetal death between 2 and 4 weeks, but including the placenta in the assessment can add some information for this time period, as discussed below. The timing of onset of these histologic changes in the fetal organs can be modified by other factors such as prolonged maternal fever, chronic fetal hypoxia, termination

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1_37

427

428

L. M. Ernst and M. K. Fritsch

Table 37.1  Fetal histologic features: good predictors of death-to-­ of pregnancy by potassium chloride injection, direct delivery interval

Organ Kidney Liver Myocardium Myocardium

Bronchus Liver Adrenal gland Trachea Gastrointestinal tract Kidney

Time lapse between fetal death and Loss of nuclear basophilia delivery Tubules ≥4 h Hepatocytes ≥24 h Inner half of the ≥24 h myocardium Inner and outer half of ≥48 h the myocardium (maximal) Epithelium ≥96 h Maximal ≥96 h Maximal ≥1 week Chondrocytes ≥1 week Maximal ≥1 week Maximal

i­schemic injury to an organ due to thrombus (e.g., renal vein thrombosis), and even delay in performance of the autopsy, with additional autolysis. On occasion, the histologic findings of loss of nuclear basophilia in various organs may be discordant and therefore may limit the accurate assessment of the duration of intrauterine fetal demise. The “gold standard” for estimating the duration of intrauterine fetal demise is an accurate clinical history. For instance, if there is clear clinical evidence of a live fetus by ultrasound 24 h prior to delivery but the histologic findings suggest a longer duration, the history should be considered more accurate.

≥4 weeks

Adapted from Genest et al. [4]; with permission

a

Fig. 37.2  Liver with moderate loss of nuclear basophilia. Note focal preservation of the nuclear detail in the extramedullary hematopoiesis and hepatocytes. Fetal death was estimated to have occurred 2–4 days prior to delivery, because this fetus also had maximal loss of nuclear basophilia in the heart (H&E, 20×)

b

Fig. 37.1  Loss of nuclear basophilia in the heart. Myocardium showing maximal loss of nuclear basophilia in the inner half (a) and moderate but submaximal loss of nuclear basophilia in the outer half (b). Fetal death was estimated to have occurred 1–2 days prior to delivery (H&E, 40×)

Fig. 37.3  Liver with maximal loss of nuclear basophilia. Note that extramedullary hematopoiesis is no longer detected, and hepatocytes are not definitely recognizable. Fetal death was estimated to have occurred at least 4 days prior to delivery (H&E, 20×)

37  Histological Changes Associated with Fetal Maceration

429

Fig. 37.4  Adrenal gland with mild to moderate loss of nuclear basophilia. Note focal preservation of the nuclear detail in the definitive and fetal cortices, although there is loss of tissue integrity and architecture in the definitive cortex. This fetus had only mild loss of nuclear basophilia in other organs, and fetal death was estimated to have occurred less than 2 days prior to delivery (H&E, 10×)

Fig. 37.5  Adrenal gland with maximal loss of nuclear basophilia. Cortical nuclei are no longer visible, but fatty change in the fetal cortical cells can be seen. Fetal death was estimated to have occurred at least 1 week prior to delivery (H&E, 20×)

Fig. 37.6  Kidney with moderate loss of nuclear basophilia. The tubular epithelium and glomeruli have lost their nuclear detail, but the renal architecture can still be detected (H&E, 10×)

Fig. 37.7  Chorionic villi showing karyorrhexis of the villous capillaries (arrows). This is an early change in the placenta, seen 6 h or more after fetal death (H&E, 40×)

 iming of Intrauterine Fetal Death Based T on Histology of the Placenta

est change of fetal vascular involution is karyorrhexis in the fetal capillaries (Fig. 37.7). This change is characterized by karyorrhectic debris within the villi in the areas where villous capillaries normally are located. The capillary lumen may or may not still be discernible. With more prolonged retention in utero after fetal death, the stem villous vessels and chorionic vessels become involved. These muscularized vessels may show fibrous ingrowth of tissue, which leads to obliteration of the vessel lumen (Figs.  37.8 and 37.9). Endothelial cells are no longer visible, and luminal blood may be absent. This obliterative change may also appear as fibrous septations within the vessel lumen, with several

The fetal vasculature of the placenta, including the umbilical vessels, chorionic vessels, stem villous vessels, and capillaries, undergoes involutional changes after fetal death with the halt of the continued circulation of the fetal blood. Since maternal perfusion to the placenta persists after fetal death until delivery, histologic involutional and maceration changes in the placenta are recognized in the fetal vasculature and include changes in the surface (chorionic) vessels, fetal stem villous vessels, and villous capillaries. The earli-

430

Fig. 37.8 Stem vessel involutional changes. This large stem villous vessel shows luminal obliteration. The arrows indicate the margins of the vessel lumen. Note the somewhat myxoid-appearing fibrous tissue that has grown into the lumen and caused its near-total obliteration. A few degenerated red blood cells persist in the center of the lumen. Endothelial cells are not seen (H&E, 20×)

small, irregular luminal spaces remaining that contain degenerating blood cells, resembling recanalized thrombi [5]. It is important to note that true fibrin thrombi within fetal vessels are not a feature of involutional change in the placenta. Ultimately, after prolonged retention in utero, the fetal vasculature is completely obliterated, and chorionic villi appear avascular, with remaining fibrotic stroma (Fig. 37.10). Because maternal perfusion persists, the trophoblast layer is preserved, as is the intervillous space. In some cases, such as early demise of a twin with extremely prolonged retention of the twin until term, the chorionic villi become collapsed upon each other and appear “infarcted,” but it is important not to characterize this finding necessarily as pathologic maternal malperfusion. The changes in the fetal vasculature after fetal death are diffuse and advance over time (Table 37.2), generally affecting the villous tree globally, with no significant regional variability in the appearance of the villi or the fetal vascular changes. Regional villous findings such as villous stromal-­ vascular karyorrhexis or groups of avascular villi most likely represent evidence of premortem fetal vascular malperfusion [6]. Because there is significant overlap between the changes of fetal vascular involution after fetal death and the changes of antemortem fetal vascular malperfusion, it can be difficult to separate true antemortem fetal vascular obstructive changes from those related to postmortem involution. Once global loss of villous capillaries is present, it is extremely difficult, if not impossible, to definitively diagnose fetal vascular malperfusion. Criteria have been established to assist the pathologist with diagnosis of “cord accidents” in stillbirth [7, 8].

L. M. Ernst and M. K. Fritsch

Fig. 37.9  Stem vessel involutional changes. This stem villous shows several profiles of stem villous vessels. The lumens are not completely obliterated, but multiple septations are present, dividing the large lumen into small lumens, mimicking recanalized thrombi (H&E, 20×)

Fig. 37.10  Low-power view of chorionic villi in a case where fetal death was estimated to have occurred 1–2 weeks prior to delivery. Note that many of the villi have complete stromal fibrosis, and only a few show remaining capillaries. The intervillous space is intact and not collapsed (H&E, 10×) Table 37.2  Placental histologic features: good predictors of death-to-­ delivery interval Placental compartment Villous capillaries Stem villous vessels Stem villous vessels Chorionic villi

Histologic feature Intravascular karyorrhexis Multifocal stem vessel luminal abnormalities (10–25%)a Extensive stem vessel luminal abnormalities (>25%)a Extensive villous fibrosis

Time lapse between fetal death and delivery ≥6 h ≥48 h ≥2 weeks ≥2 weeks

Adapted from Genest [5]; with permission a Luminal abnormalities = luminal obliteration or septation

37  Histological Changes Associated with Fetal Maceration

Special Topics Tissue Artifacts Associated with Maceration Because maceration is a unique form of autolysis, unusual tissue artifacts can be seen with maceration. Many of these artifacts do not have specific names, but they can be seen with some regularity in the tissues of macerated fetuses. These include findings such as “blue blobs” in the liver and thymus (Fig. 37.11), “mulberries” (Fig. 37.12), “basophilic dust” surrounding viscera (Fig.  37.13), and “yellow starbursts” (Fig. 37.14). These artifacts should not be confused with calcifications, bacteria, viral inclusions, hemosiderin deposits, or formalin pigment (Fig. 37.15). Brain tissue can become liquefied with prolonged retention in utero, and under pressure (such as vaginal delivery) it can be pushed out the intervertebral foramina into the retroperitoneal space. This finding should not be confused with a neuroblastic tumor in the retroperitoneum. Brain tissue can also be seen in the vasculature of the liver and other organs. If there is uncertainty, immunohistochemical stain for glial fibrillary acidic protein or CD56 can be helpful.

L imitations on Interpretation of Histologic Findings with Maceration Though the loss of nuclear basophilia can be helpful in determining the timing of fetal death, it can lead to problems interpreting the origin of some tissues. Often, however, the outline of the tissue organization does persist and some pathology such as viral inclusions, calcifications, and even some inflammation may persist.

Fig. 37.11  “Blue blobs” in the liver. These rounded blue structures, commonly seen in the macerated liver, are an artifact. They may represent collections of nuclear material (H&E, 40×)

431

Because maceration changes involve the loss of nuclear basophilia, certain fetal pathology may be difficult or impossible to interpret in the setting of maceration. The more severe the maceration, the more likely it is to hinder pathologic diagnosis, but we do recommend a thorough sampling and examination of fetal tissue at fetal autopsy because, despite the difficulties, certain pathologies such as viral inclusions (cytomegalovirus, herpes virus, and parvovirus) (Fig.  37.16), inflammation, true fetal vasculature thrombi, calcification (Fig. 37.17), and structural abnormalities may still be discernible, even with severe maceration. The use of connective tissue special stains such as Masson trichrome can often at least define overall normal organ architecture and assess for increased fibrosis. Table 37.3 indicates some of the difficulties in the interpretation of pathology with maceration.

Intracardiac Potassium Chloride (KCl) There is little literature documenting the effects on the histology of various organs of intracardiac KCl injection for termination of pregnancy, but those pathologists that perform autopsies are well aware of these histologic findings. KCl solutions used for intracardiac injection to induce fetal demise are extremely hypertonic, and the effects on fetal tissues mimic maceration changes, especially in the heart. Grossly, the endocardium and septum are usually friable; histologically, there is complete loss of nuclear basophilia. KCl injection may not be applied directly into the heart, and other sites commonly affected include the abdomen, liver, and pleural spaces or lung. The result can be accelerated tissue destruction in the affected organs, with advanced loss of nuclear basophilia [9].

Fig. 37.12  “Mulberries” are a common artifact seen in many macerated tissues. Their origin is unclear (H&E, 60×)

432

Fig. 37.13  “Basophilic dust” is a common artifact seen in sections of macerated fetal organs. In this case, the basophilic dust is seen in the soft tissues surrounding the pelvic organs in a macerated term fetus. The basophilic dust should not be confused with bacteria; a tissue Gram stain can help to clarify, if needed (H&E, 4×)

Fig. 37.15  Brown pigment in fatty tissue is a frequent finding in macerated fetal soft tissue. Its origin is unclear, but it should not be misinterpreted as hemosiderin deposition (H&E, 40×)

L. M. Ernst and M. K. Fritsch

Fig. 37.14  “Yellow starbursts” are a common artifact seen in many macerated tissues. Their origin is unclear (H&E, 60×)

Fig. 37.16  Pancreatic tissue with near maximal loss of nuclear basophilia; a distinctive cytomegalovirus viral inclusion is easily recognized (H&E 40×)

37  Histological Changes Associated with Fetal Maceration

Fig. 37.17  Adrenal gland with maximal loss of nuclear basophilia; calcification is evident in the fetal cortex (H&E 4×)

433

Fig. 37.18  Thymus gland with moderate loss of nuclear basophilia and fading of the distinction between cortex and medulla. Note that it can be difficult to distinguish loss of nuclear basophilia (maceration) from depletion of cortical lymphocytes (involutional change related to fetal stress) (H&E, 10×)

Table 37.3  Interpretation of histologic findings with maceration Organ Thymus

Difficult to interpret with maceration Involutional changes

Liver

Bile duct development

Liver

Extramedullary hematopoiesis (EMH)

Liver

General structure

Kidney

Proximal tubule assessment

Pancreas

General structure

Bone marrow

Myeloid-­erythroid ratio

Lung

General structure

GI tract

Mucosa

GI gastrointestinal, PAS periodic acid–Schiff

Comments Involutional changes of the thymus involve loss of cortical lymphocytes, which leads to loss of nuclear basophilia in the cortex (Fig. 37.18) and can overlap with loss of nuclear basophilia from maceration Biliary epithelium is lost quickly with maceration, so assessment of bile duct development could be hindered With severe maceration, nuclei of hepatocytes and hematopoietic cells are lost. With less severe maceration change, EMH may be assessed Trichrome can be helpful to determine if the structure of the liver is normal or abnormal with severe maceration changes In most cases, the proximal tubules are not well preserved, but their presence can usually be detected until maceration is severe Usually significant loss of nuclear basophilia limits evaluation of acinar and islet tissue With moderate to severe loss of nuclear basophilia, it can be difficult to assess the origin of hematopoietic precursors. PAS stain is helpful in some cases Pulmonary epithelium sloughing and loss of nuclear basophilia can preclude assessment of epithelial features, but determination of the stage of pulmonary development can usually be made based on overall architecture (Fig. 37.19), which usually persists The GI mucosa is often sloughed and difficult to assess, but characteristics of the wall such as the presence of muscularis mucosa, muscularis propria, and ganglionation can often be determined

434

Fig. 37.19  Lung tissue with mild maceration changes. Note the loss of the epithelial lining of the airspaces. The general structure of the lung is still discernible; the formation of secondary crests is visible, consistent with the saccular stage of development (H&E, 10×)

References 1. “Macerate.” Merriam-Webster, www.merriam-webster.com/dictionary/macerate. Accessed 25 Apr 2018. 2. Genest DR, Singer DB.  Estimating the time of death in stillborn fetuses: III.  External fetal examination; a study of 86 stillborns. Obstet Gynecol. 1992;80:593–600.

L. M. Ernst and M. K. Fritsch 3. Gold KJ, Abdul-Mumin AR, Boggs ME, Opare-Addo HS, Lieberman RW.  Assessment of “fresh” versus “macerated” as ­accurate markers of time since intrauterine fetal demise in lowincome countries. Int J Gynaecol Obstet. 2014;125:223–7. 4. Genest DR, Williams MA, Greene MF.  Estimating the time of death in stillborn fetuses: I. Histologic evaluation of fetal organs; an autopsy study of 150 stillborns. Obstet Gynecol. 1992;80:575–84. 5. Genest DR.  Estimating the time of death in stillborn fetuses: II.  Histologic evaluation of the placenta; a study of 71 stillborns. Obstet Gynecol. 1992;80:585–92. 6. Khong TY, Mooney EE, Ariel I, Balmus NC, Boyd TK, Brundler MA, et al. Sampling and definitions of placental lesions: Amsterdam placental workshop group consensus statement. Arch Pathol Lab Med. 2016;140:698–713. 7. Parast MM, Crum CP, Boyd TK.  Placental histologic criteria for umbilical blood flow restriction in unexplained stillbirth. Hum Pathol. 2008;39:948–53. 8. Ryan WD, Trivedi N, Benirschke K, Lacoursiere DY, Parast MM.  Placental histologic criteria for diagnosis of cord accident: sensitivity and specificity. Pediatr Dev Pathol. 2012;15:275–80. 9. Royal College of Obstetricians and Gynaecologists and Royal College of Pathologists. Fetal and perinatal pathology: report of a joint working party. London: Royal College of Obstetricians and Gynaecologists; 2001. p. 1–38.

Index

A Abnormal tubular differentiation, 117 Accessory spleens, 266 Acidophils, 243–245 Acinotubular stage, 169 Acute thymic involution, 258 Adenohypophysis, 241 Adenoid change, 220 Adnexa apocrine gland, 383 eccrine gland, 382, 383 hair follicle, 381, 382 hypodermis, 383, 384 nail, 383 sebaceous glands, 382 Adrenal corticotropin hormone (ACTH), 242–244 Adrenal cytomegaly, 219, 220 Adrenal gland 12 weeks gestation, 214, 216, 217 17 month old child, 217 17 weeks gestation, 215, 218 20 weeks gestation, 215, 216, 218 26 weeks gestation, 215 35 weeks gestation, 215, 216 39 weeks gestation, 214, 216, 218, 219 adenoid change, 220 adrenal cytomegaly, 219, 220 calcifications, 220, 221 capsule, 214 CD31 stain, 217 cortex definitive zone, 214, 215 fetal zone, 215 transitional zone, 216 embryology, 213, 214 extramedullary hematopoiesis, 221 fatty change in adrenal cortex, 221 fetal cortex/zone involution, 217, 218 heterotopic adrenal tissue, 219 layers, 214 medulla, 216–219 Adrenal weight, 213 Adventitia, 126, 193, 194 Allantoic remnant, 407, 409 Alveolar stage embryology, 36 histology, 42, 44 Amnioblasts, 400 Ampulla, of vas deferens, 158, 159 Ampullary cristae, 334 Anal canal, 51, 53, 67, 74 Androgen insensitivity syndrome, 167

Angioblast, 21 Angiogenesis, 21 Anitschkow cells, 6, 7, 9 Apocrine gland, 383 Apocrine metaplasia, 389, 394 Appendix less than 14 weeks, 64, 70 14–30 weeks, 65, 71–73 30–40 weeks, 69, 74, 75 epididymis, 143 cystic, sessile, 143 solid, pedunclated, 143 surface epithelium, 144 testis cystic, 143 epithelium, 143 solid filiform, 142 Atrioventricular (AV) valve, 11–13 Atypical giant cells, 394 B Bag fibers, 373 Bartholin glands, 204–206 Basal cells, 161, 163, 164 Basal plate, 408, 410–413 Basophil cells, 243–245 Basophilic dust, 432 Basophilic dye (basophils), 243 Beckwith-Weidemann syndrome, 220 Bessis islands, 79 Biliary tract extrahepatic biliary tree, 85–89 intrahepatic biliary tree, 81, 84, 85 Blood flow patterns, 22 Blood islands, 281 B-lymphocyte, 255, 266 Body weight, 213 Bone bone formation endochondral ossification, 362–365 membranous ossification, 365 cell types osteoblasts, 361, 364 osteoclasts, 362, 364 osteocytes, 361 osteoprogenitor cells, 361 histological types lamellar bone, 359, 361 woven bone, 360, 361 lateral plate mesoderm, 357 long-bone ossification, 358, 359

© Springer Nature Switzerland AG 2019 L. M. Ernst et al. (eds.), Color Atlas of Human Fetal and Neonatal Histology, https://doi.org/10.1007/978-3-030-11425-1

435

436 Bone (cont.) lower limb 38-week fetus, 359 40-week fetus, 359 metacarpal bone ossification, 358 neural crest cells, 358 paraxial mesoderm, 357 somites, 358 structure cortical bone, 360, 362 medullary canal, 360, 363 periosteum, 360, 362 vascular supply, 360, 363 Bone marrow 19 weeks gestation, 283 24 weeks gestation, 283 32 weeks gestation, 283 41 weeks gestation, 283 embryology, 281, 282 during fetal life, 282 hematopoiesis, 284 hemophagocytosis, 287 myelopoiesis erythropoiesis, 285 granulopoiesis, 284 lymphopoiesis, 286, 287 megakaryopoiesis, 286 monopoiesis, 284 structure, 283 Bone morphogenic proteins (BMPs), 377 Bony labyrinth, 332 Bovine parathyroid hormone, 233–234 Bowman’s layer (BL), 312, 313 Brainstem, 299, 300 Branching morphogenesis, 167 Bronchus-associated lymphoid tissue (BALT), 37, 45, 46 Brown pigment, 432 Buck's fascia, 200, 201 Bud stage, 168 Bud-tubule stage, 169 Bulbourethral (Cowper) glands, 203 Bulbourethral glands, 161, 167 Bulbous urethra, 203 C Calcifications, 220, 221 Calcitonin, 223, 226, 229–231 Canalicular stage early, 39, 41 embryology, 36 late, 40, 42 Cardiac conduction system AV node, 9, 10, 15, 18 bundle of His, 9, 11–13, 16–18 components, 4 fibrous annulus, 5, 11, 16 incisions, 13 method of removal, 12 SA node, 10, 14, 18 Cardiac myocytes, 7 Caudal pouch, 233 C cells, 226, 229–231 Central nervous system cytology astrocytic development, 294 choroid plexus, 291, 293 meninges, 297, 298

Index microglia, 297, 298 myelination-associated glia, 295, 296 neuroepithelium, 291, 292 oligodendrocytes, 295, 296 radial glia, 291, 293 unmyelinated tracts, 295, 296 developmental anatomy and histology brainstem, 299, 300 cerebellum, 299, 301, 302 cerebral hemispheres, 305 cerebral neocortex, 304, 307 cortical mantle development, 302, 303, 306 external cerebral development, 302, 303 hippocampal development, 304, 308 midbrain, 302 spinal cord, 297, 299 sulcal development, 302, 304 embryology, 291 Papillae of Retzius, 304, 305, 309 Centroacinar cells, 93 Cerebellum, 299, 302 Chain fibers, 373 Chicken ovalbumin upstream promoter transcription factor 2 (COUP-TFII), 22 Chordae tendineae, 4, 8, 11 Chorionic plate, 407–410 Chorionic-type intermediate trophoblast cells, 404 Chorionic villi 14 weeks gestation, 416 16 weeks gestation, 417 20 weeks gestation, 418 21 weeks gestation, 417, 418 23 to 24 weeks gestation, 418 27 weeks gestation, 419 28 weeks gestation, 419 32 weeks gestation, 419, 420 39 weeks gestation, 420 gestational age, 415 villous types and characteristics, 415 Chorion laeve, 401–403, 407, 423 Choroid, 314, 316 Chromaffin cells, 217–219 Chromatin, 233 Chromophobe, 243–245 Cilia, 183, 184 Ciliary body, 314–316 Ciliary epithelium, 314 Ciliary processes, 316 Clitoral glans, 198, 205–208 Clitoral hood, 198 Clitoris, 198, 204, 206, 207 Cochlear canaliculus, 333 Colloid, 223, 225–228, 230 Columnar cell, 161–163 Common excretory duct, 113 Complex migrational pathways, 295 Congenital blood cyst, 9, 13 Congenital erosion, 192 Coni vasculosi, 148, 150 Connexins, 7 Cords of Billroth, 266 Cornea embryology, 312 histology, 312 Corneal epithelium, 312, 313 Corneal stroma, 312 Coronal sulcus, 199, 200

Index Corpora cavernosa, 198, 200, 202, 204, 206 Corpus spongiosum, 197–202, 204, 206, 207 Cowper glands, 167, 203, 204 Crura, 198, 202, 206–208 Cryptorchidism, 147 Crystallins, 319 Cumulus oophorus, 179, 180 Cupola, 346, 347 Cystic appendix testis, 143 Cystic hygroma, 29 Cystic, sessile appendix epididymis, 143 Cytomegaly, 219, 220 D Dartos fascia, 200, 204 Decidua spp. D. capsularis, 404 D. parietalis, 404 Deep dorsal vein, 198, 200, 202 Definitive cortex, 214, 216, 217, 219 Definitive zone, 214 Dehydroepiandrosterone (DHEA), 213 Dermis, 380, 381 Dermomyotome, 367 Descemet’s membrane (DM), 312 Dichorionic diamnionic twin placentas, 422, 423 Diffuse gut-associated lymphoid tissue, 52, 59, 60 Dihydrotestosterone (DHT), 197 Distal penis, 198–200 Dorsal trunk dermis, 378 Ducts, 93 Ductus arteriosus, 24, 25 Ductus deferens, see Vas deferens Ductus epididymis, 147–149, 151 Ductus reuniens, 328, 329, 333–335 Ductus venosus, 27, 78 Dyserythropoiesis/erythroid dysplasia, 285, 286 E Ear anatomy arcuate eminence, 325 external auditory canal, 329 inner ear, 326, 332–334 inner, middle, and external auditory canal, 327 internal acoustic meatus, 326 jugular fossa, 326 middle ear, 327, 331, 332 ostium, 326 petrous bone, 325, 326 subarcuate fossa, 325 surface landmarks, 325 tympanic membrane, 329 embryology bony labyrinth or otic capsule, 336 cecum, 335 ductus reuniens, 335 Meckel’s cartilage, 335 otic placode, 334, 335 Reichert’s cartilage, 335 tubotympanic recess, 335 utriculosaccular duct, 335 histology Eustachian tube, 343 external auditory canal, 336

437 inner ear, 343–350 middle ear, 339 tympanic membrane, 337 perinatal period, 350, 351 temporal bone, 352, 353 Eccrine gland, 382, 383 Ectocervix, 190, 192 Ectodermal ring, 385 Ectopic adrenal nodules, 145 Efferent ductules, 147–150 Ejaculatory ducts, 169 Embryonic epithelial remnants, 145, 146, 243 Embryonic stage embryology, 35 histology, 39 Embryonic vessels, 22 Endocardium, 8, 10 Endocervix, 187 Endochondral ossification, 362–365 Endocrine cell nucleomegaly, 96 Endocrine pancreas, 93–95 Endolymphatic duct, 349 Endometrium, 188–190 21 weeks, 189 28 weeks, 189 35 weeks, 190 Endomitosis, 286 Endomysium, 373 Epicardium, 6, 7 Epidermis 10 weeks gestation, 378 13 weeks gestation, 378, 379 23 weeks gestation, 379, 380 30 weeks gestation, 379 37 weeks gestation, 379, 380 Epididymis 17 weeks gestation anatomy, 149 body of, 149 efferent ductules, 149 tail of, 150 22 weeks gestation anatomy, 150 body, 151 coni vasculosi, 150 tail, 151 34 weeks gestation anatomy, 152 body, 153 efferent ductules, 152 head, 152 tail, 153 39 weeks gestation body, 154 efferent ductules, 154 head, 153 tail, 154, 155 body of, 149 congenital anomalies, 147 embryology, 147 histology, 148 primary function, 147 tail of, 149 Epimysium, 371 Epithelial elements, 168, 170 Epoophoron, 180, 181 ER71 gene, 22 Erythropoiesis, 285

438 Esophagus 14–30 weeks, 54–57 30–40 weeks, 55–58 less than 14 weeks, 54, 55 Estrogen, 198, 205 ETS translocation variant 2 (ETV2), 22 Exocervix, 190 Exocrine pancreas, 92, 93 External auditory canal, 329 External spermatic fascia, 204 Extra-adrenal chromaffin cells, 30 Extraembryonic mesoderm, 400 Extraembryonic somatopleuric mesoderm, 400–401 Extramedullary hematopoiesis (EMH), 15, 221 Extraplacental membranes, 402–404 Eye cornea embryology, 312 histology, 312 lens embryology, 319 histology, 320, 321 optic nerve embryology, 316–318 histology, 318 retina embryology, 316–319 histology, 318–321 sclera embryology, 312 histology, 313 uvea embryology, 313–315 histology, 315 vitreous embryology, 319 histology, 321, 322 Eye-field transcription factors (EFTFs), 316 F Fallopian tube 19 weeks, 184 21 weeks, 184, 185 23 weeks, 184–186 32 weeks, 185 38 weeks, 185, 186 first segment, 183 second segment, 183 third segment, 183 Fatty change, adrenal cortex, 221 Female external genitalia clitoris labia majora, 208 labia minora, 207 vulvar vestibule, 204, 205 Fetal Leydig cells 10–20 weeks postmenstrual age 15 weeks gestation, 131, 132 17 weeks gestation, 133 20–40 weeks postmenstrual age 21 weeks gestation, 134, 135 23 weeks gestation, 134–135 26 weeks gestation, 135–137 41 weeks gestation, 135, 139 39 weeks postmenstrual age 21 weeks gestation, 135 39 weeks gestation, 137, 138

Index Fetal maceration intracardiac potassium chloride, 431 intrauterine fetal death adrenal gland, 427, 429 heart, 427, 428 kidney, 427, 429 liver, 427, 428 placenta, 429, 430 limitations, 431–434 tissue artifacts, 431, 432 Fetal zone, 215 Fetal/neonatal thyroid stroma, 227 Fetus papyraceous, 423, 427 Fibroblast growth factors (FGFs), 316, 377 Fibrous annulus, 5, 11 Follicle-stimulating hormone (FSH), 242 Follicular atresia, 179 Follicular cysts, 179 Folliculogenesis, 178 Foramen ovale, 4 Foreskin, 197–201, 204 Fossa navicularis, 203 G Gartner duct cysts, 181 Gastrointestinal tract 14–30 weeks anal canal, 67, 74 appendix, 65, 71–73 esophagus, 54–57 large intestine, 64, 70, 71 small intestine, 62, 64–69 stomach, 59–61 timing and sequence of, 53 30–40 weeks anal canal, 69, 74, 75 appendix, 69, 74, 75 esophagus, 55–58 large intestine, 69, 74, 75 small intestine, 64, 69, 70 stomach, 60–62 timing and sequence of, 53 embryology, 51 less than 14 weeks anal canal, 64 appendix, 64, 70 esophagus, 54, 55 large intestine, 64, 70 small intestine, 62, 63 stomach, 57–60 timing and sequence of, 52, 53 wall organization mucosa, 51 muscularis mucosae, 52 muscularis propria, 52 submucosa, 52 subserosa or adventitia, 52 Gerlach’s tubal tonsils, 343 Germ cells, 130 Glans penis, 197–200, 202–204, 206 Glial fibrillary acidic protein (GFAP), 246 Graafian follicle, 180 Granulopoiesis, 284 Growth hormone (GH), 242 Gut-associated lymphoid tissue, 65

Index H Hair follicles, 381, 382 Hart's line, 204 Hassall’s corpuscles, 251, 252, 255–259 Heart embryology cardiac conduction system, 4, 5 cardiac loop formation, 3 cardiac valves formation, 4 foramen ovale, 4 membranous septum, 4 ostium primum, 3 ostium secundum, 3 extramedullary hematopoiesis, 15 histology endocardium, 8, 10 epicardium, 6, 7 myocardium, 6–9 pericardium, 6 lymphocytes, 17 specialized structures chordae tendineae, 8 conduction system, 9–15, 18 fetal valve leaflets and cusps, 8, 11–13 fetal valve stroma, 8 papillary muscles, 8, 11 vacuolation, 15 Hemangioblasts, 21, 281 Hematogones, 286, 287 Hematopoiesis, 78, 95, 96, 267, 281, 284 Hematoxylin and eosin (H&E) stain, 157, 200, 202, 243, 294, 312 Hemogenic endothelium, 21 Hemophagocytosis, 272, 287 Hepatic diverticulum, 77 Hepatic lobule 16 weeks gestation, 79 19 weeks gestation, 79 22 weeks gestation, 80 23 weeks gestation, 80 28 weeks gestation, 81 30 weeks gestation, 81 37 weeks gestation, 82 endothelial sinusoids, 80, 83 hepatic cords, 79, 82 iron accumulation, 81, 83 Hepatic morphogenesis, 77 Hepatic primordium, 77 Hepatobiliary morphogenesis, 77 Heterotopic adrenal tissue, 219 Hilus cells, 180 Hydatid of Morgagni, 142 Hymen, 204, 205 Hymenal ring, 193, 194 Hypodermis, 383, 384 Hypotympanum, 331 Hypoxic stress, 221 Hyrtl’s anastomosis, 29 I Inner ear anatomy membranous labyrinth, 333 osseous labyrinth, 332 perilymphatic space, 333 periotic duct, 334 sensory end organs, 334

439 histology cochlea and cochlear duct, 344–346 cochlear aqueduct, 343, 344 endolymphatic duct, 349 otic capsule, 343 perilymphatic space, 343 periotic duct, 343, 344 vestibular system, 346, 347 Inner neuroblastic layer (INBL), 317 Internal acoustic canal, 333 Internal acoustic meatus, 333 Internal spermatic fascia, 204 Intracardiac potassium chloride (KCl), 431 Intrapulmonary karyorrhectic cells, 46, 47 Intrapulmonary lymphatics, 44, 45 Intrapulmonary squamous cells, 46, 47 Intrathyroidal inclusions, 227, 228 Intrathyroidal parathyroid gland, 238 Intrauterine fetal death adrenal gland, 427, 429 heart, 427, 428 kidney, 427, 429 liver, 427, 428 placenta, 429, 430 Iridial muscles, 314 Iris, 315 Iris pigmented epithelium (IPE), 314 Iris stroma, 314 Isthmus, 331 K Keratinocytes, 377 Kidney common excretory duct, 113 histology collecting ducts, 121, 124 cortex, 115–117, 122 glomeruli, 120 juxtaglomerular apparatus, 117, 122 medulla, 118, 123 nephrogenesis, 119, 120 nephrogenic zone, 115, 117 nephron segments, 117, 121 proximal convoluted tubules, 117, 120 renal lobe, 115 renin-expressing cells, 118, 122 rows of glomeruli, 115, 118 sclerotic glomeruli, 120 tubular growth and maturation, 117, 122 weight, 213 nephron, eighth week, 114 S-shaped nephron, 114 L Labia majora, 197, 198, 204, 208 Labia minora, 207, 208 Labioscrotal folds, 197, 198, 204 Labioscrotal swellings, 197 Lactotrophs, 243 Lamellar bone, 359, 361 Lamina cribrosa, 313 Lamina propria, 126, 161–164, 193–195, 199, 200 Langerhans cells, 378 Langhans’ fibrin, 408

440 Large intestine 14–30 weeks, 64, 70, 71 30–40 weeks, 69, 74, 75 less than 14 weeks, 64, 70 Lateral plate mesoderm, 357 Lens embryology, 319 histology, 320, 321 Lens pit, 319 Lingual tonsil, 279 Lipofuscin pigment, 225 Littre glands, 203, 205 Liver biliary tract extrahepatic biliary tree, 85–89 intrahepatic biliary tree, 81, 84, 85 ductus venosus, 78 embryology mechanism of development, 77 vascular architecture, 78 hepatic lobule 16 weeks gestation, 79 19 weeks gestation, 79 22 weeks gestation, 80 23 weeks gestation, 80 28 weeks gestation, 81 30 weeks gestation, 81 37 weeks gestation, 82 endothelial sinusoids, 80, 83 hepatic cords, 79, 82 iron accumulation, 81, 83 Lung alveolar stage development, 46, 47 embryology, 36 histology, 42, 44 bronchus-associated lymphoid tissue, 45, 46 canalicular stage embryology, 36 histology, 39, 41, 42 developmental stages, 35 embryonic stage embryology, 35 histology, 39 intrapulmonary karyorrhectic cells, 46, 47 intrapulmonary lymphatics, 44, 45 intrapulmonary squamous cells, 46, 47 postnatal pulmonary histology, 36–39 pseudoglandular stage embryology, 35 histology, 39–41 pulmonary arteries and arterioles, 43, 45 pulmonary neuroendocrine cells, 45, 46 saccular stage embryology, 36 histology, 41–44 tight clusters, 45 Lung alveolar development, 46, 47 Luteinizing hormone (LH), 242 Lymph nodes 19 weeks gestation, 270, 271 34 weeks gestation, 270, 271 41 weeks gestation, 272, 273 blood supply, 270 embryology, 269 hematopoiesis, 272

Index hemophagocytosis, 272 lymphatics, 270 Lymphatic endothelium, 23 Lymphatic system, 269 Lymphatic vessels, 29 Lymphocyte inversion, 256 Lymphocytes, 17, 251–254 Lymphoid follicles, 262, 266, 267 Lymphoid tissue, 94, 96 Lymphopoiesis, 286, 287 M Male external genitalia, 198 distal penis, 198–200 penile shaft, 200, 202, 203 scrotum, 203, 204 Male genital system, 161, 167 Mammary gland apocrine metaplasia, 389, 394 atypical giant cells, 394 complex branching, 390, 391 early branching morphogenesis, 389 exaggerated cystic change during regression, 389, 395 myoepithelial cells, 389, 393 near-term hyperplasia with early regression, 394 neonatal hyperplasia and secretion, 389, 393 neonatal regression, 391, 394, 395 nipple, lobular phase, 386 postmenstrual gestational ages, 385 prenatal and neonatal development, 385 primary ducts, 387, 392 primordia, 385 progressive branching morphogenesis, 390 specialized mesenchyme, 387, 388 tubercles of Montgomery, 387 Mature intermediate villi, 412 Mature/Graafian follicle, 179 Meckel’s cartilage, 335 Meckel’s diverticulum, 96 Medulla, 216–219 Megakaryopoiesis, 286 Melanocytes, 377 Membranous labyrinth, 333 Membranous ossification, 365 Membranous septum, 4 Meninges, 297 Merkel cells, 377, 378 Mesenchymal villi, 412, 413 Mesonephric duct remnants, 143–145, 147, 157, 161, 180 Metanephric blastema, 113, 114 Microglia, 297 Mini-puberty, 139, 140 Monochorionic diamnionic twins, 422, 423 Monopoiesis, 284 Mons pubis, 198, 204, 206, 208 Mucoid wedge, 243 Mucosa, 51, 183 Mucosal stroma, 183–185 Müllerian (paramesonephric) ducts, 187 Müllerian columnar epithelium, 193 Multiple clefts, 23 Multiple-gestation pregnancies, 421 Muscle spindles, 373 Muscularis (myosalpinx), 161–164, 183, 184, 186 Muscularis mucosae, 51–67, 69–71, 74

Index Muscularis propria, 51–67, 70, 71, 73, 125–127, 193–195 Myelopoiesis erythropoiesis, 285 granulopoiesis, 284 lymphopoiesis, 286, 287 megakaryopoiesis, 286 monopoiesis, 284 Myoblasts, 368 Myocardium, 6–9 Myocytolysis/colliquative myocytolysis, 15 Myosalpinx, 185 N Nephrogenesis, 119, 120 Nesidioblastosis, 91 Neural crest cells, 214 Neuroblastic nodules, 217 Neurohypophysis, 241, 246 Neurosensory epithelial end organs, 334 Neurosensory epithelium, 334 Nitabuch’s fibrinoid, 410 Nodular splenic heterotopia, 96 Non-keratinized squamous corneal epithelium, 312 Notch signaling pathway, 22 Nuclear basophilia, loss of adrenal gland, 427, 429, 433 heart, 427, 428 histologic features, 428 kidney, 427, 429 liver, 427, 428 maceration changes, 431 pancreatic tissue, 432 thymus gland, 433 Nucleated red blood cells (NRBCs), 416 O Oligodendrocytes, 295, 296 Oogonia, 176, 177 Optic nerve embryology, 316–318 histology, 318 Organs of Haller, 144 Osseous labyrinth, 332 Osteoblasts, 361, 364 Osteoclasts, 362, 364 Osteocytes, 361 Osteoprogenitor cells, 361 Ostium, 183 Ostium primum, 3 Ostium secundum, 3 Otic capsule, 326, 332 Otolithic membranes, 347 Outer neuroblastic layer (ONBL), 317 Ovary 19 weeks gestation, 176 central zone, 175 features, 175 follicles, 178 follicular atresia, 179 graafian follicle, 180 hilus follicle, 180 mesonephric duct remnants, 180 oocytes, 176–178 oogonia, 176, 177

441 peripheral zone, 175 primary follicles, 178–179 primordial follicles, 178 secondary/preantral follicles, 179 stroma, 176, 177 surface epithelium, 176 tertiary/antral follicle, 179 tunica albuginea, 176 Oxyphil cells, 233 P Palatine tonsils 16 weeks gestation, 275, 276 17 weeks gestation, 276 21 weeks gestation, 276, 277 27 weeks gestation, 276, 277 embryology, 275 intraepithelial lymphocytes, 276, 278 location, 275 lymphoid tissue, 277, 278 posterior tonsillar pillar, 275 tonsillar epithelium, 277, 278 Palatopharyngeal arch, 275 Paler basal cells, 164 Pancreas embryology, 91 histology endocrine, 93–95 exocrine, 92, 93 hematopoiesis, 95, 96 lymphoid tissue, 94, 96 nodular splenic heterotopia, 96 pancreatic stroma, 92 small-bowel pancreatic heterotopia, 96 Pancreatic stroma, 92 Papillary muscles, 8, 11 Para-aortic bodies, 30 Paradidymis/organ of Giraldés, 144 Paraganglia, 30, 31 Paramesonephric (Müllerian) duct, 167, 183, 187, 193 Paramesonephric remnants, 142, 143 Parathyroid gland 18 weeks gestation, 235 24 weeks gestation, 235, 236 39 weeks gestation, 237 vs. autolyzed thyroid tissue, 235 ectopic, undescended thymic lobule, 239 embryology, 233 intrathyroidal parathyroid gland, 238 vs. lymph node, 234 term gestation with identification of, 238 Parathyroid hormone, 233, 234 Paratracheal/paraesophageal tissue, 234 Paraxial mesoderm, 357, 367 Parenchymal cell, 233, 235–237 Parietal pericardium, 6 Paroophoron, 181 Pars distalis, 241–244 Pars tuberalis, 241 Pedunculated appendix epididymis, 143 Pendulous urethra, 203 Penile shaft, 200, 202, 203 Penile urethra, 197, 203 Periaortic or retroperitoneal soft tissues, 30–31 Pericardium, 6

442 Periderm, 377 Perilymphatic space, 333 Perimysium, 371, 373 Perinatal thyroid discharge, 228 Periotic duct, 334 Periurethral Skene ducts, 204, 205 Pharyngeal pituitary, 243 Pharyngeal pouch, 233 PIN-4 immunohistochemistry, 173 Pituicytes, 243, 245, 246 Pituitary gland 17 weeks gestation, 241, 242 26 weeks gestation, 242–244 35 weeks gestation, 244–246 anterior lobe, 242, 243 embryology, 241, 242 posterior lobe, 242 posterior pituitary, 243, 246 sampling, 242 Placenta embryology amnioblasts, 400 blastocyst stage, 400 embryoblast, 399 extraembryonic mesoderm, 400 extraembryonic somatopleuric mesoderm, 400–401 morula, 399, 400 syncytiotrophoblast, 399 trophoblast differentiation, 399, 400 fetal-maternal exchange, 399 fetus papyraceous, 423 histology basal plate, 408, 410–413 chorionic plate, 407–410 extraplacental membranes, 402–404 umbilical cord, 404–409 villous parenchyma, 411, 416 multiple-gestation pregnancies, 421 yolk sac remnant, 424 Placental blood vessels, 22 Plicae, 184, 185 Posterior tonsillar pillar, 275 Posterior urethral valves, 167 Postnatal pulmonary histology, 36–39 Postnuchal fluid accumulation, 30 Primary follicles, 178 Primary logettes, 282 Primitive mesenchyme, 148–151 Primordial follicles, 178 Prolactin (PRL), 242 Promontory, 332 Prospero-related homeobox protein 1 (PROX1), 23 Prostate gland, 161 17 weeks gestation, 168 20 weeks gestation, 170 24 weeks gestation, 170, 171 29 weeks gestation, 171 34 weeks gestation, 172 39 weeks gestation, 172, 173 embryology, 167 epithelial elements, 168, 170 purpose, 167 stromal elements, 170, 171 Prostatic urethra, 168–170, 172 Prostatic utricle (PU), 167–170 Proximal convoluted tubules (PCT), 117, 120 Prune belly syndrome, 167, 197 Pseudofollicular change, 220

Index Pseudoglandular stage embryology, 35 histology, 39–41 Pseudostratified columnar epithelium, 158 Pulmonary neuroendocrine cells, 45, 46 R Radial alveolar count (RAC), 46, 47 Radial migration, 303 Rathke cleft cyst, 243, 245 Rathke’s pouch, 241–243, 246 Red pulp, spleen, 266 Reichert’s cartilage, 335 Rete ovarii, 180, 181 Retina embryology, 316–319 histology, 318–321 Retinal ganglion cell (RGC) differentiation, 316 Retinal pigment epithelium (RPE), 316 Rohr’s fibrinoid, 410 Rostral-to-caudal gradient, 148, 152 S Saccular otolithic macula, 334 Saccular stage early, 41–43 embryology, 36 late, 42–44 Saccules, 41 Salivary glands, 246 bud stage, 99 canalicular stage, 99 pre-bud stage, 99 pseudoglandular stage, 99 submandibular gland 16 weeks, 100 19 weeks, 100 23 weeks, 101 27 weeks, 101, 102 35 weeks, 102 41 weeks, 102, 103 terminal bud stage, 99 “Salt and pepper” appearance, 233 Sattler’s layer, 316 Schiff method, 243 Schlemm’s canal, 314 Sclera, 313 embryology, 312 histology, 313 Sclerotic glomeruli, 120 Sclerotome, 357, 367 Scrotum, 203, 204 Sebaceous glands, 382 Secondary crests, 41 Secondary follicle, 179 Sella turcica, 241, 242, 246 Seminal colliculus, 168 Seminal vesicles 17 weeks gestation, 162 24 weeks gestation, 162, 163 29 weeks gestation, 163 34 weeks gestation, 163, 164 38 weeks gestation, 163, 164 appearance, 161 function, 161

Index Seminiferous tubules, 129, 130 Separation zone, 410 Septal islets, 93, 95 Serosa, 183, 184 Sex-determining region Y (SRY) protein, 129 Sinovaginal bulb, 193 Sinus development, 262 Sinusoids, 28 Skeletal muscle embryology, 367 epimysium, 371 fetal muscle development 17 weeks gestation, 369 23 weeks gestation, 370 fascicular arrangement, 370 fiber types, 371 muscle fascicles, 371 histology, 368–372 muscle spindles, 373 perimysium, 371 postnatal muscle development, 369, 370, 372 Skene gland, 204, 205 Skin adnexa apocrine gland, 383 hair follicle, 381, 382 hypodermis, 383, 384 nail, 383 sebaceous glands, 382 dermis, 380, 381 embryology, 377–378 epidermis 10 weeks gestation, 378 13 weeks gestation, 378, 379 23 weeks gestation, 379, 380 30 weeks gestation, 379 37 weeks gestation, 379, 380 Small intestine 14–30 weeks, 62, 64–69 30–40 weeks, 64, 69, 70 less than 14 weeks, 62, 63 Small-bowel pancreatic heterotopia, 96 Smooth muscle actin (SMA), 170 Solid cell nests, see Ultimobranchial body remnants Solid filiform appendix testis, 142 Somites, 357, 358, 367 Somitic mesoderm, 357 Sonic Hedgehog (Shh) signaling, 316 Spermatic duct, see Vas deferens Spinal cord, 297, 299 Spleen 16 weeks gestation, 262, 263 21 weeks gestation, 263, 264 23 weeks gestation, 264 24 weeks gestation, 264, 265 32 weeks gestation, 265 35 weeks gestation, 266, 267 accessory spleens, 266 capsule, 261 embryology, 261 fetal, 267 hematopoiesis, 267 histological appearance, 261 postnatal, 267 red pulp, 266 vascular tree, 261, 262 white pulp, 262–264, 266

443 Splenic lymphoid, 262 Spongiosa, 8 Spongy layer, 402 Squamous metaplasia, 168–170 Stem villi, 412, 413 Stomach 14–30 weeks, 59–61 30–40 weeks, 60–62 less than 14 weeks, 57–60 Stratified squamous epithelium, 199 Stress involution, of thymus, 258, 259 Stroma, 176, 177, 227 Stromal elements, 170, 171 Submucosa, 52 Subserosa/adventitia, 52 Superior and inferior aberrant ducts, 144 Sympathetic ganglion, 31 Syncytiotrophoblastic knots or syncytial knots, 413 T Tangential migration, 303 Tegmen tympani, 332 Terminal villi, 412, 413 Tertiary/antral follicles, 179 Testicular descent, 157 Testicular differentiation 10–20 weeks postmenstrual age fetal Leydig cells, 131–133 germ cells, 131–134 urogenital union, 134 20–40 weeks postmenstrual age fetal Leydig cells, 134 germ cells, 135 mini-puberty, 139–141 onset of meiosis, 141 puberty and complete spermiogenesis, 141, 142 tubuli recti, 138 adrenal cortical rests, 145 bipotential gonad, 130 eighth postfertilization week, 130 embryonic remnants, 145, 146 germ cell maturation, 129, 130 mesonephric remnants, 143–145 paramesonephric remnants, 142, 143 seminiferous tubules, 129, 130 tunica albuginea, 131 tunica vaginalis, 131 tunica vasculosa, 131 Testis, 129, 197, 198, 204 Testosterone, 197, 198 Thoracic duct, 29 Thymic cortical lymphocytes, 254 Thymic medulla, 255, 256 Thymic stress involution, 258, 259 Thymic tissue, 255, 256 Thymocytes, 251, 254, 256–258 Thymus gland age-associated/physiologic thymic involution, 256–258 capsule, 252 connective tissue trabeculae, 252 cortex, 252–254 embryology, 251 medulla, 255 stress-associated involution, 258, 259 thymic architecture, 251, 252 thymic tissue in ectopic sites, 255, 256

444 Thyroid colloid stains, 226 Thyroid follicles, 225, 226 Thyroid gland 15 weeks gestation, 225 18 weeks gestation, 226 21 weeks gestation, 227 26 weeks gestation, 228 34 weeks gestation, 229 autolysis of, 224 C cells, 226, 230 embryology, 223 fetal neck structures, 223, 224 fetal vs. adult, 223 intrathyroidal inclusions, 227, 228 stroma, 227 thyroid follicles, 225, 226 ultimobranchial body remnants, 229 Thyroid-stimulating hormone (TSH), 242 Tingible body macrophages, 254 Tissue artifacts, 431, 432 T lymphocytes, 254, 255, 262, 264 Trabecular meshwork, 314 Transient thymic involution, 257 Transitional zone, 216 Trophoblast cell columns, 410 Tubercles of Montgomery, 387 Tubotympanic recess, 335 Tubuli recti, 138 Tubulogenesis, 148 Tubulotympanic recess, 335 Tunica albuginea, 129–138, 143, 145, 148, 149, 176, 179, 198, 200–202, 204 Tunica dartos, 204 Tunica vaginalis, 131 Tunica vasculosa, 131, 133–138, 143, 321 Type I pneumocytes, 37 Type II pneumocytes, 37 U Ultimobranchial body remnants, 229, 231 Umbilical arteries, 25–27 Umbilical cord, 404 16 weeks gestation, 405–407 24 weeks gestation, 405 36 weeks gestation, 405 40 weeks gestation, 405 allantoic remnant, 407, 409 internal elastic lamina, 406 scattered peripheral elastic fibers, 407 vitelline duct remnant, 405, 408 Umbilical vein, 25 Umbilical vitelline duct remnant, 405, 408 Urethral orifice, 204, 205 Urinary bladder embryology, 125 histology, 126, 127 Urogenital sinus, 193 Urogenital union, 134, 147 Urothelium, 126 Uterine cervix, 190, 192 23 weeks, 191 24 weeks, 191

Index 32 weeks, 191 35 weeks, 192 37 weeks, 192 38 weeks, 192 Uterine corpus, 188, 189 18 days, 190 19 weeks, 188 21 weeks, 188, 189 35 weeks, 189 38 weeks, 190 Uterus developmental anomalies, 187 postmenstrual weeks, 187 Utricular otolithic macula, 334 Uvea embryology, 313–315 histology, 315 V Vacuolation, 15 Vacuoles, 54 Vacuolization, 53 Vagina 18 days of postnatal life, 196 18 weeks gestation, 194 23 weeks gestation, 194 25 weeks gestation, 195 32 weeks gestation, 195 embryology, 193 Vaginal introitus, 198, 204, 205 Vaginal wall, 193 Valvular dysplasia, 13 Vas deferens, 147, 148, 151 embryology, 157 histology 17 weeks gestation, 157, 158 20 weeks gestation, 158 24 weeks gestation, 158 29 weeks gestation, 158 34 weeks gestation, 158 38 weeks gestation, 158, 159 ampulla, 34 weeks gestation, 159 Vascular and lymphatic structures cystic hygroma, 29 embryology angioblast, 21 angiogenesis, 21 blood flow patterns, 22 COUP-TFII, 22 embryonic vessels, 22 ETS translocation variant 2, 22 hemangioblasts, 21 hemogenic endothelium, 21 lymphatic formation, 23 placental blood vessels, 22 vasculogenesis, 21 vitelline vascular system, 22 histology aorta, 23, 24 capillaries, 28 ductus arteriosus, 24, 25 ductus venosus, 27

Index inferior vena cava and renal veins, 28 lymphatic vessels, 29 muscular arteries, 27, 28 sinusoids, 28 umbilical arteries, 25–27 umbilical vein, 25 Hyrtl’s anastomosis, 29 periaortic or retroperitoneal soft tissues, 30–31 postnuchal fluid accumulation, 30 Vascular architecture, 78 Vasculogenesis, 21, 36 Vasculosyncytial membranes, 421 Verumontanum, 168, 169 Vestibular canaliculus, 333 Vestibular system, 346, 347 Villous parenchyma, 411, 416 Visceral pericardium, 6 Vitelline vascular system, 22 Vitreous

445 embryology, 319 histology, 321, 322 Vulvar vestibule, 204, 205 W Wharton’s jelly, 404 Woven bone, 360, 361 Y Yellow starbursts, 432 Yolk sac erythropoiesis, 281 Yolk sac remnant, 424 Z Zona glomerulosa, 215, 216 Zonular fibers, 314