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Ellis D. Avner
William E. Harmon
Patrick Niaudet
Norishige Yoshikawa
Francesco Emma
Stuart l. Goldstein 
Editors

Pediatric
Nephrology
Seventh Edition

OFFICIALLY
ENDORSED BY

1 3Reference

Pediatric Nephrology

Ellis D. Avner • William E. Harmon
Patrick Niaudet • Norishige Yoshikawa
Francesco Emma • Stuart L. Goldstein
Editors

Pediatric Nephrology
Seventh Edition

With 506 Figures and 310 Tables

Editors
Ellis D. Avner
Department of Pediatrics
Medical College of Wisconsin
Children’s Research Institute
Children’s Hospital
Health System of Wisconsin
Milwaukee, WI, USA

William E. Harmon
Boston Children’s Hospital
Harvard Medical School
Boston, MA, USA

Patrick Niaudet
Service de Néphrologie Pédiatrique
Hôpital Necker-Enfants Malades
Université Paris-Descartes
Paris, France

Norishige Yoshikawa
Department of Pediatrics
Wakayama Medical University
Wakayama City, Japan

Francesco Emma
Division of Nephrology
Bambino Gesù Children’s Hospital – IRCCS
Rome, Italy

Stuart L. Goldstein
Division of Nephrology and
Hypertension
The Heart Institute
Cincinnati Children’s Hospital
Medical Center, College of
Medicine
Cincinnati, OH, USA

ISBN 978-3-662-43595-3
ISBN 978-3-662-43596-0 (eBook)
ISBN 978-3-662-43597-7 (print and electronic bundle)
DOI 10.1007/978-3-662-43596-0
Library of Congress Control Number: 2015954467
Springer Heidelberg New York Dordrecht London
# Springer-Verlag Berlin Heidelberg 2009, 2016
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, express or implied, with respect to the material contained
herein or for any errors or omissions that may have been made.
Printed on acid-free paper
Springer-Verlag GmbH Berlin Heidelberg is part of Springer Science+Business Media (www.
springer.com)

Preface

Through its past six editions, Pediatric Nephrology has become the standard
medical reference for health care professionals treating children with kidney
disease. This new edition, published 6 years since the previous version,
reflects the tremendous increase in critical information required to translate
molecular and cellular pathophysiology into the prevention, diagnosis, and
therapy of childhood renal disorders. This text is particularly targeted to
pediatricians, pediatric nephrologists, pediatric urologists, and physicians in
training. It is also targeted to the increased number of health care professionals
comprising the multidisciplinary teams required to provide comprehensive
care for children with kidney disease and their families: geneticists, genetic
counselors, nurse specialists, dialysis personnel, nutritionists, social workers,
and mental health professionals. Finally, this reference is designed to serve the
needs of primary care physicians (internists and family practitioners) as well as
internist nephrologists who are increasingly involved in the initial evaluation
and/or the longitudinal care of children with renal disease under different
health care delivery systems evolving throughout the world.
To keep pace with the dramatic changes in pediatric renal medicine since
the publication of the previous edition, the content of the seventh edition of
Pediatric Nephrology has been completely revised, updated, and enlarged.
The seventh edition contains 83 chapters in 3 volumes, which are organized
into 12 main sections. The text begins with an overview of the basic developmental anatomy, biology, and physiology of the kidney, which provides the
basic information necessary to understand the developmental nature of pediatric renal diseases. This is followed by a comprehensive coverage of the
evaluation, diagnosis, and therapy of specific childhood kidney diseases,
including the extensive use of clinical algorithms. Of special note is a section
focused on rapidly evolving research methodologies, which are being translated into new clinical approaches and therapies for many childhood renal
diseases. The final sections focus on comprehensive, state-of-the-art reviews
of acute and chronic renal failure in childhood.
Many chapters of the seventh edition have been completely rewritten by
new authors, all recognized as global authorities in their respective areas. The
remainder of the text has been totally revised, with junior authors often joining
senior authors from the previous edition. The number of contributors has
increased by 20 %. In addition to the new section on Global Pediatric
Nephrology, which focuses on unique aspects of pediatric nephrology practice
v

vi

and the epidemiology of pediatric renal disease in different regions of the
world, all of the chapters reflect a global perspective. This has been achieved
through a dynamic, evolving relationship between the Editors and the International Pediatric Nephrology Association (IPNA). This has led to the continued endorsement of Pediatric Nephrology as the standard global
reference text in the field of childhood kidney disease. We are proud
that the IPNA logo adorns the cover of Pediatric Nephrology in recognition
of this endorsement. The Editors look forward to this dynamic interaction
with IPNA to take advantage of future opportunities that such collaboration
may provide in the areas of education and outreach.
Other major changes are also evidenced by the new publication of the
seventh edition of Pediatric Nephrology as a basic reference handbook in the
SpringerReference series. Representing advances in state-of-the-art electronic
publishing, there are regularly updated online versions of each chapter of this
text at SpringerLink.com. The new publishing format has led to a welcome
expansion of the published text to three volumes and the increased utilization
of high-resolution, color figures. Further, two new Editors, Professor
Francesco Emma of Ospedale Pediatrico Bambino Gesu in Rome, Italy, and
Professor Stuart L. Goldstein of the University of Cincinnati, USA, have
joined the Editorial Team. The addition of new Editors continues to provide
a dynamic mixture of continuity, new ideas, new perspectives, and globalism,
which has distinguished each new edition of the text. Professors Emma and
Goldstein join the four Editors from the sixth edition: Senior Editor
and Emeritus Professor Ellis D. Avner, from the Medical College of Wisconsin, USA, and Professors William E. Harmon M.D. of Harvard University,
USA, Patrick Niaudet, from the Hôpital Necker-Enfants Malades in
Paris, France; and Norishige Yoshikawa from Wakayama, Japan. The current
Editors are internationally recognized leaders in complementary areas of
pediatric nephrology and along with the more than 150 contributors reflect
the global nature of the text and the subspecialty it serves.
The Editors wish to thank a number of individuals whose efforts were
critical in the success of this project. The book would never have reached
this seventh edition without the dedication of our professional colleagues at
Springer, Gabriele Schroder, Sandra Lesny, Gregory Sutorius, and particularly
Mr. Andrew Spencer, Senior Editor of Major Reference Works, who served as
our “guide for the perplexed” in all aspects of project management. We thank
our families, and particularly our wives (Jane, Diane, Claire, Hiro, and
Elizabeth) for their support and understanding. In particular, the Senior Editor
wishes to recognize his lifetime partner in all endeavors, Jane A. Avner, Ph.D.,
for her extraordinary editorial assistance. And finally, we thank our mentors,
our students, and most importantly, our patients and their families. Without
them, our work would lack purpose.

Preface

Preface

vii

Finally, the Editors wish to dedicate this seventh edition of Pediatric Nephrology to three former Editors who passed away in 2014. Professors Martin Barratt,
Malcolm A. “Mac” Holiday, and Robert Vernier were extraordinary physicianscientists who served as mentors to a generation of pediatric nephrologists. This
text would not exist without their efforts, commitment, and selfless contributions.
Ellis D. Avner
William E. Harmon
Patrick Niaudet
Norishige Yoshikawa
Francesco Emma
Stuart L. Goldstein

Contents

Volume 1
Part I

Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

1

Embryonic Development of the Kidney . . . . . . . . . . . . . . . . .
Carlton Bates, Jacqueline Ho, and Sunder Sims-Lucas

3

2

Development of Glomerular Circulation and Function
Alda Tufro and Ashima Gulati

....

37

3

Renal Tubular Development . . . . . . . . . . . . . . . . . . . . . . . . . .
Michel Baum

61

4

Clinical Perinatal Urology . . . . . . . . . . . . . . . . . . . . . . . . . . . .
David A. Diamond and Richard S. Lee

97

5

Renal Dysplasia/Hypoplasia . . . . . . . . . . . . . . . . . . . . . . . . . . 115
Paul Goodyer and Indra R. Gupta

6

Developmental Syndromes and Malformations of the
Urinary Tract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135
Chanin Limwongse

Part II

Homeostasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

179

7

Physiology of the Developing Kidney: Sodium and Water
Homeostasis and Its Disorders . . . . . . . . . . . . . . . . . . . . . . . . 181
Nigel Madden and Howard Trachtman

8

Physiology of the Developing Kidney: Potassium
Homeostasis and Its Disorder . . . . . . . . . . . . . . . . . . . . . . . . . 219
Lisa M. Satlin and Detlef Bockenhauer

9

Physiology of the Developing Kidney: Acid-Base
Homeostasis and Its Disorders . . . . . . . . . . . . . . . . . . . . . . . . 247
Peter D. Yorgin, Elizabeth G. Ingulli, and Robert H. Mak

10

Bone Developmental Physiology . . . . . . . . . . . . . . . . . . . . . . . 279
MH Lafage-Proust
ix

x

Contents

11

Physiology of the Developing Kidney: Disorders and
Therapy of Calcium and Phosphorous Homeostasis . . . . . . . 291
Amita Sharma, Rajesh V. Thakker, and Harald J€uppner

12

Nutrition Management in Childhood Kidney Disease:
An Integrative and Lifecourse Approach . . . . . . . . . . . . . . . . 341
Lauren Graf, Kimberly Reidy, and Frederick J. Kaskel

13

Physiology of the Developing Kidney: Fluid and
Electrolyte Homeostasis and Therapy of Basic
Disorders (Na/H2O/K/Acid Base) . . . . . . . . . . . . . . . . . . . . . . 361
Isa F. Ashoor and Michael J. G. Somers

Part III Translational Research Methods

....................

423

14

Translational Research Methods: Basics of Renal Molecular
Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 425
Gian Marco Ghiggeri, Maurizio Bruschi, and
Simone Sanna-Cherchi

15

Translational Research Methods: The Value of Animal
Models in Renal Research . . . . . . . . . . . . . . . . . . . . . . . . . . . . 447
Jordan Kreidberg

16

Basics of Clinical Investigation . . . . . . . . . . . . . . . . . . . . . . . . 473
Susan L. Furth and Jeffrey J. Fadrowski

17

Genomic Methods in the Diagnosis and Treatment of
Pediatric Kidney Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 499
Karen Maresso and Ulrich Broeckel

18

Translational Research Methods: Renal Stem Cells . . . . . . . 525
Kenji Osafune

19

Translational Research Methods: Tissue Engineering of the
Kidney and Urinary Tract . . . . . . . . . . . . . . . . . . . . . . . . . . . 571
Austin G. Hester and Anthony Atala

Part IV Clinical Approach to the Child with Suspected Renal
Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

593

20

Clinical Evaluation of the Child with Suspected
Renal Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 595
Mohan A. Shenoy and Nicholas J. A. Webb

21

Laboratory Investigation of the Child with Suspected
Renal Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 613
George van der Watt, Fierdoz Omar, Anita Brink, and
Mignon McCulloch

Contents

xi

22

Growth and Development of the Child with
Renal Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 637
Bethany Foster

23

Diagnostic Imaging of the Child with Suspected
Renal Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 667
Jonathan Loewen and Larry A. Greenbaum

24

Pediatric Renal Pathology . . . . . . . . . . . . . . . . . . . . . . . . . . . . 705
Agnes B. Fogo

Part V

Glomerular Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

751

25

Congenital Nephrotic Syndrome . . . . . . . . . . . . . . . . . . . . . . . 753
Hannu Jalanko and Christer Holmberg

26

Inherited Glomerular Diseases . . . . . . . . . . . . . . . . . . . . . . . . 777
Michelle N. Rheault and Clifford E. Kashtan

27

Idiopathic Nephrotic Syndrome in Children: Genetic
Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 805
Olivia Boyer, Kálmán Tory, Eduardo Machuca, and
Corinne Antignac

28

Idiopathic Nephrotic Syndrome in Children: Clinical
Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 839
Patrick Niaudet and Olivia Boyer

29

Immune-Mediated Glomerular Injury in Children . . . . . . . . 883
Michio Nagata

30

Complement-Mediated Glomerular Injury in
Children . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 927
Zoltán Prohászka, Marina Vivarelli, and
George S. Reusz

31

Acute Postinfectious Glomerulonephritis in Children . . . . . . 959
Bernardo Rodríguez-Iturbe, Behzad Najafian,
Alfonso Silva, and Charles E. Alpers

32

Immunoglobulin A Nephropathies in Children
(Includes HSP) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 983
Koichi Nakanishi and Norishige Yoshikawa

33

Membranoproliferative and C3-Mediated
GN in Children . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1035
Christoph Licht, Magdalena Riedl, Matthew C. Pickering, and
Michael Braun

34

Membranous Nephropathy in Children . . . . . . . . . . . . . . . . . 1055
Rudolph P. Valentini

xii

Contents

Volume 2
Part VI

Tubular Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1077

35

Nephronophthisis and Medullary Cystic Kidney Disease in
Children . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1079
Friedhelm Hildebrandt

36

Childhood Polycystic Kidney Disease . . . . . . . . . . . . . . . . . . . 1103
William E. Sweeney Jr., Meral Gunay-Aygun,
Ameya Patil, and Ellis D. Avner

37

Aminoaciduria and Glycosuria in Children . . . . . . . . . . . . . . 1155
Israel Zelikovic

38

Renal Tubular Disorders of Electrolyte Regulation
in Children . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1201
Olivier Devuyst, Hendrica Belge, Martin Konrad,
Xavier Jeunemaitre, and Maria-Christina Zennaro

39

Renal Tubular Acidosis in Children . . . . . . . . . . . . . . . . . . . . 1273
Raymond Quigley and Matthias T. F. Wolf

40

Nephrogenic Diabetes Insipidus in Children . . . . . . . . . . . . . 1307
Nine V. A. M. Knoers and Elena N. Levtchenko

41

Cystinosis and Its Renal Complications in Children . . . . . . . 1329
William A. Gahl and Galina Nesterova

42

Pediatric Fanconi Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . 1355
Takashi Igarashi

43

Primary Hyperoaxaluria in Children . . . . . . . . . . . . . . . . . . . 1389
Pierre Cochat, Neville Jamieson, and
Cecile Acquaviva-Bourdain

44

Pediatric Tubulointerstitial Nephritis . . . . . . . . . . . . . . . . . . . 1407
Uri S. Alon

Part VII

Kidney Involvement in Systemic Diseases . . . . . . . . . . . 1429

45

Renal Involvement in Children with Vasculitis . . . . . . . . . . . 1431
Seza Ozen and Diclehan Orhan

46

Renal Involvement in Children with Systemic Lupus
Erythematosus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1449
Patrick Niaudet, Brigitte Bader-Meunier, and Rémi Salomon

47

Renal Involvement in Children with HUS . . . . . . . . . . . . . . . 1489
Carla M. Nester and Sharon P. Andreoli

Contents

xiii

48

Sickle Cell Nephropathy in Children . . . . . . . . . . . . . . . . . . . 1523
Connie Piccone and Katherine MacRae Dell

49

Diabetic Nephropathy in Children . . . . . . . . . . . . . . . . . . . . . 1545
M. Loredana Marcovecchio and Francesco Chiarelli

50

Renal Manifestations of Metabolic Disorders
in Children . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1569
Francesco Emma, William G. van’t Hoff, and
Carlo Dionisi Vici

51

Infectious Diseases and the Kidney in Children . . . . . . . . . . 1609
Jennifer Stevens, Jethro A. Herberg, and Michael Levin

52

Nephrotoxins and Pediatric Kidney Injury . . . . . . . . . . . . . . 1655
Takashi Sekine

Part VIII

Urinary Tract Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1693

53

Urinary Tract Infections in Children . . . . . . . . . . . . . . . . . . . 1695
Elisabeth M. Hodson and Jonathan C. Craig

54

Vesicoureteral Reflux and Renal Scarring in Children . . . . . 1715
Tej K. Mattoo, Ranjiv Mathews, and Indra R. Gupta

55

Pediatric Obstructive Uropathy . . . . . . . . . . . . . . . . . . . . . . . 1749
Bärbel Lange-Sperandio

56

Pediatric Bladder Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . 1779
Etienne Berard

57

Urolithiasis in Children . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1821
Vidar Edvardsson

58

Pediatric Renal Tumors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1869
Elizabeth Mullen, Jordan Kreidberg, and
Christopher B. Weldon

Volume 3
Part IX

Hypertension . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1905

59

Epidemiology of Hypertension in Children . . . . . . . . . . . . . . 1907
Midori Awazu

60

Pathophysiology of Pediatric Hypertension . . . . . . . . . . . . . . 1951
Ikuyo Yamaguchi and Joseph T. Flynn

61

Evaluation of Hypertension in Childhood Diseases . . . . . . . . 1997
Eileen D. Brewer and Sarah J. Swartz

xiv

62

Contents

Management of the Hypertensive Child . . . . . . . . . . . . . . . . . 2023
Demetrius Ellis and Yosuke Miyashita

Part X

Acute Renal Injury . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2099

63

Pathogenesis of Acute Kidney Injury . . . . . . . . . . . . . . . . . . . 2101
David P. Basile, Rajasree Sreedharan, and
Scott K. Van Why

64

Evaluation and Management of Acute Kidney Injury in
Children . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2139
Stuart L. Goldstein and Michael Zappitelli

Part XI

Chronic Renal Failure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2169

65

Pathophysiology of Progressive Renal Disease
in Children . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2171
H. William Schnaper

66

Management of Chronic Kidney Disease in Children . . . . . . 2207
Rene G. VanDeVoorde, Craig S. Wong, and
Bradley A. Warady

67

Handling of Drugs in Children with Abnormal
Renal Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2267
Guido Filler, Amrit Kirpalani, and Bradley L. Urquhart

68

Endocrine and Growth Abnormalities in Chronic
Kidney Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2295
Franz Schaefer

69

Mineral and Bone Disorders in Children with Chronic
Kidney Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2349
Katherine Wesseling-Perry and Isidro B. Salusky

70

Peritoneal Dialysis in Children . . . . . . . . . . . . . . . . . . . . . . . . 2381
Enrico Verrina and Claus Peter Schmitt

71

Hemodialysis in Children . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2433
Lesley Rees

72

Immunology of Pediatric Renal Transplantation . . . . . . . . . 2457
Elizabeth G. Ingulli, Stephen I. Alexander, and David M. Briscoe

73

Pediatric Renal Transplantation . . . . . . . . . . . . . . . . . . . . . . . 2501
Nancy M. Rodig, Khashayar Vakili, and William E. Harmon

74

Immunosuppression for Pediatric Renal
Transplantation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2553
Jodi M. Smith, Thomas L. Nemeth, and Ruth A. McDonald

75

Complications of Pediatric Renal Transplantation . . . . . . . . 2573
Vikas R. Dharnidharka and Carlos E. Araya

Contents

xv

Part XII

IPNA: Global Pediatric Nephrology . . . . . . . . . . . . . . . . . 2605

76

IPNA: Global Pediatric Nephrology, Introduction and
Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2607
Pierre Cochat and Isidro B. Salusky

77

AFPNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2613
Mignon McCulloch, Hesham Safouh, Amal Bourquia, and
Priya Gajjar

78

ALANEPE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2631
Vera Koch, Nelson Orta, and Ramon Exeni

79

AsPNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2639
Hui-Kim Yap, Man-Chun Chiu, Arvind Bagga, and
Hesham Safouh

80

Pediatric Nephrology in North America
Victoria F. Norwood and Maury Pinsk

81

ANZPNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2673
Deborah Lewis

82

ESPN . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2681
Rosanna Coppo

83

JSPN . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2687
Kazumoto Iijima

. . . . . . . . . . . . . . . . 2665

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2697

Contributors

Cecile Acquaviva-Bourdain Centre de référence des maladies rénales rares
Néphrogones, Hôpital Femme Mère Enfant, Hospices Civils de Lyon &
Université de Lyon, Lyon, France
Service Maladies Héréditaires du Métabolisme et Dépistage Néonatal, Centre
de Biologie et Pathologie Est, Hospices Civils de Lyon, Lyon, France
Stephen I. Alexander Discipline of Paediatrics and Child Health, The
University of Sydney, Sydney, New South Wales, Australia
Division of Nephrology, Children’s Hospital at Westmead, Sydney, New
South Wales, Australia
Uri S. Alon Section of Pediatric Nephrology, The Children’s Mercy Hospital
and Clinics, University of Missouri at Kansas City, School of Medicine,
Kansas City, MO, USA
Charles E. Alpers Department of Pathology, University of Washington,
Seattle, WA, USA
Sharon P. Andreoli Division of Nephrology, Indianapolis, IN, USA
Corinne Antignac Laboratory of Hereditary Kidney Diseases, Inserm UMR
1163, Paris, France
Paris Descartes, Sorbonne Paris Cité University, Imagine Institute, Paris,
France
Department of Genetics, MARHEA reference center, Necker – Enfants
Malades Hospital, Paris, France
Carlos E. Araya University of Central Florida and Nemours Children’s
Hospital, Orlando, FL, USA
Isa F. Ashoor Division of Nephrology, Children’s Hospital, New Orleans,
LA, USA
Anthony Atala School of Medicine, Department of Urology, Wake Forest
Institute for Regenerative Medicine, Wake Forest University, Winston Salem,
NC, USA

xvii

xviii

Contributors

Ellis D. Avner Department of Pediatrics, Medical College of Wisconsin,
Children’s Research Institute, Children’s Hospital Health System of Wisconsin, Milwaukee, WI, USA
Midori Awazu Department of Pediatrics, Keio University School of Medicine, Shinjuku-ku, Tokyo, Japan
Brigitte Bader-Meunier Service d’Immunologie et
Pédiatrique, Hôpital Necker-Enfants Malades, Paris, France

Rhumatologie

Arvind Bagga Division of Nephrology, All India Institute of Medical
Sciences, New Delhi, India
David P. Basile Indiana University School of Medicine, Indianapolis, IN, USA
Carlton Bates Department of Pediatrics, Division of Pediatric Nephrology,
Children’s Hospital of Pittsburgh of UPMC, University of Pittsburgh School
of Medicine, Pittsburgh, PA, USA
Michel Baum Departments of Pediatrics and Internal Medicine, University
of Texas Southwestern Medical Center at Dallas, Dallas, TX, USA
Hendrica Belge Zurich Institute for Human Physiology, University of
Zurich, Institute of Physiology, Zurich Center for Integrative Human Physiology (ZIHP), Z€
urich, Switzerland
Etienne Berard Pediatric Nephrology Unit, Universitary Hospital of Nice
(France), Nice, France
Detlef Bockenhauer Great Ormond Street Hospital, Institute of Child
Health, University College London, London, UK
Amal Bourquia Paediatric Nephology, Red Cross War Memorial Children’s
Hospital, Dept. of Paediatric Medicine, University of Cape Town, Western
Cape, South Africa
Olivia Boyer Service de Néphrologie Pédiatrique, Hôpital Necker-Enfants
Malades, Université Paris-Descartes, Paris, France
Michael Braun Renal Section, Department of Pediatrics, Texas Children’s
Hospital, Balyor College of Medicine, Houston, TX, USA
Eileen D. Brewer Department of Pediatrics Renal Section, Baylor College of
Medicine and Texas Children’s Hospital, Houston, TX, USA
Anita Brink Department of Pediatrics and Child Health (Nuclear Medicine),
University of Cape Town, Red Cross War Memorial Children’s Hospital, Cape
Town, South Africa
David M. Briscoe Department of Pediatrics, Harvard Medical School,
Boston, MA, USA
Division of Nephrology, Transplant Research Program, Boston Children’s
Hospital, Boston, MA, USA

Contributors

xix

Ulrich Broeckel Department of Pediatrics, Medical College of Wisconsin
and Children’s Hospital of Wisconsin, Milwaukee, WI, USA
Maurizio Bruschi Laboratory of Physiopathology of Uremia, Division of
Nephrology, Dialysis and Transplantation, Istituto Giannina Gaslini, Genoa,
Italy
Francesco Chiarelli University of Chieti, Chieti, Italy
Man-Chun Chiu Department of Pediatrics and Adolescent Medicine, Princess Margaret Hospital, Hong Kong University, Kowloon, Hong Kong
Pierre Cochat Centre de référence des maladies rénales rares Néphrogones,
Hôpital Femme Mère Enfant, Hospices Civils de Lyon & Université de Lyon,
Lyon, France
IBCP-UMR 5305 CNRS, Université Claude-Bernard Lyon 1, Lyon, France
Rosanna Coppo Nephrology Dialysis and Transplantation Unit, Azienda
Ospedaliera-Universitaria Città della Salute e della Scienza di Torino, Regina
Margherita Children’s University Hospital, Turin, Italy
Jonathan C. Craig Centre for Kidney Research, The Children’s Hospital at
Westmead, Westmead, Sydney, NSW, Australia
Sydney School of Public Health, University of Sydney, Sydney, NSW,
Australia
Katherine MacRae Dell Center for Pediatric Nephrology, Department of
Pediatrics, Cleveland Clinic Children’s and Case Western Reserve University,
Cleveland, OH, USA
Olivier Devuyst Zurich Institute for Human Physiology, University of
Zurich, Institute of Physiology, Zurich Center for Integrative Human Physiology (ZIHP), Z€urich, Switzerland
Vikas R. Dharnidharka Pediatric Nephrology, Washington University
School of Medicine and St. Louis Children’s Hospital, St. Louis, MO, USA
David A. Diamond Department of Urology, Harvard Medical School,
Boston Children’s Hospital, Boston, MA, USA
Carlo Dionisi Vici Division of Metabolic Diseases, Bambino Gesù
Children’s Hospital – IRCCS, Rome, Italy
Vidar Edvardsson Landspitali – The National University Hospital of Iceland, Reykjavik, Iceland and Faculty of Medicine, School of Health Sciences,
University of Iceland, Reykjavik, Iceland
Demetrius Ellis Department of Pediatrics, Division of Pediatric Nephrology,
Children’s Hospital of Pittsburgh of UPMC, University of Pittsburgh School
of Medicine, Pittsburgh, PA, USA
Francesco Emma Division of Nephrology, Bambino Gesù Children’s
Hospital – IRCCS, Rome, Italy

xx

Ramon Exeni Department of Nephrology, Children’s Hospital “San Justo”,
Buenos Aires, Argentina
Jeffrey J. Fadrowski Department of Pediatrics, John Hopkins University
School of Medicine, Baltimore, MD, USA
Guido Filler Departments of Pediatrics, Medicine, and Pathology Laboratory
Medicine, Schulich School of Medicine and Dentistry, Western University,
London, ON, Canada
Children’s Hospital of Western Ontario, Children’s Health Research Institute
(CHRI), London, ON, Canada
Joseph T. Flynn Division of Nephrology, Seattle Children’s Hospital;
Department of Pediatrics, University of Washington School of Medicine,
Seattle, WA, USA
Agnes B. Fogo Department of Pathology, Microbiology and Immunology,
Vanderbilt University Medical Center, Nashville, TN, USA
Bethany Foster The Research Institute of the McGill University Health
Centre, Montreal, QC, Canada
Susan L. Furth Departments of Pediatrics and Epidemiology, Perelman
School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA
William A. Gahl Section on Human Biochemical Genetics, Medical Genetics Branch, National Human Genome Research Institute, National Institutes of
Health, Bethesda, MD, USA
Priya Gajjar Paediatric Nephology, Red Cross War Memorial Children’s
Hospital, Dept. of Paediatric Medicine, University of Cape Town, Western
Cape, South Africa
Gian Marco Ghiggeri Laboratory of Physiopathology of Uremia, Division
of Nephrology, Dialysis and Transplantation, Istituto Giannina Gaslini,
Genoa, Italy
Stuart L. Goldstein Division of Nephrology and Hypertension, The Heart
Institute, Cincinnati Children’s Hospital Medical Center, College of Medicine,
Cincinnati, OH, USA
Paul Goodyer Division of Pediatric Nephrology, Montreal Children’s
Hospital, McGill University, Montreal, QC, Canada
Lauren Graf Children’s Hospital at Montefiore, Albert Einstein College of
Medicine, Bronx, NY, USA
Larry A. Greenbaum Department of Pediatric Radiology, Emory University
School of Medicine, Atlanta, GA, USA
Ashima Gulati Department of Pediatrics, Nephrology Section, Yale School
of Medicine, New Haven, CT, USA

Contributors

Contributors

xxi

Meral Gunay-Aygun Medical Genetics Branch, The Intramural Program of
the Office of Rare Diseases, National Human Genome Research Institute,
Bethesda, MD, USA
Department of Pediatrics, McKusick-Nathans Institute of Genetic Medicine,
Johns Hopkins University School of Medicine, Baltimore, MD, USA
Indra R. Gupta Department of Pediatrics and Department of Human Genetics, Division of Pediatric Nephrology, Montreal Children’s Hospital, McGill
University, Montréal, QC, Canada
William E. Harmon Boston Children’s Hospital, Harvard Medical School,
Boston, MA, USA
Jethro A. Herberg Imperial College London, London, UK
Austin G. Hester School of Medicine, Department of Urology, Wake Forest
Institute for Regenerative Medicine, Wake Forest University, Winston Salem,
NC, USA
Friedhelm Hildebrandt Harvard Medical School, Boston, MA, USA
Jacqueline Ho Department of Pediatrics, Division of Pediatric Nephrology,
Children’s Hospital of Pittsburgh of UPMC, University of Pittsburgh School
of Medicine, Pittsburgh, PA, USA
Elisabeth M. Hodson Centre for Kidney Research, The Children’s Hospital
at Westmead, Westmead, Sydney, NSW, Australia
Sydney School of Public Health, University of Sydney, Sydney, NSW,
Australia
Christer Holmberg Hospital for Children and Adolescents, University of
Helsinki, Helsinki, Finland
Takashi Igarashi National Center for Child Health and Development
(NCCHD), Tokyo, Japan
Kazumoto Iijima Department of Pediatrics, Kobe University Graduate
School of Medicine, Kobe, Japan
Elizabeth G. Ingulli Department of Pediatrics, University of California,
San Diego, CA, USA
Division of Nephrology, Rady Children’s Hospital San Diego, San Diego,
CA, USA
Kidney Transplant Program, Rady Children’s Hospital, San Diego, CA, USA
Hannu Jalanko Hospital for Children and Adolescents, University of
Helsinki, Helsinki, Finland
Neville Jamieson Department of Surgery, Addenbrookes Hospital,
Cambridge University Teaching Hospitals, Cambridge, UK
Xavier Jeunemaitre Department of Molecular Genetics, Hôpital Européen
George Pompidou, Paris, France

xxii

Harald J€
uppner Departments of Medicine and Pediatrics, Endocrine Unit
and Pediatric Nephrology Unit, Massachusetts General Hospital and Harvard
Medical School, Boston, MA, USA
Clifford E. Kashtan Department of Pediatrics, Division of Pediatric
Nephrology, University of Minnesota Masonic Children’s Hospital, Minneapolis, MN, USA
Frederick J. Kaskel Division of Pediatric Nephrology, Children’s Hospital
at Montefiore, Albert Einstein College of Medicine, Bronx, NY, USA
Amrit Kirpalani Departments of Pediatrics, Schulich School of Medicine
and Dentistry, Western University, London, ON, Canada
Nine V. A. M. Knoers Departments of Medical Genetics, University Medical
Centre Utrecht, Utrecht, The Netherlands
Vera Koch Instituto da Criança- Pediatric Nephrology Unit, Department of
Pediatrics, University of Sao Paulo Medical School, Sao Paulo, Brazil
Martin Konrad Department of General Pediatrics, Pediatric Nephrology,
University Hospital, M€unster, Germany
Jordan Kreidberg Children’s Hospital Boston, Boston, MA, USA
MH Lafage-Proust INSERM U 1059, Université de Lyon, Saint-Etienne,
France
Bärbel Lange-Sperandio Dr. v. Hauner Children’s Hospital, Department of
Pediatric Nephrology, LMU, Munich, Germany
Richard S. Lee Department of Urology, Harvard Medical School, Boston
Children’s Hospital, Boston, MA, USA
Michael Levin Department of Medicine, Imperial College London, London,
UK
Elena N. Levtchenko Department of Pediatric Nephrology, Department of
Growth and Regeneration, University Hospitals Leuven, Katholieke
Universiteit Leuven, Leuven, Belgium
Deborah Lewis Sydney, NSW, Australia
Christoph Licht Division of Nephrology, The Hospital for Sick Children,
University of Toronto, Toronto, ON, Canada
Research Institute, Cell Biology Program, The Hospital for Sick Children,
Toronto, ON, Canada
Department of Paediatrics, University of Toronto, Toronto, ON, Canada
Chanin Limwongse Department of Medicine, Division of Medical Genetics,
Faculty of Medicine, Siriraj Hospital, Mahidol University, Bangkoknoi,
Bangkok, Thailand
Jonathan Loewen Department of Pediatric Radiology, Emory University
School of Medicine, Atlanta, GA, USA

Contributors

Contributors

xxiii

Eduardo Machuca Department of Nephrology, Medical School, Pontificia
Universidad Católica de Chile, Santiago, Chile
Nigel Madden NYU Langone Medical Center and NYU School of
Medicine, New York, NY, USA
Robert H. Mak Pediatric Nephrology, University of California, San Diego,
CA, USA
Pediatric Nephrology Division, Rady Children’s Hospital, San Diego, CA, USA
M. Loredana Marcovecchio University of Chieti, Chieti, Italy
Karen Maresso Section of Genomic Pediatrics, Medical College of
Wisconsin, Milwaukee, WI, USA
Ranjiv Mathews The Nevada School of Medicine, The Johns Hopkins
School of Medicine, Las Vegas, NV, USA
Tej K. Mattoo Pediatric Nephrology and Hypertension, Wayne State
University School of Medicine, Children’s Hospital of Michigan, Detroit,
MI, USA
Mignon McCulloch Department of Paediatric Intensive Care/Nephrology,
University of Cape Town, Red Cross War Memorial Children’s Hospital, Cape
Town, Western Cape, South Africa
Ruth A. McDonald Division of Nephrology, Seattle Children’s, University
of Washington, Seattle, WA, USA
Yosuke Miyashita Department of Pediatrics, Division of Pediatric Nephrology, Children’s Hospital of Pittsburgh of UPMC, University of Pittsburgh
School of Medicine, Pittsburgh, PA, USA
Elizabeth Mullen Hematology Oncology, Dana-Farber/Boston Children’s
Blood Disorders and Cancer Center, Boston, MA, USA
Michio Nagata Graduate School of Comprehensive Human Sciences,
University of Tsukuba, Tsukuba, Japan
Behzad Najafian Department of Pathology, University of Washington,
Seattle, WA, USA
Koichi Nakanishi Department of Pediatrics, Wakayama Medical University,
Wakayama City, Japan
Thomas L. Nemeth Department of Pharmacy, Seattle Children’s, University
of Washington, Seattle, WA, USA
Carla M. Nester Stead Family Department of Pediatrics, Department of
Internal Medicine, Divisions of Nephrology, University of Iowa Children’s
Hospital, Carver College of Medicine, Iowa City, IA, USA
Galina Nesterova Section on Human Biochemical Genetics, Medical Genetics Branch, National Human Genome Research Institute, National Institutes of
Health, Bethesda, MD, USA

xxiv

Patrick Niaudet Service de Néphrologie Pédiatrique, Hôpital NeckerEnfants Malades, Université Paris-Descartes, Paris, France
Victoria F. Norwood University of Virginia, Charlottesville, VA, USA
Fierdoz Omar Chemical Pathology, University of Cape Town and National
Health Laboratory Service, Red Cross Children’s Hospital and Groote Schuur
Hospital, Cape Town, South Africa
Diclehan Orhan Department of Pediatric Pathology, Hacettepe University,
Sihhiye, Ankara, Turkey
Nelson Orta Service of Pediatric Nephrology, Children’s Hospital “Jorge
Lizarraga”, University of Carabobo, Valencia, Venezuela
Kenji Osafune Center for iPS Cell Research and Application (CiRA), Kyoto
University, Sakyo-ku, Kyoto, Japan
Seza Ozen Department of Pediatrics, Faculty of Medicine, Hacettepe
University, Sihhiye, Ankara, Turkey
Ameya Patil Department of Pediatrics, Medical College of Wisconsin, Children’s Research Institute, Children’s Hospital Health System of Wisconsin,
Milwaukee, WI, USA
Connie Piccone Rainbow Babies and Children’s Hospital, Cleveland,
OH, USA
Matthew C. Pickering Centre for Complement and Inflammation Research,
Imperial College, London, UK
Maury Pinsk University of Alberta, Edmonton, AB, Canada
Zoltán Prohászka 3rd Department of Medicine, Faculty of Medicine,
Semmelweis University, Budapest, Hungary
Raymond Quigley Department of Pediatrics, University of Texas Southwestern Medical Center at Dallas, Dallas, TX, USA
Lesley Rees Department of Nephrology, Great Ormond Street Hospital for
Children NHS Trust, London, UK
Kimberly Reidy Division of Pediatric Nephrology, Children’s Hospital at
Montefiore, Albert Einstein College of Medicine, Bronx, NY, USA
George S. Reusz 1st Department of Pediatrics, Faculty of Medicine,
Semmelweis University, Budapest, Hungary
Michelle N. Rheault Department of Pediatrics, Division of Pediatric
Nephrology, University of Minnesota Masonic Children’s Hospital, Minneapolis, MN, USA
Magdalena Riedl Research Institute, Cell Biology Program, The Hospital
for Sick Children, Toronto, ON, Canada
Department of Paediatrics, Innsbruck Medical University, Innsbruck, Tyrol,
Austria

Contributors

Contributors

xxv

Nancy M. Rodig Boston Children’s Hospital, Harvard Medical School,
Boston, MA, USA
Bernardo Rodríguez-Iturbe Nephrology Service, Hospital Universitario,
Maracaibo, Estado Zulia, Venezuela
Hesham Safouh Faculty of Medicine, Center for Pediatric Nephrology and
Transplantation (CPNT), Cairo University, Orman, Giza, Egypt
Rémi Salomon Service de Néphrologie Pédiatrique, Hôpital Necker-Enfants
Malades, Université Paris-Descartes, Paris, France
Isidro B. Salusky Division of Pediatric Nephrology, Clinical Translational
Research Center, David Geffen School of Medicine at UCLA, Los Angeles,
CA, USA
Simone Sanna-Cherchi Division of Nephrology, Columbia University,
College of Physicians and Surgeons, New York, NY, USA
Lisa M. Satlin Department of Pediatrics, Division of Pediatric Nephrology,
Icahn School of Medicine at Mount Sinai, New York, NY, USA
Franz Schaefer Division of Pediatric Nephrology, University Children’s
Hospital, Heidelberg, Germany
Claus Peter Schmitt Centre for Pediatric and Adolescent Medicine, Heidelberg, Germany
H. William Schnaper Division of Kidney Diseases, Ann and Robert H. Lurie
Children’s Hospital of Chicago, Department of Pediatrics, Northwestern
University Feinberg School of Medicine, Chicago, IL, USA
Takashi Sekine Department of Pediatrics, Toho University Faculty of
Medicine, Meguro-ku, Tokyo, Japan
Amita Sharma Department of Pediatrics, Pediatric Nephrology Unit, Massachusetts General Hospital and Harvard Medical School, Boston, MA, USA
Mohan A. Shenoy Department of Pediatric Nephrology, Royal Manchester
Children’s Hospital, Manchester, UK
Alfonso Silva Nephrology Service, Hospital Universitario, Maracaibo,
Estado Zulia, Venezuela
Sunder Sims-Lucas Department of Pediatrics, Division of Pediatric
Nephrology, Children’s Hospital of Pittsburgh of UPMC, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA
Jodi M. Smith Division of Nephrology, Seattle Children’s, University of
Washington, Seattle, WA, USA
Michael J. G. Somers Division of Nephrology, Boston Children’s Hospital,
Harvard Medical School, Boston, MA, USA
Rajasree Sreedharan Medical College of Wisconsin, Milwaukee, WI, USA
Jennifer Stevens University Hospital Wales, Cardiff, S. Wales, UK

xxvi

Sarah J. Swartz Department of Pediatrics Renal Section, Baylor College of
Medicine and Texas Children’s Hospital, Houston, TX, USA
William E. Sweeney Jr. Department of Pediatrics, Medical College of Wisconsin, Children’s Research Institute, Children’s Hospital Health System of
Wisconsin, Milwaukee, WI, USA
Rajesh V. Thakker Radcliffe Department of Medicine, Academic Endocrine
Unit, University of Oxford, OCDEM (Oxford Centre for Diabetes, Endocrinology and Metabolism), The Churchill Hospital Headington, Oxford, UK
Kálmán Tory Laboratory of Hereditary Kidney Diseases, Inserm UMR
1163, Paris, France
Department of Pediatrics, Semmelweis University, Budapest, Hungary
Howard Trachtman NYU Langone Medical Center and NYU School of
Medicine, New York, NY, USA
Alda Tufro Department of Pediatrics, Nephrology Section, Yale School of
Medicine, New Haven, CT, USA
Bradley L. Urquhart Departments of Pediatrics, Department of Medicine,
Physiology and Pharmacology, Western University, London, ON, Canada
Children’s Health Research Institute (CHRI), London, ON, Canada
Khashayar Vakili Boston Children’s Hospital, Harvard Medical School,
Boston, MA, USA
Rudolph P. Valentini Pediatric Nephrology, Children’s Hospital of Michigan, Detroit, MI, USA
Wayne State University School of Medicine, Detroit, MI, USA
George van der Watt Chemical Pathology, University of Cape Town and
National Health Laboratory Service, Red Cross Children’s Hospital and
Groote Schuur Hospital, Cape Town, South Africa
Scott K. Van Why Medical College of Wisconsin, Milwaukee, WI, USA
William G. van’t Hoff Great Ormond Street Hospital, London, UK
Rene G. VanDeVoorde Cincinnati Children’s Hospital Medical Center,
Cincinnati, OH, USA
Enrico Verrina Nephrology, Dialysis and Transplantation Unit, Giannina
Gaslini Childrens Hospital, Genoa, Italy
Marina Vivarelli Division of Nephrology and Dialysis, Children’s Hospital
Bambino Gesù-IRCCS, Rome, Italy
Bradley A. Warady Pediatric Nephrology, Children’s Mercy Hospital,
Kansas City, MO, USA
Nicholas J. A. Webb Department of Pediatric Nephrology, Royal Manchester Children’s Hospital, Manchester, UK

Contributors

Contributors

xxvii

Christopher B. Weldon Department of Surgery, Boston Children’s Hospital
and Harvard Medical School, Boston, MA, USA
Katherine Wesseling-Perry Division of Pediatric Nephrology, David
Geffen School of Medicine at UCLA, Los Angeles, CA, USA
Matthias T. F. Wolf Department of Pediatrics, University of Texas Southwestern Medical Center at Dallas, Dallas, TX, USA
Craig S. Wong Pediatric Nephrology, University of New Mexico Health
Sciences Center, Albuquerque, NM, USA
Ikuyo Yamaguchi Division of Pediatric Nephrology, University of Texas
School of Medicine at San Antonio, University of Texas Health Science Center
at San Antonio, San Antonio, TX, USA
Hui-Kim Yap Yong Loo Lin School of Medicine, National University of
Singapore, Singapore, Singapore
Peter D. Yorgin Pediatric Nephrology, University of California, San Diego,
CA, USA
Pediatric Nephrology Division, Rady Children’s Hospital, San Diego,
CA, USA
Norishige Yoshikawa Department of Pediatrics, Wakayama Medical
University, Wakayama City, Japan
Michael Zappitelli Pediatrics, Division of Nephrology, Montreal Children’s
Hospital, McGill University Health Center, Montreal, QC, Canada
Israel Zelikovic Department of Physiology and Biophysics, Faculty of
Medicine, Technion – Israel Institute of Technology, Haifa, Israel
Division of Pediatric Nephrology, Rambam Medical Center, Haifa, Israel
Maria-Christina Zennaro Inserm U970, Paris Cardiovascular Research
Center, Paris, France

Part I
Development

1

Embryonic Development of the Kidney
Carlton Bates, Jacqueline Ho, and Sunder Sims-Lucas

Contents
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

4

Studying the Kidney and Urinary Tract
Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

5

Development of the Metanephros . . . . . . . . . . . . . . . . . .

7

Nephron Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Specification of Nephron Progenitors/Cap
Mesenchyme . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Nephron Induction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Nephron Segmentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Glomerulogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Molecular Control of Podocyte Terminal
Differentiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Molecular Control of Glomerular Capillary Tuft
Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

8
10
11
12
12
13
13

Renal Stroma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

Vascular Development of the Kidney . . . . . . . . . . . . . .
Angiogenesis Versus Vasculogenesis . . . . . . . . . . . . . . . . .
Origins of the Peritubular Capillary Endothelia . . . . . .
Molecular Control of Renal Vascular
Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Collecting System Development . . . . . . . . . . . . . . . . . . . .
Ureteric Bud Induction and Outgrowth . . . . . . . . . . . . . . .
Renal Branching Morphogenesis . . . . . . . . . . . . . . . . . . . . .
Patterning of the Medullary and Cortical Collecting
Ducts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Lower Urinary Tract Development . . . . . . . . . . . . . . . .
Anatomic and Functional Development . . . . . . . . . . . . . .
Molecular Control of Ureter and Bladder
Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Bladder . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Ureter–Bladder Anastomosis . . . . . . . . . . . . . . . . . . . . . . . . .

14
15
16
17
17
17
19
20
21
21
23
25
25

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26

C. Bates (*) • J. Ho • S. Sims-Lucas
Department of Pediatrics, Division of Pediatric
Nephrology, Children’s Hospital of Pittsburgh of UPMC,
University of Pittsburgh School of Medicine, Pittsburgh,
PA, USA
e-mail: batescm@upmc.edu; jacqueline.ho2@chp.edu;
sunder.sims-lucas@chp.edu
# Springer-Verlag Berlin Heidelberg 2016
E.D. Avner et al. (eds.), Pediatric Nephrology,
DOI 10.1007/978-3-662-43596-0_1

3

4

Introduction
The mammalian kidney functions as a key regulator of water balance, acid–base homeostasis,
maintenance of electrolytes, and waste excretion.
The performance of these activities depends on
the development of specific cell types in a precise
temporal and spatial pattern, to produce a sufficient number of nephrons. Over the past several
decades, considerable advances have been made
in understanding the molecular basis for this
developmental program. Defects in this program
result in congenital anomalies of the kidney and
urinary tract, which are the leading causes of
chronic kidney disease and renal failure in children. These developmental disorders range from
renal malformations, such as renal aplasia
(absence of the kidney), dysplasia (failure of normal renal differentiation), and hypoplasia (smaller
kidneys), to urinary tract abnormalities such as
vesicoureteral reflux and duplicated collecting
systems. This chapter describes the embryology
of the kidney and urinary tract, as a means to
understand the developmental origins of these
disorders.
Human kidney development starts in the fifth
week of gestation, and new nephrons are formed
until approximately 32–34 weeks gestation
[1–3]. Remarkably, nephron endowment is quite
variable ranging from 200,000 to 1.8 million
nephrons per person [4]. While the human kidney
continues to grow after 34 weeks gestation, this
occurs due to the growth and maturation of
existing nephrons, rather than the formation of
new nephrons. The mature mammalian kidney
cannot compensate for nephron loss due to renal
injury by the de novo generation of nephrons
[2, 5]. Therefore, the number of nephrons present
at birth in an individual is an important determinant of long-term kidney health. Therefore,
reduced nephron number is associated with hypertension and chronic kidney disease [6, 7].
Critical determinants of nephron endowment
are structural development and three-dimensional
nephron patterning. The formation of kidneys in
utero involves the coordinated regulation of
critical developmental processes: differentiation,

C. Bates et al.

morphogenesis, and regulation of cell number.
Differentiation is the process by which precursor
cells or tissues mature into more specialized cells.
During kidney development, renal mesenchymal
cells have the potential to differentiate into
nephron epithelia or stromal cells and interstitial
fibroblasts. Morphogenesis describes the
process whereby cells and tissues acquire threedimensional patterns. This is particularly important in the kidney, as the three-dimensional relationship between the nephrons, the vasculature,
and the collecting system is critical for
normal kidney structural function. Finally, the
regulation of cell number at different stages of
development is crucial. Such regulation maintains
a balance between cellular proliferation and
programmed cell death or apoptosis. All of these
processes are integrated tightly and regulated both
spatially and temporally in normal renal
development.
The molecular control of these developmental
processes has been the subject of several recent
comprehensive reviews [8–14]. Genetic, epigenetic, and environmental factors all regulate differentiation, morphogenesis, and cell number
within the developing kidney. Mutational analyses in animal models have provided significant
insights into the genetic control of renal development. Genes critical for kidney development in
animal models include transcription factors that
act as master regulators of other genes, growth
factors that signal to other cells, and adhesion
molecules that regulate how cells interact with
each other and with the extracellular matrix.
Increasingly, analyses of humans with congenital
renal malformations (such as renal aplasia or
duplex kidneys) have identified gene mutations
originally described in animal models [15]. Recent
studies have also implicated epigenetic mechanisms (defined as heritable changes in gene activity that are not caused by changes in DNA
sequence) in regulating nephron formation. Two
major classes of epigenetic molecules that appear
to regulate renal development include chromatin
remodeling proteins and regulatory RNAs such as
miRNAs [16–22]. Finally, environmental influences that may interact with genetic and

1

Embryonic Development of the Kidney

epigenetic factors are also important in determining nephron number and patterning. For example,
vitamin A deficiency leads to decreased nephron
number in rodents and has been implicated in
decreased renal size in humans [23, 24].

Studying the Kidney and Urinary Tract
Development
The methods for studying the molecular and
genetic control of kidney development have continued to evolve over the past several decades.
Visualization of tissue morphology and expression of individual genes and proteins in the developing kidney have traditionally been performed
on tissue sections by general staining (e.g., hematoxylin and eosin), detecting messenger RNA
(mRNA) via in situ hybridization or protein via
immunohistochemistry (Fig. 1a–c). The advent of
high-throughput technologies to detect gene
expression with microarrays and more recently
by high-throughput RNA sequencing has resulted
in the ability to assay the transcriptome of the
developing kidney and/or different kidney tissue
compartments in an unbiased fashion. These techniques have led to large public databases that
describe the gene expression of the developing
kidney, including the GenitoUrinary Development Molecular Anatomy Project (URL: www.
gudmap.org) and Eurexpress (URL: www.
eurexpress.org) [25–27].
A classical technique to analyze kidney development is to culture rodent embryonic kidneys
in vitro as explants. Studies using these methods
were the first to show that reciprocal interactions
between the metanephric mesenchyme and the
ureteric bud are critical to induce the formation
of new nephrons and ureteric branching (Fig. 1d)
[28]. Moreover, kidney explants still allow one to
modulate the expression and function of specific
genes and proteins using reagents such as antisense oligonucleotides or blocking antibodies
[29, 30]. While these experiments have been illuminating, the growth of embryonic kidney
explants differs from kidney development
in vivo in several key ways: lack of blood flow,

5

growth limitations from diffusion of the culture
media across the air–media interface, and distortions in the three-dimensional kidney architecture
as explants flatten in culture.
Recently, several methods to generate more
physiological and quantifiable three-dimensional
reconstructions of developing kidneys and urinary
tracts have been developed. One method utilizes
exhaustive serial sectioning through developing
kidneys, histological staining, projection of each
serial image onto a monitor to identify each tissue
lineage, and rendering of the serial images into a
three-dimensional image (Fig. 1e) [31, 32]. This
technique allows for the quantification of both
developing nephron structures and the branching
ureteric tree. Another method uses optical projection tomography to image through the full thickness of a developing kidney that has been whole
mount stained for a specific tissue and also permits the quantification of these elements [33].
Physical, chemical, and genetic strategies can
be used to manipulate developing kidneys in vivo.
For example, ureteric obstruction in utero in sheep
and monkeys results in hydronephrotic kidneys
with renal dysplasia [34, 35]. In addition, dietary
manipulations including high doses of vitamin A or
dietary protein restriction result in kidney and urinary tract defects [36, 37]. Transgenic approaches
have also been used to drive gene expression in
specific spatiotemporal patterns, usually in the
mouse. In these experiments, a transgenic construct
consisting of a tissue-specific promoter and the
gene of interest is randomly inserted into the
genome, leading to expression of that gene in a
tissue-specific pattern. The limitations of this
approach include: (1) that the random insertion
can result in unintended changes in gene expression (due to other nearby promoters/enhancers near
the site of integration), (2) that the insertion of the
transgene into the genome may lead to loss of
function of an endogenous gene, and that (3) epigenetic factors may silence the construct. The
increased utilization of bacterial artificial chromosome (BAC) constructs, which contain more of
endogenous promoter elements than are found in
traditional plasmid constructs, has led to more
faithful and reliable transgene expression.

6

Fig. 1 Experimental methods utilized to study kidney
development. (a) Hematoxylin and eosin (H&E)-stained
tissue section of a control postnatal day 0 mouse kidney.
The ureteric bud is outlined in yellow, and the arrow points
to the cap metanephric mesenchyme. (b) In situ hybridization in a control mouse embryonic day 16.5 tissue section
for the transcription factor, Wt1, which stains the metanephric mesenchyme and developing glomeruli. (c) Immunofluorescent staining in a control embryonic day 14.5
mouse tissue section for the transcription factor, Wt1
(red), and lotus tetragonolobus lectin (LTL, green), which
stains the proximal tubule. (d) Embryonic culture of a
transgenic embryonic day 11.5 HoxB7GFP mouse kidney,

C. Bates et al.

demonstrating branching ureteric structures (green) after
5 days of growth. (e) 3D reconstruction of an embryonic
day 13.5 mouse kidney with the ureteric epithelium
depicted in pink and developing nephron types including
renal vesicles (blue), comma-shaped bodies (red),
S-shaped bodies (purple), and glomeruli (green)
(Reproduced with kind permission from Springer Science
+Business Media: Sims-Lucas S. Kidney Development:
Methods and Protocols, Methods in Molecular Biology,
vol. 886, 2012, pp 81, Figure 3F) (f) H&E-stained tissue
section of a postnatal day 0 mouse kidney lacking
microRNAs in the ureteric lineage using a conditional
knockout approach (HoxB7Cre; Dicerflx/flx)

1

Embryonic Development of the Kidney

As opposed to transgenic approaches, homologous recombination is the method whereby a
gene is “knocked-out of” or “knocked-into” the
mouse genome. Using these methods, a gene of
interest is deleted so that it becomes
nonfunctional, or a gene (such as a green fluorescent reporter) is added to the genome at a specific
locus. A limitation to traditional knockout techniques is that global loss of function of the gene
may result in extrarenal effects (such as early
embryonic lethality), which can impact or
severely limit the study of the gene’s function in
the kidney. Given these limitations, it has become
more common to perform conditional gene
targeting (e.g., with the Cre–loxP system)
(Fig. 1f) and/or inducible gene targeting (e.g.,
with tamoxifen), allowing for kidney- and/or urinary tract-specific gene deletion (using a kidney
zebrafish specific Cre-) and/or at a particular time
(driving induction of gene targeting with a drug).
While most investigators using genetically
modified animals utilize mice, a growing number
of scientists study kidney development in other
model systems, such as avians, zebrafish, and
Xenopus. These simple systems are obviously
limited by their inability to recapitulate the complex regulation of development previously
described in three-dimensional mammalian kidney formation. However, their simplicity has
advantages in isolating possible molecular pathways involved in specific renal developmental
processes. These systems produce larger number
of embryos in a shorter period of time than in
mammals. Many of these models, such as
zebrafish and Xenopus, have only a
one-dimensional pronephric tubule as a kidney.
However, many of the genes which pattern such
simple structures have important roles in mammalian metanephric kidney development. Increasingly, investigators using have utilized more
sophisticated techniques such as transgenic fish
to examine the roles of genes or drugs in modifying nephron progenitor populations [38]. Finally,
the advent of clustered regularly interspaced short
palindromic
repeats
(CRISPR)/CRISPRassociated protein (Cas) techniques should eventually allow relatively quick and easy genetic
modifications of any animal model desired [39].

7

Development of the Metanephros
The mesoderm forms as one of the three embryonic
germ layers during gastrulation. The mammalian
kidney develops from the intermediate mesoderm,
lying between the somites and lateral plate mesoderm, on the posterior abdominal wall of the developing embryo. In mammals, three pairs of
embryonic kidneys develop from the intermediate
mesoderm: the pronephros, the mesonephros, and
the metanephros (Fig. 2). At their maximal development, the pronephros and mesonephros extend
from the cervical to the lumbar levels of the developing embryo. The pronephric and mesonephric
nephrons are induced to differentiate by signals
from the adjacent pronephric/mesonephric ducts,
paired epithelial tubules running in a longitudinal
course along the embryo on either side of the
midline. The mesonephric duct (also known as
the Wolffian duct) continues to grow caudally in
the embryos to eventually fuse with the cloaca,
which eventually gives rise to the bladder. The
pronephros is not functional in mammals, but is
the functional kidney in larval fish [40] and frogs
[41]. The mesonephros becomes the mature kidney
in these lower species and is functional in mammals during embryogenesis. Ultimately, the pronephros and mesonephros largely degenerate in
mammals. Portions of the mesonephros and mesonephric duct persist in mammals as the rete testis,
efferent ducts, epididymis, vas deferens, seminal
vesicle, and prostate in males [42]. In mammalian
females, remnants of the mesonephric tubules persist as the epoophroron and paroophoron.
The mature mammalian kidney, the metanephros, is derived from two tissues, the ureteric bud
and metanephric mesenchyme (see a recent
review in Ref. [13]). Starting at approximately
embryonic day 10.5 in the mouse and the 5th
week of gestation in humans, reciprocal inductive
signals cause the ureteric bud to grow out from the
caudal portion of the mesonephric duct and the
metanephric mesenchyme to condense around the
ureteric bud to form nephron progenitors. The
ureteric bud ultimately gives rise to the collecting
system, including the collecting ducts, renal
calyces, renal pelvis, and ureters [2, 43]. In turn,
the nephrogenic metanephric mesenchyme

8

Fig. 2 Schematic overview of kidney development.
Mammalian kidney development begins with the formation of the nephric duct, which is divided into three segments: pronephros, mesonephros, and metanephros. The
pronephros degenerates in mammals, whereas the mesonephros forms the male reproductive organs (rete testis,
efferent ducts, epididymis, vas deferens, seminal vesicles,
and prostate). The metanephros becomes the mature mammalian kidney and is derived from inductive interactions
between the metanephric mesenchyme and the ureteric bud
(Reproduced with kind permission from Springer Science
+Business Media: Moritz K, et al. Factors Influencing
Mammalian Kidney Development: Implications for Health
in Adult Life, Morphological Development of the Kidney,
Advances in Anatomy and Cell Biology, volume
196, 2008, pp 9–16, figure number 1)

C. Bates et al.

limbs of the loops of Henle, and the distal convoluted tubule [2, 43]. Just after the condensation of
nephrogenic mesenchyme around the ureteric
bud, stromal metanephric mesenchyme (or renal
stroma) develops adjacent to the nephrogenic
mesenchyme. The renal stroma develops into
perivascular cells, vascular smooth muscle, fibroblasts, mesangial cells, renin-producing cells, and
even some peritubular endothelial cells (see
below).
The transcription factor, Odd-skipped related 1
(Osr1), is one of the several key molecules necessary to specify portions of the posterior intermediate
mesoderm to become the mesonephric duct, ureteric
bud, and metanephric mesenchyme (nephrogenic
and stromal) [44]. Osr1-expressing cells in the intermediate mesoderm have been shown to give rise to
most of the cellular components of the metanephric
kidney, including the nephron, vascular cells, interstitial cells, and the mesonephric duct (including its
derivatives, viz., the ureteric bud/collecting system)
[45]. Other molecules critical for specification and
development of the mesonephric duct include the
transcription factors Paired box 2 (Pax2) [46], Pax8
[47], Lim homeobox 1 (Lhx1) [48], Gata binding
protein 3 (Gata3) [49], and the receptor tyrosine
kinase, Ret [50].
Many key signaling pathways have been shown
to drive reciprocal interactions between the ureteric
epithelium, the renal mesenchymal lineages, and
the renal vasculature. Ureteric bud outgrowth
depends on inductive signals from nephron progenitors [51–53], stromal cells [54–58], and
angioblasts [59, 60], as well as from itself
[61]. The nephrogenic mesenchyme relies in part
on signaling from the ureteric bud and renal stroma
for self-renewal and for initiation of nephron formation [8, 11, 62–64]. Subsequent nephrogenesis
(i.e., patterning and differentiation of nephron epithelia) is highly dependent on factors from both
ureteric epithelial and stromal cells [57, 65, 66].

Nephron Formation
differentiates into the epithelial cells that comprise
the mature nephron, including the parietal cells
and podocytes of the glomerulus, the proximal
convoluted tubule, the ascending and descending

The metanephric mesenchyme gives rise to
nephrogenic mesenchyme or nephron progenitors, which self-renew and have the capacity to

1

Embryonic Development of the Kidney

9

form the multiple epithelial cell types of the nephron. Much research is geared toward understanding this progenitor cell population, which is
critical for determining nephron endowment and
thus long-term kidney health.
Anatomically, there are several steps that take
place in nephron formation. After the ureteric bud
has penetrated the metanephric mesenchyme,
nephron progenitors condense around the first
ureteric ampulla, forming “cap mesenchyme.”
As the ureteric bud continues to branch and elongate, the nephrogenic mesenchyme continues to
form new caps surrounding each ureteric tip. After
the initial few ureteric branches, the earliest cap
mesenchymal cells receive spatiotemporal cues to
begin the differentiation process to form
epithelialized renal vesicles [67]. Subsequent
growth and differentiation of the renal vesicle
results in formation of the comma-shaped body,
which then lengthens to form the S-shaped body.
The lower limb of the S-shaped body begins to
differentiate into glomerular podocytes. During
this time, endothelial cells migrate into the cleft
of the lower limb of the S-shaped body and will
ultimately form the glomerular capillary loops
[68, 69]. Simultaneously, nascent mesangial
cells, derived from renal stroma (see below), also
migrate into this cleft. Thus, the lower limb of the
S-shaped body forms the immature glomerulus.

Concurrently, the middle and upper limbs of the
S-shaped body elongate and differentiate into
nephron tubules including proximal tubules,
loops of Henle (including descending and ascending limbs), and distal convoluted tubules. The
terminal ends of the distal convoluted tubules
eventually connect to ureteric epithelia, which
ultimately form the collecting system (collecting
ducts, renal pelvis, and ureters) (Fig. 3).
Nephrogenesis repeats in a radial fashion with
the first nephrons forming in the juxtamedullary
regions and last in the peripheral cortex, until the
full complement of nephrons is reached.
During prenatal and/or postnatal life, each
nephron increases in size and complexity as it
matures. Starting in the first month of life, maturing proximal tubules transition from a columnar to
cuboidal epithelium, develop microvilli, and
increase their tubular dimensions [70, 71]. While
the earliest limbs of the Henle loop are located in
the renal cortex, subsequent maturation and elongation of these limbs in utero results in the loops
pushing through the corticomedullary boundary
in term infants [72–74]. Postnatal maturation
results in the Henle loops eventually reaching
the inner renal medulla in the mature kidney.
Thus, the urinary concentrating capacity of newborn infants is limited by a reduced medullary
tonicity gradient, due to the relatively shorter

Fig. 3 H&E-stained sections showing the four stages of
nephron formation in mice. (a) Image of a renal vesicle,
the first stage of nephron formation. (b) Image of a commashaped body that has differentiated from a vesicle. (c)
Image of an S-shaped body, the third stage of nephron
formation. (d) Image of an immature glomerulus that

differentiated from the lower limb of the S-shaped body
(Reproduced with kind permission from Springer Science
+Business Media: Sims-Lucas S. Analysis of 3D
Branching Pattern: Hematoxylin and Eosin Method,
Methods in Molecular Biology, volume 886, pp 73–86,
2012, Figure 3, panels A–D)

10

loops of Henle. Finally, as the distal convoluted
tubule matures, a portion of the cells are found in
close proximity to the future vascular pole of the
developing glomerulus, where they develop into
the macula densa [72].

Specification of Nephron Progenitors/
Cap Mesenchyme
Differentiation of the intermediate mesoderm and
metanephric mesenchyme into nephron progenitors and their derivatives is genetically defined by
the sequential upregulation of several transcription factors, cell adhesion molecules, and growth
factors. The intermediate mesoderm and early
metanephric mesenchyme express the transcription factors Sal-like 1 (Sall1) [75], Sine oculis
homeobox homolog 1 (Six1) [76], Eyes absent
homolog 1 (Eya1) [77, 78], and the secreted peptide growth factor, Glial-derived neurotrophic
factor (Gdnf) [79]. Induction of cap mesenchyme/nephron progenitors by the ureteric bud
tips is marked by expression of transcription factors such as Wilms tumor 1 (Wt1) [80]; Cbp/p300interacting transactivator, with Glu/Asp-rich
carboxy-terminal domain, 1 (Cited1) [81]; and
Sine oculis homeobox homolog 2 (Six2) [82], as
well as the transmembrane molecules cadherin-11
[83] and α8 integrin [84]. The earliest epithelial
derivative of the cap mesenchyme, the renal vesicle, is marked by the transcription factors Pax2
[85] and Lhx1 [86].
The use of tissue-specific gene knockouts generally via conditional transgenic techniques has
revealed the importance of several transcription
factors (including many mentioned above) in the
specification of the cap mesenchyme. Conditional
homozygous deletion of Eya1 [78], Six1 [76],
Pax2 [87, 88], Wt1 [80], Sall1 [75], Six2 [82], or
Lhx1 [86, 89] leads to bilateral renal aplasia or
severe renal dysgenesis from defects in cap mesenchyme specification and/or differentiation;
these mesenchymal defects are often accompanied by ureteric induction and/or branching
abnormalities due in large part to loss of GDNF
signaling from the metanephric mesenchyme.
Pax2 mutant mice generate a metanephric

C. Bates et al.

mesenchyme that is unable to differentiate into
nephrons and fail to form the mesonephric duct,
which is required for ureteric bud induction
[88]. In Lhx1 [86, 89] and Sall1 [75] mutants,
the metanephric mesenchyme does induce ureteric bud formation, but it fails to elongate and
branch, and the mesenchyme is once again unable
to differentiate into nephrons. In Wt1 mutants, a
defective metanephric mesenchyme is formed,
but rapidly undergoes apoptosis [80]. Six2, a specific marker of nephron progenitors, is required
for the maintenance of progenitor cells, but not for
nephron differentiation; deletion of Six2 in mice
results in the formation of ectopic nephron tubules
and the rapid depletion of nephron progenitor
cells [82]. Similarly, the p53–E3 ubiquitin ligase,
murine double minute 2 (Mdm2), appears critical
for the maintenance of nephron progenitor
cells [91].
Recent studies have identified factors that govern the delicate balance of self-renewal and differentiation of nephron progenitors, including
WNT genes. Studies from the mid-1990s showed
that lithium chloride, a potent inducer of Wnt
signaling, was able to drive tubulogenesis in isolated rodent metanephric mesenchyme cultures
[92–94]. More recently Wnt9b, secreted from ureteric bud cells, was shown to be required for the
differentiation of nephron progenitor cells.
Genetic deletion of Wnt9b resulted in a failure of
nephron progenitors to undergo the mesenchymal
to epithelial transition that is required to form the
renal vesicle [95]. A subsequent study revealed
that Wnt9b, along with signals from the renal
stroma (see below), plays an essential role in
mediating the decision of nephron progenitors to
self-renew or differentiate [64, 96].
Another major signaling pathway that has been
shown to mediate nephron progenitor survival is
the fibroblast growth factor (FGF) signaling pathway. FGF ligands are secreted peptides that bind
and signal through their receptor tyrosine kinases,
FGF receptors (FGFRs). Isolated nephrogenic
zone cell culture studies revealed that addition of
FGF1, 2, 9, and 20 ligands drives the expression
of nephron progenitor markers and progenitor
proliferation [97]. More recently, in vivo mouse
studies showed that Fgf9 and Fgf20 are critical for

1

Embryonic Development of the Kidney

maintaining nephron progenitor survival, proliferation, and competence to respond to inductive
signals; furthermore, FGF20 mutations in humans
were shown to be associated with severe renal
dysplasia [98]. Other work in mice has identified
that the Fgfrs critical for metanephric mesenchyme development are Fgfr1 and Fgfr2. Conditional deletion of Fgfr1 and Fgfr2 in the
metanephric mesenchyme leads to severe renal
dysgenesis [99–101].
Several studies have shown that a balance
between cell survival and apoptosis is necessary
for normal nephron progenitor function. Classic
in vitro studies using isolated metanephric mesenchyme have shown that nephron progenitors
undergo massive apoptosis when cultured without
an inducer [102]. Coculture with isolated ureteric
buds or heterologous inducers, such as embryonic
spinal cords, dampens apoptosis and drives progenitor survival [43, 52]. Other studies have identified several factors that when added to
metanephric mesenchyme or nephrogenic zone
cell cultures drive progenitor survival, including
transforming growth factor-β2 (TGF-β2), TGFα,
leukemia inhibitory factor (LIF), epidermal
growth factor (EGF), FGF2, and bone morphogenetic protein 7 (BMP7) [62, 103–105]. Whether
some or all of these factors act as endogenous
nephron progenitor inducers remains to be determined. Counterbalancing cell survival, apoptosis
is required for normal nephron progenitor function. Moreover, suppression of apoptosis via pharmaceutical or genetic means leads to kidney
malformations including abnormal ureteric
branching and defective nephrogenesis [106,
107]. Relative “overabundance” of nephron progenitors also leads to epithelial and/or stromal cell
defects [108, 109].
Recent studies have revealed the importance of
epigenetic mechanisms that regulate nephron progenitor specification, survival, and potential for
differentiation. Histone deacetylases (HDACs)
are enzymes that remove acetyl groups from histones, which then modulate (usually stimulate)
gene transcription. Recent work has revealed
that Class I HDACs are highly expressed in nephron progenitors and are required for the proper
expression of several key developmental genes

11

including Osr1, Eya1, Pax2, Wt1, and Wnt9b
(among others) [16]. MicroRNAs (miRNAs) are
small noncoding RNAs that bind to specific
mRNA targets to block translation and promote
mRNA degradation. Conditional targeting of
dicer, an enzyme required for the processing of
all miRNAs, in mouse nephron progenitors, led to
a loss of the progenitors due to excessive apoptosis, likely from upregulation of the proapoptotic
protein, Bim [22]. A more recent study revealed
that conditional deletion of a specific miRNA
cluster, the miR-17~92 complex, in nephron progenitors led to decreases in progenitor proliferation, fewer numbers of nephrons, proteinuria, and
podocyte damage. Moreover, this report was the
first to identify a specific miRNA cluster essential
for kidney development [110].

Nephron Induction
The initial differentiation step of nephron progenitors is to undergo a mesenchymal to epithelial
transition to form the renal vesicle. In vitro experiments utilizing isolated rodent metanephric mesenchymal rudiments (similar to and including
some of the survival studies noted above) have
identified exogenous factors that stimulate nephron progenitors to undergo tubulo-epithelial differentiation [52]. Some of these growth factors
can act alone or in concert with others and include
FGF2 [111], LIF [63, 105, 112], TGFβ2 [105,
113], growth/differentiation factor-11 (GDF-11)
[105, 114], and WNT1/4 [115, 116]. Sequestration
of Wnt ligands in intact rodent kidney explants by
addition of secreted frizzled-related proteins
(sFrps) leads to decreases in mesenchyme-derived
tubulogenesis [117]. More recent mouse genetic
experiments have identified at least some of the
critical endogenous pathways and growth factors
necessary for the induction of the mesenchymal to
epithelial transition. For example, global deletion
of Wnt4, normally expressed in renal vesicles, did
not perturb cap mesenchyme formation; however,
the mutant nephron progenitors were completely
unable to form renal vesicles [95]. Conditional
deletion of Fgf8 in the metanephric mesenchyme
leads to a block of nephrogenesis beyond the renal

12

vesicle stage. Fgf8 likely normally acts with Wnt4
to drive Lhx1 expression [118, 119]. Interestingly,
global deletion of Fgfr-like 1, a membrane-bound
Fgf receptor that lacks an intracellular tyrosine
kinase domain, also leads to a block in
nephrogenesis similar to Fgf8 conditional
mutants [120].

C. Bates et al.

glomeruli and proximal tubules [127]. In vivo,
conditional deletion of Notch2 or Rbpsuh in
mouse metanephric mesenchyme leads to an
absence of proximal tubules and glomerular epithelium [128]. Finally, ectopic Notch expression
in nephron progenitor cells results in the premature differentiation of the progenitors into proximal nephron epithelia [130].

Nephron Segmentation
Glomerulogenesis
Establishment of a proper proximal–distal axis is
critical for normal nephron segmentation. Negative reciprocal interactions between Wt1 and
Pax2 at early stages of nephron development
appear vital to proximal–distal axis patterning
[121–123]. In the S-shaped body, Wt1 is localized
to the lower limb and inhibits Pax2 expression,
which together drives cells toward podocyte fates
[124]. Transgenic mice with overexpression of
Pax2 throughout the embryo including developing nephrons develop glomerular defects and
renal cystic dysplasia [125]. In contrast Pax2 is
expressed in the upper limb of the S-shaped body
and represses Wt1 expression, stimulating these
cells to become proximal and distal tubular nephron segments [118, 126].
Two other transcription factors critical for
proximal–distal axis patterning of the nephron
include Lhx1 and Brain specific homeobox 1
(Brn1), both of which are expressed at the renal
vesicle stage. Conditional deletion of Lhx1
throughout the metanephric mesenchyme blocks
nephrogenesis at the renal vesicle stage and also
leads to a loss of Brn1 expression [86]. Conditional targeting of Brn1 in the metanephric mesenchyme does not block proximal nephrogenesis;
however, the loop of Henle fails to form, and
distal convoluted tubules fail to terminally differentiate [73]. These results suggest that Lhx1 acts
earlier in nephron patterning than Brn1, which is a
critical distal nephron patterning.
Notch receptor signaling, mediated largely by
Recombining binding protein suppressor of hairless (Rbpsuh), appears critical for proximal nephron patterning [127–129]. Use of a Notch
inhibitor in mouse metanephric kidney explants
led to a loss of proximal cell fates, including

Glomerulogenesis is initiated when the commashaped body differentiates into the S-shaped body
[2, 131]. During this time, immature podocytes
along the lower limb are highly proliferative and
have a columnar shape with apical cell attachments and a single-layer basement membrane
[131]. Concurrently, endothelial and mesangial
cell progenitors are recruited into the lower cleft
of the S-shaped body, which will become the
vascular pole [132]. While mesangial cells originate from the renal stroma (see below), the developmental origin of the glomerular endothelium is
still unclear. Transplantation of avascular rodent
embryonic kidney rudiments under neonatal kidney capsule led to the formation of endothelial
precursors or angioblasts originating from the
graft metanephric mesenchyme [132–134]. However, engraftment of embryonic rat kidney rudiments onto avian chorioallantoic membrane led to
vascular ingrowth of avian vessels into the rat
glomeruli [135]. As will be expanded on in the
Vascular Development section below, it is likely
that both processes/sources contribute to the formation of glomerular capillaries. As the S-shaped
body matures, the lower cleft transforms into a
cup shape configuration.
At this time the podocytes lose their proliferative ability [136] and differentiate, forming foot
processes and slit diaphragms, specialized intracellular junctions critical for proper glomerular
filtration [137, 138]. Concurrently, the composition of the glomerular basement membrane
changes from laminin-1 to laminin-11 and from
α -1 and α -2 type IV collagen chains to α -3, α -4,
and α -5 type IV collagen chains [139]. Several
mouse knockout mice models have shown how

1

Embryonic Development of the Kidney

failure in these transitions leads to structural and
functional glomerular basement membrane
defects [140–142]. At this stage of development,
the nascent mesangial cells act as a scaffold for the
formation of the glomerular capillary loops and
ultimately form the supportive core for the entire
glomerulus via the deposition of extracellular
matrix [143, 144]. Developing glomerular endothelial cells branch extensively during this time
and begin differentiating into fenestrated endothelia [2] (see Vascular section below).
By 32–34 weeks gestation, glomerulogenesis/
nephrogenesis ceases in humans, whereas it persists in mice and rats for 7–10 days following full
gestation [2]. In newborn humans, the superficial
glomeruli are the most immature and are smaller
than the deeper juxtamedullary glomeruli
[71]. While no new glomeruli are formed after
birth, they continue to grow and mature postnatally, reaching their adult size at approximately
three and a half years of age [71].

Molecular Control of Podocyte
Terminal Differentiation
Transcription factors and epigenetic factors,
including microRNAs, have been shown to be
critical for podocyte differentiation. Examples of
essential transcription factors include Wt1,
podocyte expressed 1 (Pod1), Lim homeobox 1b
(Lmx1b), and Mafb. Several studies utilizing
murine genetic knockout models of Wt1 have
demonstrated its critical roles in mediating
podocyte differentiation [145–148]. In humans,
WT1 mutations can lead to diffuse mesangial sclerosis, characterized by podocyte differentiation
defects resulting in varied glomerular lesions and
proteinuria, and can occur as an isolated disease or
in association with Denys–Drash or Frasier syndromes [149–152]. Deletion of Pod1, expressed
in stromal cells, leads to nonautonomous
podocyte defects at the capillary loop stage in
mice [153]. Genetic deletion of Lmx1b or Mafb
leads to podocyte differentiation defects past the
capillary loop stage [154, 155]. Furthermore,
mutations in the LMX1b gene in humans lead to
nail–patella syndrome, which is often associated

13

with glomerular basement membrane thickening
and proteinuria that can progress to chronic kidney disease [156, 157]. Three studies recently
showed the importance of microRNAs in
maintaining differentiated podocytes in the
mouse. Targeted ablation of dicer in murine
podocytes, resulting in a loss of all miRNAs, led
to podocyte injury, severe proteinuria, and tubular
damage starting 2 weeks after birth [20, 158, 159].

Molecular Control of Glomerular
Capillary Tuft Development
There are several signaling cascades that have
been implicated in the homing and maturation of
the endothelial and mesangial precursors to form
the glomerular capillary tuft. Vascular endothelial
growth factor (VEGF), which is secreted from the
podocytes at the S-shaped body stage, promotes
recruitment of endothelial precursors to the vascular cleft [160, 161]. Angiopoietin-1 and -2,
growth factors expressed by podocytes and
mesangial cells, respectively, are also critical for
normal glomerular capillary development
[162]. Mesangial cell recruitment into the cleft is
largely mediated by the secretion of plateletderived growth factor (PDGF)-B by endothelial
cells, which binds PDGF receptor-β (PDGFRβ)
on the mesangial cell progenitors [163]. Mice that
lack either Pdgfβ or Pdgfrβ fail to form glomerular capillary tufts demonstrating the importance of
mesangial cell recruitment [164, 165]. Finally,
Notch2 and its ligand Jagged1 are critical for
glomerular endothelial and mesangial cell development. Notch2 hypomorphic mice and Notch2/
Jagged1 compound heterozygous mice develop
glomerular aneurysms and possess no mesangial
cells [166].

Renal Stroma
The renal stroma, like the nephrogenic mesenchyme, is derived from Osr1-positive intermediate mesoderm and the metanephric mesenchyme
[167, 168]. A hallmark of the initial renal stroma
is the expression of the transcription factor,

14

Foxd1, which is seen as early as E11.5 in the
mouse. The renal stroma is initially located at
the periphery of the kidney and interdigitates
between the developing nephron units and ureteric tips. One function of the early renal stroma
is to support framework for the developing vessels, nephron progenitors, and ureteric epithelia.
As embryonic kidney development progresses,
stromal cells are present in both the peripheral
renal cortex and the medulla surrounding developing collecting ducts. At this time, the cortical
stroma expresses Foxd1, Aldehyde dehydrogenase 1 family, member A2 (Raldh2), Retinoic
acid receptor α (Rarα), and Rarβ2, while the
medullary stroma expresses Fgf7, Pod1, and
Bmp4. Many of these stromally expressed genes
have been shown to be critical for nephrogenesis
and ureteric branching morphogenesis by virtue
of mouse knockout studies [54, 55, 57, 58, 153,
169]. At birth, many of the developmental stromal
cells have undergone apoptosis and are replaced
by nephron segments such as loops of Henle
[170]. Many stromal derivatives do survive giving
rise to fibroblasts, lymphocyte-like cells, glomerular mesangial cells, renin-expressing cells, vascular smooth muscle cells, pericytes, and a
subpopulation of peritubular endothelial cells
[168, 170, 171].
As noted, signaling from the renal stroma is
critical for ureteric morphogenesis. Three genes/
pathways expressed within the stroma, retinoic
acid, Foxd1, and Pod1, modulate ureteric
branching by regulating expression of Ret, a
receptor tyrosine kinase expressed in ureteric
tips and required for ureteric development (see
below). Vitamin A is converted to its active
form, retinoic acid, by the enzyme Raldh2 in the
renal stroma. Moreover, blockade of retinoic acid
signaling in mice, by deletion of Raldh2 or by
combined deletion of the retinoic acid receptors,
Rarα and Rarβ2, leads to hypoplastic kidneys
with a reduction in the number of ureteric
branches; the ureteric branching defects are linked
to the downregulation of Ret expression in mutant
embryos, which in the case of the retinoic acid
receptor mutants can be rescued by forced
re-expression of Ret in the ureteric tissues [55,
56, 172]. Foxd1 (expressed in cortical stroma

C. Bates et al.

and the renal capsule) or Pod1 (found in medullary stroma) appears to appropriately restrict Ret
expression to ureteric tips; genetic deletion of
either gene leads to mis-expression of Ret
throughout the entire ureteric tree and subsequent
ureteric branching defects [54, 57, 153, 173].
Cross talk from the stroma is also critical for
nephron development. Mouse genetic studies
show that Foxd1 and Pod1 are necessary for normal nephron patterning (in addition to ureteric
morphogenesis) [54, 153]. Loss of Foxd1 in
mice leads to premature differentiation of stromal
cells, which inhibits Bmp7-mediated nephron
progenitor differentiation [174]. Two recent
studies have shown how complete ablation of
renal cortical stroma with diphtheria toxin leads
to abnormally thickened nephron progenitor
caps and a decreased ability of progenitors to
differentiate [64, 175]. Mechanistically, it
appears that loss of the protocadherin Fat4 in the
stroma perturbs the activity of the transcription
factors, Yap and Taz, which in turn disrupts
Wnt9b signaling and nephron differentiation
[64]. Thus, in addition to providing a “framework” for the rest of the developing kidney, the
renal stroma actively signals to other renal lineages and differentiates into cells that populate the
mature kidney.

Vascular Development of the Kidney
The adult kidney receives approximately 25 % of
the cardiac output. Furthermore, the adult kidney
has a high complex vascular network with different functions and therefore different specialized
endothelia depending on location [176]. Specifically, three major types of endothelial cells are
present within the kidney, including fenestrated
(in glomerular capillaries), fenestrated with diaphragms (in peritubular capillaries and ascending
vasa recta), and continuous capillaries
(in descending vasa recta) (Fig. 4). Not surprisingly, these various endothelial cell types have
heterogeneous expression profiles and often
appear to have different developmental origins
[21, 177, 178].

1

Embryonic Development of the Kidney

15

Fig. 4 Electron microscopy demonstrating the varied
renal endothelium. (a, e) Glomerular capillaries contain
fenestrated endothelium without diaphragms (e, arrows)
and share a basement membrane (*) with podocyte foot
processes (large arrowhead) that are separated by slit
diaphragms (small arrowhead). (b, f, g) Peritubular capillaries have fenestrated endothelial cells that are covered
with diaphragms (f, g, arrows) and have a thick basement
membrane (*) separating them from the tubular cells.
(c, h, i) Ascending vasa recta (AVR) also have fenestrated

endothelium with diaphragms (c, h, i, arrow). (d, h, i)
Descending vasa recta (DVR) possess endothelium that is
non-fenestrated, thick, and continuous (d, h, i). RBC red
blood cell, EC endothelial cell. Panels (a–d) scanning
electron micrographs. Panels (e–i) transmission electron
micrographs (Reproduced with kind permission from
Springer Science+Business Media: Stolz DB and SimsLucas S. Unwrapping the origins and roles of the renal
endothelium, Pediatr Nephrol. 2015;30(6):865–72,
Figure 2)

Angiogenesis Versus Vasculogenesis

progenitors (marked by Flk1/Vegfr2) form primitive vascular networks, particularly within the
renal stroma, that subsequently join with and are
pruned by the angiogenic vessels [182]. The
vasculogenic endothelial cell progenitors within
the kidney appear to arise from the
Osr1-expressing intermediate mesoderm, as is
the case with the rest of the metanephric
kidney [45]. Finally, specification of the
endothelium
(whether
angiogenic
or
vasculogenic in origin), including arterial,
venous, capillary, or lymphatic fates, is driven
by growth factor signaling pathways and

Blood vessels can form by angiogenesis, in which
new vessels sprout from existing vessels, or by
vasculogenesis, in which de novo vessels form
from endothelial progenitors (Fig. 5). Extensive
linage tracing experiments and transplantation
studies have shown that both of likely occur in
renal vascular formation [176, 179–181].
The early renal artery and efferent arterioles
appear to be primary sites from which new
angiogenic vessels sprout within the developing
kidney. Simultaneously, renal endothelial

16

C. Bates et al.

Fig. 5 Schematic diagram of vascular formation in the
developing mouse kidney. Top panels. Angiogenic vessels (red) grow out from the major branches of the renal
artery and track with the branching ureteric epithelium
(orange). Middle panels. Vasculogenic vessels form from
progenitor cells (yellow, middle panel) within the metanephric mesenchyme (blue) and form a primitive vascular
plexus (yellow, right panel). Bottom panels. Schematic

diagrams depicting how a combination of angiogenesis
and vasculogenesis likely leads to vessel formation in the
kidney. E10.5–12.5 = embryonic days 10.5–12.5
(Reproduced with kind permission from Springer Science
+Business Media: Stolz DB and Sims-Lucas
S. Unwrapping the origins and roles of the renal endothelium, Pediatr Nephrol. 2015;30(6):865–72, Figure 1)

transcription factors, including Vegf, Ephrin,
Notch, and Sox [183].

peritubular capillaries has been less well defined.
Recent studies, however, have found that
peritubular capillaries arise from a combination
of resident endothelial progenitors as well as
invading angiogenic vessels [182, 184,
185]. One intriguing study found that Foxd1-positive renal cortical stroma cells give rise to a subset
of the peritubular endothelia but not the glomerular endothelia [184].

Origins of the Peritubular Capillary
Endothelia
While the formation of glomerular capillaries has
been extensively studied (see above), the origin of

1

Embryonic Development of the Kidney

Molecular Control of Renal Vascular
Development
A key signaling pathway mediating renal vascular
development is the VEGF pathway. VEGF
ligands are expressed early in the metanephric
mesenchyme and later in the developing glomerular podocytes, distal tubules, and collecting
ducts and at low levels in the proximal tubules
[68, 69]. Developing endothelial cells, including
those that arise from existing vessels and those
forming de novo, express VEGF receptors; thus,
VEGF signaling appears to drive both angiogenesis and vasculogenesis within the kidney. Interestingly, Vegfr2 is present on the apical surface of
ureteric epithelium, which likely accounts for
the stimulatory role of Vegf on ureteric growth
[132, 186].
Hypoxia-inducible factors (HIFs), a family of
transcriptions factors, are likely master regulators
of angiogenesis and vasculogenesis within the
developing kidney [187]. These molecules are
activated during periods of low oxygenation, as
occurs during embryogenesis, and are
downregulated postnatally. The HIF genes are
largely located in the nephrogenic zone, including
podocytes, developing collecting ducts, and
developing endothelial cells [187, 188]. HIF proteins induce expression of VEGF ligands, Vegfr1,
and Vegfr2 during kidney development by directly
binding to hypoxia-responsive elements on those
genes [189–192].
Angiopoietin (Ang) growth factors that bind to
Tie receptors also appear to have critical roles in
renal vascular development and are at least in part
regulated by HIF and VEGF signaling [193,
194]. Ang1, which is expressed in the metanephric mesenchyme, maturing nephron tubules, and
podocytes, signals through Tie2, which is
expressed on endothelial cells [195]. Conditional
deletion of Ang1 or Tie2 in mice leads to glomerular capillary defects including endothelial cells
that do not attach to the basement membrane [196,
197]. Ang2, expressed in vascular smooth muscle
cells and pericytes, binds to Tie1 that is expressed
by endothelial cells. Genetic deletion of Ang2
leads to upregulation of Tie2 signaling and significant defects in renal peritubular capillaries [198].

17

The Notch signaling pathway appears to regulate renal angiogenic vessel outgrowth [199].
Notch receptors induce sprouting by stimulating
the expression of Vegfr2 in vascular tip cells.
Simultaneously Notch inhibits Vegfr2 signaling
in adjacent vascular stalk cells, causing them to
remain dormant. Thus Notch regulates the pattern
of branching in angiogenic vessels.

Collecting System Development
Ureteric bud formation begins in the 5th week of
gestation in humans and at embryonic day 10.5 in
mice. As noted previously, signals from the metanephric mesenchyme cause the ureteric bud to
form from the mesonephric duct and then invade
the mesenchyme. Overall, collecting duct system
development includes (i) ureteric bud outgrowth,
(ii) branching of the ureteric bud, and (iii) patterning of the collecting duct system, all of which is
discussed in more detail below.

Ureteric Bud Induction and Outgrowth
Failure of ureteric bud outgrowth results in renal
aplasia, which can occur unilaterally or bilaterally
[200]. The GDNF–RET signaling pathway is crucial for bud outgrowth. The receptor tyrosine
kinase RET and its coreceptor GFRα1 are
expressed in the mesonephric duct, the initial ureteric bud, and later in the branching ureteric tips,
while its ligand, GDNF, is present in the metanephric mesenchyme (Fig. 6) [201–203]. Targeted
deletion of Gdnf, Ret, or Gfrα1 in mice generally
results in bilateral renal aplasia due to a lack of
ureteric bud outgrowth [202, 204–209]. Heterozygous mutations of RET have also been identified
in humans with bilateral renal aplasia, and a rare
RET polymorphism has been reported in individuals with nonsyndromic vesicoureteral reflux
[210, 211]. Moreover, the renal aplastic phenotype is not fully penetrant in a subset of Gdnf /
or Ret / mutant mice [202, 204], suggesting that
other molecular pathways play a role in ureteric
bud outgrowth. Some examples of other pathways
include signaling through integrins such as

18

C. Bates et al.

Fig. 6 Schematic diagram of the molecular control of
ureteric bud induction. GDNF is secreted from the metanephric mesenchyme and binds to its receptor, Ret (and its
coreceptor GFRα1) on the mesonephric duct, to induce
ureteric bud formation. Slit2/Robo2 and FoxC1 inhibit
the domain of GDNF expression and thus limit ureteric

bud induction to a single site from the mesonephric duct.
Sprouty1 (in the mesonephric duct) and Bmp4 (in tailbudderived mesenchyme around the mesonephric duct)
repress GDRF–Ret signaling and thus restrict ureteric
bud induction to its proper site

α8 integrin [84] and enzymes involved in proteoglycan synthesis such as heparan sulfate
2-sulfotransferase (Hs2st) [212].
Many studies have focused on the molecular
mechanisms that regulate GDNF–RET expression
and/or signaling. Prior to kidney development,
Ret is expressed throughout the mesonephric
duct, and Gdnf is present throughout the intermediate mesoderm adjacent to the mesonephric duct
[50, 201]. At the time of ureteric bud induction,
Gdnf expression becomes restricted to the posterior intermediate mesoderm next to the site of
ureteric bud outgrowth; once the ureteric bud has
invaded the mesenchyme, Ret expression
becomes restricted to ureteric bud tips [50]. In
vitro studies with Gdnf-soaked agarose beads
show that the entire length of the mesonephric
duct is competent to respond to Gdnf by initiating
ectopic ureteric bud formation [201, 213]. Moreover, mice that ectopically express Gdnf or Ret
in vivo develop renal malformations such as
duplex kidney and hydronephrosis [214,
215]. Together, these data show that GDNF–RET
signaling is highly spatially regulated for a single
ureteric bud to form in the correct location from
the mesonephric duct.

At least three genes, Foxc1, Slit2, and Robo2,
are thought to be crucial in restricting Gdnf to the
posterior intermediate mesoderm (Fig. 6). Homozygous mutant mice for all three genes develop
ectopic ureteric buds, multiple ureters,
hydroureter, and anterior expansion of Gdnf
expression [216, 217]. Foxc1 encodes a transcription factor co-expressed with Gdnf in the metanephric mesenchyme [216]. In the central nervous
system, the secreted protein Slit2 functions as a
chemorepellent during migration of axons that
express its receptor Robo2 [218, 219]. In the
developing kidney, Slit2 is expressed in the mesonephric duct, and Robo2 is detected in the metanephric mesenchyme [220]. ROBO2 missense
mutations in humans have been identified in families with vesicoureteral reflux and/or duplex
kidneys [221].
Two other genes, Sprouty1 (Spry1) and Bmp4,
act in a negative feedback loop with GDNF–RET
signaling (Fig. 6). Loss of Spry1, which is normally expressed in the mesonephric duct, results
in ectopic ureteric bud induction, multiple ureters,
multiplex kidneys, and hydroureter [222,
223]. Spry1 mutant embryonic kidneys have
increased expression of Gdnf and GDNF–RET

1

Embryonic Development of the Kidney

target genes and have increased sensitivity to
GDNF-induced ureteric induction in organ culture. Bmp4 is expressed in the tailbud-derived
mesenchyme (different than renal mesenchyme)
immediately next to the mesonephric duct and
ureteric bud [169, 224]. Mice heterozygous for
Bmp4 have ectopic or duplicated ureteric buds,
resulting in hypodysplastic kidneys, hydroureteronephrosis, and ureteral duplications [225,
226]. In vitro, Bmp4 has been shown to block
the ability of Gdnf to induce ureteric bud outgrowth from the mesonephric duct [85,
227]. BMP4 mutations have also been described
in humans with renal tract malformations [228].
The downstream effects of GDNF–RET signaling, namely, ureteric bud proliferation, survival, and ureteric outgrowth and branching, are
mediated by the transcription factors, Etv4 and
Etv5. Combined deletion of Etv4 and Etv5 causes
bilateral renal aplasia in mice [229]. Etv4 and Etv5
drive expression of several critical genes in the
ureteric bud tip, including Wnt11, Cxcr4, Mmp14,
Myb, and Met [229]. Furthermore, genetic deletion studies in mice have shown that Wnt11 is
necessary for normal Gdnf expression in the metanephric mesenchyme [230].

Renal Branching Morphogenesis
After growing into the metanephric mesenchyme,
the ureteric bud bifurcates into a T-shaped structure. The ureteric bud then continues to branch,
ultimately generating about 15 generations of
branches, with the earliest branches remodeling
to form the calyces and renal pelvis [231]. The
process of ureteric branching includes (1) expansion of the ureteric bud at its leading tip (termed
the ampulla), (2) division of the ampulla to form
new branches, and (3) elongation of newly formed
branches [232, 233]. In humans, during the first
9 generations of branching, ureteric bud tips
induce formation of new nephrons from the surrounding cap mesenchyme at about a 1:1 ratio
[2]. Ureteric bud branching is completed by the
20th–22nd week of human gestation, and subsequent collecting duct maturation occurs by elongation of peripheral (cortical) segments and

19

remodeling of central (medullary) segments
[2]. At this stage, four to seven new nephrons
are induced around each tip of a terminal
collecting duct branch [2, 43].
Localized cell proliferation contributes to initial ureteric bud outgrowth from the mesonephric
duct, formation and growth of ampullae, and elongation of ureteric branches [61, 108, 234,
235]. Cell survival is also critical for normal
renal branching morphogenesis; defects in cell
survival are associated with renal cystic dysplasia
and urinary tract dilatation. Moreover, targeted
deletion of bcl2 [236] and AP-2 [237], genes
critical for cell survival, results in increased apoptosis and collecting duct cysts in mice. In addition, experimental models of fetal and neonatal
urinary tract obstruction lead to apoptosis in
dilated collecting ducts [238, 239].
Several signaling pathways are necessary for
branching morphogenesis. In addition to its role in
ureteric bud induction, GDNF–RET signaling has
been shown to be critical for ureteric branching
in vivo and in vitro [213, 215]. As noted, Wnt11,
expressed in ureteric tips, is necessary for
maintaining normal Gdnf expression; conversely,
Wnt11 expression is reduced when Gdnf signaling
is absent. Furthermore, Wnt11 mutant mice
have defective ureteric branching morphogenesis
and
thus
develop
renal
hypoplasia
[230, 240]. Finally, conditional targeting of
β-catenin, a key mediator of canonical Wnt signaling, in the ureteric lineage results in aberrant
branching, loss of ureteric bud tip gene expression, and premature expression of differentiated
collecting duct genes [241, 242].
Fibroblast growth factor signaling is also critical for ureteric branching. In vitro studies have
shown that exogenous Fgf ligands differentially
modulate ureteric bud growth and proliferation
[243]. FGF10 preferentially stimulates proliferation at ureteric bud tips, whereas FGF7 increases
cell proliferation throughout the developing
collecting ducts [243]. In vivo, global deletion of
Fgf7 or Fgf10 in mice results in ureteric branching
defects and hypoplastic kidneys [58, 244]. Conditional targeting studies in mice have revealed that
Fgfr2 is likely the key Fgf receptor mediating
effects on ureteric branching [245–247]. Loss of

20

Fgfr2 in the ureteric bud results in hypoplastic
ureteric ampullae with reduced proliferation and
increased apoptosis, ultimately leading to a significant reduction in ureteric branching and hypoplastic kidneys. In addition, an interesting study
revealed that while combined loss of Sprouty1 and
Gdnf in mice largely rescues ureteric defects compared to when either locus is deleted alone, additional loss of Fgf10 in the combined mutants led
to complete loss of ureteric branching and renal
aplasia [248]; thus, FGF signals appear to be able
to largely substitute for GDNF in promoting ureteric morphogenesis in the absence of Sprouty1.
Finally, a combination of in vitro and in vivo
experiments revealed that Fgf signaling acts in a
coordinated fashion with Wnt11 and Gdnf to regulate ureteric morphogenesis, in concert with
Sprouty genes [249].

Patterning of the Medullary
and Cortical Collecting Ducts
From the 22nd–34th week of human gestation [2]
and embryonic day 15 birth in mice [43], the
cortical (peripheral) and medullary (central)
regions of the kidney become established. The
relatively compact, circumferential renal cortex
comprises approximately 70 % of the mature kidney volume [250]. The renal medulla develops
modified a cone shape and occupies the remainder
of the mature kidney volume [250]. The apex of
the medullary cone consists of collecting ducts
converging in the inner medulla and is termed
the papilla. Ultimately, medullary collecting
ducts become morphologically distinct from cortical collecting ducts. Medullary collecting ducts
become elongated and linear and remain relatively
unbranched in a region devoid of glomeruli. In
contrast, collecting ducts in the renal cortex
remain branched and induce nephrogenic cap
mesenchyme to form nephrons throughout
nephrogenesis.
These morphological differences are likely due
in part to distinct axes of growth in the developing
renal cortex and medulla. The renal cortex grows
circumferentially, which preserves the organization of the peripheral tissues, including

C. Bates et al.

differentiating glomeruli, nephron tubules, and
collecting ducts [250]. In contrast, the developing
renal medulla expands longitudinally, perpendicular to the axis of cortical growth, due to elongation of outer medullary collecting ducts
[250]. Stromal cells may be a source of stimulatory cues for the medullary growth [250]; studies
have shown that mice lacking the stromal transcription factors Foxd1 and Pod1 have abnormal
medullary collecting duct patterning [54, 66,
251]. Finally, apoptosis appears to participate in
remodeling the branched medullary ureteric tissues into elongated tubules, as programmed cell
death normally occurs prominently in developing
medullary ureteric epithelia that become the
papilla, calyces, and renal pelvis [108].
Multiple genes have been implicated in the
differentiation of cortical and medullary
collecting ducts, including those that encode for
soluble growth factors (Fgf7, Fgf10, Bmp4,
Bmp5, and Wnt7b), proteoglycans (Gpc3), cell
cycle regulatory proteins (p57KIP2), and components of the renin–angiotensin axis (angiotensin
and angiotensin type 1 and 2 receptors). Fgf7
mutant mice have marked papillary underdevelopment, while Fgf10 null kidneys exhibit medullary dysplasia with fewer loops of Henle and
medullary collecting ducts, increased medullary
stroma, and enlargement of the renal calyx [58,
244]. The ability of FGF ligands to bind properly
to their receptors requires interactions with cell
surface proteoglycans, including glypicans
[252]. Glypican-3 (GPC3) is required for normal
medullary patterning in humans and mice [253,
254]. Moreover, the medullary dysplasia observed
in Gpc3-deficient mice appears to result from
unrestrained proliferation and overgrowth of the
ureteric bud and collecting ducts, followed by
aberrant apoptosis [253, 254]. The Gpc3 / medullary defects appear to be driven by altered
responses of mutant collecting duct cells to
growth factors including Fgfs [254–256]. Finally,
mice lacking the cell cycle protein p57KIP2 demonstrate medullary dysplasia, with fewer inner
medullary collecting ducts [257]. Together, these
studies reveal the importance of balanced cell
proliferation and apoptosis in medullary
collecting duct patterning.

1

Embryonic Development of the Kidney

Proper elongation and growth of medullary
collecting ducts also appears to rely on oriented
cell divisions. Studies have shown that canonical
Wnt signaling in collecting ducts via Wnt7b leads
to proper oriented cell division and survival [258,
259]. Furthermore, α3β1 integrin and the receptor
tyrosine kinase c-Met act in concert to regulate
Wnt7b expression and signaling in medullary
collecting ducts [259].
Finally, angiotensin (Agt) and angiotensin
receptors (Agtrs) appear critical for the development of the renal calyces, pelvis, and ureter. Mice
lacking Agt or Agtr1 genes demonstrate progressive widening of the calyx and atrophy of the
papillae and underlying medulla [260,
261]. These defects appear to be caused by
decreased proliferation of the smooth muscle
cells that line the renal pelvis. Loss of Agtr2
causes a range of renal anomalies secondary to
ureteric mispatterning, including vesicoureteral
reflux, duplex kidney, renal ectopia, ureteropelvic
Fig. 7 H&E-stained
sections from E15.5 and
P1 mouse ureters and
bladders. E15.5 ureters and
bladders have early
urothelium (u), an inner
layer of mesenchyme
(future lamina propria,
arrows), and an outer layer
of condensing mesenchyme
(future muscle, m). P1
ureters and bladders have a
more stratified urothelium
(u) and well-developed
lamina propria (arrows) and
outer muscle (m) layers.
The adventitial layer of
fibroblasts that surrounds
the muscle in both tissues is
not labeled. Ureters =100
magnification; bladders
=40 magnification

21

or ureterovesical junction stenoses, renal dysplasia or hypoplasia, multicystic dysplastic kidney,
and renal aplasia [107].

Lower Urinary Tract Development
Anatomic and Functional Development
Concurrent with metanephric kidney formation,
the embryonic ureter and bladder develop the
former functioning to propel urine into the latter
which stores urine until an appropriate time to
expel it via the urethra. Similar to the kidney, the
ureter and bladder undergo maturation largely due
to reciprocal interactions between an epithelium
(i.e., urothelium) and surrounding mesenchyme
that forms the lamina propria, muscle, and adventitia (Fig. 7). For a recent detailed review on
anatomic and molecular control of lower urinary
tract development, see [262].

22

Ureter development begins simultaneously
with metanephric kidney development around
the 5th week of gestation in humans and at
E10.5 in the mouse, when the ureteric bud arises
from the mesonephric duct [262]. Thus the
embryonic origin of the ureteral urothelium is
the intermediate mesoderm, the same as the metanephric kidney. By E11.5 in the mouse, the ureteric bud has been segmented into a distal portion
that will develop into the ureter and a proximal
end that has invaded the metanephric mesenchyme to eventually branch and form the
collecting ducts and renal pelvis (see above).
The mesenchyme surrounding the early developing ureter consists largely of tailbud-derived mesenchyme that appears to be crucial for directing
the distal ureteric bud toward a ureter fate
[263]. Between E10.5 and E13.5, the nascent
ureter transitions from attaching to the nephric
duct to emptying directly into the early bladder
(see below). By E13.5, a thin outer ring of ureteral
mesenchyme condenses and expresses alpha
smooth muscle actin (αSMA) mRNA, the first
marker of differentiation toward a smooth muscle
fate [264]. αSMA protein expression is not noted
until E14.5 in the proximal portion of the ureter
(nearest the kidney) and then throughout the entire
length of ureter by E16.5; thus, ureter muscle
development progresses in a rostral to caudal
direction [265]. Concurrent with development of
the mesenchymal layers, the ureteral urothelium
gradually matures from a simple epithelium to a
stratified epithelium consisting of at least three
cell types: basal, intermediate, and superficial/
umbrella cells. Each of these cell types has distinct
structural features and molecular markers [266];
moreover, recent data strongly suggests that
urothelial basal cells, arising from the original
ureteric epithelium, serve as the progenitors for
other ureteral urothelial cell types [266]. A unique
feature of urothelium (compared with other epithelia) is the apical expression of urothelial
plaques, consisting of uroplakins, which likely
have several functions, including providing a permeability barrier and acting as a binding site for
uropathogenic E. coli [266].
An important function of the ureter is to continuously propel urine from the renal pelvis to the

C. Bates et al.

bladder. The ability of the ureter to undergo peristaltic waves of contraction followed by relaxation appears to be intrinsic and not dependent on
urine flow; cultured explants of E13.5 mouse ureters attached to kidneys begin to undergo spontaneous peristaltic contractions within a few days,
as do isolated and cultured E15.5 ureters
[268]. Elegant studies recently identified a population of pacemaker cells at the junction of the
renal pelvis and the kidney that express
hyperpolarization-activated cation-3 (Hcn3)
channels (a family of channels that are also present in cardiac pacemakers) [269]. Loss of Hcn3
activity in mice leads to abnormal coordination
and frequency of ureter contractions [269]. Following this study, another described a population
of secondary pacemakers that are located in the
muscle of the mouse proximal ureter (starting at
the ureteropelvic junction) and that have morphological and molecular features similar to intestinal
pacemakers including expression of c-kit
[268]. Thus, like the cardiac conduction system,
the ureter has primary and secondary pacemakers
that act to drive urinary propulsion in a coordinated fashion.
The embryonic bladder initially forms around
the 5th gestational week in humans and at
E11.5–12.5 in the mouse [262, 270]. Unlike the
ureter, bladder urothelium is derived from the
endodermal urogenital sinus, formed from the
ventral region of the cloaca. At E11.5–12.5, the
urogenital sinus further subdivides into an anterior portion, which will become the bladder, and a
posterior portion that forms the urethra and portions of the female vagina. The mesenchyme that
surrounds the bladder urothelium is largely
thought to be from splanchnic mesoderm,
although fate-mapping studies reveal that tailbud
mesenchyme also contributes to bladder mesenchyme (similar to the ureter) [263]. By E13.5, the
bladder is recognized as a distinct structure that is
attached directly to the ureters. As is the case with
ureters, αSMA mRNA expression in the developing bladder muscle precedes protein expression;
mRNA expression appears as early as E11.5 in
mice [264], while protein expression begins at
E13.5 [270]. Unlike the ureter (and many other
organs with smooth muscle such as the intestine)


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